Dear All This is all fun. Lost a bit of skin myself. Latex gloves does help a bit. Home made clamping devices I pressume will help, but I prefer to have a "hands on" onto the sample. For large amounts of samples automation is a option.
} Everett Ramer wrote: } } } } We have a manual metalographic grinding/polishing wheel. The sample } } } being prepared is held in the hand as it is pressed against the rotating } wheel. } } } Our safety people have asked us to provide finger protection for this } device. } } } Does anyone have any solution/suggestion? } } } Everett Ramer } } Everett, } Mary Mager's response, while funny, is actually quite accurate. There are } not too } many ways to protect fingers while properly holding a small (1-1/4", } typically) } sample for grinding/polishing. } } I preface my next remarks by pointing out that I work for a manufacturer of } } metallographic equipment and consumables, and therefore have a financial } interest in solving your problem: } } We at BUEHLER, do offer a simple grinding fixture which might help. This } fixture } is a squat, stainless steel, hollow cylinder with a carbide ring around } it's base. } The sample is clamped within another hollow cylinder seated within the } first. } The two cylinders are threaded, so that the inner can be raised or lowered } with respect to the carbide 'stop' of the outer. Engraved markings allow } material removal in increments as fine as 20microns. While this is not } actually } finger protection, per se, it will allow you to grasp something larger so } that your } fingers are not in such close proximity to the grinding wheel. We also } offer } a motor system which allows the fixture to rotate, in place, on the wheel} Mr. S H Coetzee Electron Microscope Unit Private bag X3 Wits Johannesburg 2050 Tell: +27 11 716 2419 Fax : +27 11 339 3407 E-mail stephan-at-gecko.biol.wits.ac.za
I am curious if anyone is using an automatic section stainer for TEM. If so, what brand are you using. We are using an LKB section stainer (15 years old now). I just wonder if there is anything else in the market. Thanks,
Cora Bucana ******************************************************* Corazon D. Bucana, Ph.D. Department of Cancer Biology U.T. M.D. Anderson Cancer Center 1515 Holcombe Blvd. Box 173 Houston, Texas 77030 Phone: (713) 792-8106 Email:bucana-at-audumla.mdacc.tmc.edu FAX: (713) 792-8747
May 20-22 and May 24-26, 1999 North Carolina State University Raleigh, North Carolina, USA
and
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This highly regarded hands-on course taught by expert faculty has been presented annually for more than 15 years. It deals with all phases of quantitative and computer-assisted imaging from acquisition and processing through measurement and stereological interpretation. Attendees receive The Image Processing Handbook plus a CD-ROM containing images, algorithms (Photoshop-compatible for Mac and Windows) and an extensive on-line tutorial and course notes on stereology and statistical analysis. The course is appropriate for scientists, technicians and administrators using or intending to use these techniques. Attendees typically come from materials science, geology, biological and medical sciences, pharmaceuticals, food science, industrial quality control, remote sensing, and other disciplines. You are encouraged to bring your own images for the hands-on lab sessions.
For detailed information and registration contact Cindy Allen, Dept. of Continuing and Professional Education, N. C. State University, Raleigh, NC 27695-7401, 919-515-8171, fax 919-515-7614, email: Cindy_Allen-at-NCSU.edu
Information is available on-line at the following sites:
Seadoohog-at-aol.com-at-sparc5.microscopy.com wrote: } } My next question concerns Energy Dispersive X-ray Spectroscopy (EDS). The } baseline on the EDS spectra is influenced by inelasic scattering of the } incident electron beam by the atomic nuclei of the sample, which results in a } peaked background. This is called the bremsstrahlung effect. What causes } this??? } } Thanks!
Dear Seadoohog, Brehmsstrahlung, or "braking radiation" is caused by the ac- celeration of the electron by a large mass (nucleus). Both Maxwell's equations and quantum mechanics predict that accelerated charges will give off electromagnetic radiation. The large mass is necessary so that conservation of both energy and momentum can be satisfied. Yours, Bill Tivol
There are some second hand equipment outlets here in the US who occasionally get bits and pieces for Zetopans. One is John Oren, in VT.
Re: illuminators - OptiQuip has suggested some interesting alternatives for an upgrade path. We will be working on this in Apr/May. Anyone interested is welcome to email privately for further info.
Best regards, 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 providing customized on-site workshops in all areas of microscopy, sample preparation, and image analysis.
At 01:41 PM 2/28/99 +0100, Yvan Lindekens wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
In the process of trying to produce some Ag coated samples, I instead obtained some silver beads, which would be dandy if that's what I was after ;^ { ...
The specifics were: Denton DV502A evaporator, 4cm of 0.2mm dia Ag wire wrapped around ~1mm dia section of a standard carbon rod, 5x10^-7 Torr. Melting of the wire occured abruptly at well under 20A.
Can anyone suggest conditions or parameters to that will yield evaporation rather than melting???
Later I expect to coat with Al, which like Ag melts and boils at much lower temperatures than Pd and Pt (which are no problem with the above...) so if anyone can provide similar information regarding Al, that would also be helpful.
(This is probably pushing my luck, but if anyone knows any rule of thumb for how thick the films of the above are as a function of conditions & time, that would be super to hear....)
Thanks.
_______________________________________________________ Get your free, private email at http://mail.excite.com/
O.k., I've lost my cheat sheet which had the sizes of Tobbaco Mosaic Virus components, and I can't locate the information presently. Can anyone help me out here? I know the particles are 18.0 nm in diameter "300 nm" in length (o.k. some of them break). What is the diameter of the central core?
What is the spacing between the sprialing sub units?
Thanks!
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 352 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu
Silver has a melting point of 961C. Its vapor pressure is 847C = 1e-8 torr 958C = 1e-6 torr 1105C = 1e-4 torr
An equilibrium vapor pressure of 1e-6 will give ~1 monolayer per second deposition rate (using a kinetic theory of gas model with a sticking coefiicient of 1). Since this will undoubtedly not be an equilibrium situation, you can this value to be an upper limit. So, at the melting point of silver, you will get much less than 0.2nm/sec deposition rate near the sample. Farther away, it will drop as 1/r^2.
Aluminum too, will melt long before you get much evaporation 660C = melting point 677C = 1e-8 torr 821C = 1e-6 torr 1010C = 1e-4 torr
Also, thermal evaporation of Ag, Au, Al will tend to produce metal islands on the sample which can interfere with high mag imaging. Sputtering often will give a smaller grain size.
Cheers, Henk
At 08:40 AM 3/1/99 -0800, you wrote: } {snip} } } Can anyone suggest conditions or parameters to that will yield evaporation } rather than melting??? } } Later I expect to coat with Al, which like Ag melts and boils at much lower } temperatures than Pd and Pt (which are no problem with the above...) so if } anyone can provide similar information regarding Al, that would also be } helpful. } } (This is probably pushing my luck, but if anyone knows any rule of thumb for } how thick the films of the above are as a function of conditions & time, } that would be super to hear....) } } Thanks.
Hendrik O. Colijn colijn.1-at-osu.edu Campus Electron Optics Facility Ohio State University (614) 292-0674 http://web.ceof.ohio-state.edu An optimist believes that we live in the best of all possible worlds. A pessimist fears that this is true.
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There was a request recently for information on a new EM techniques book. = The book is scheduled to be published in mid March so there is no = information yet other than the chapter titles. I post them here.
Electron microscopy methods and protocols / edited by M.A. Nasser Hajibagheri. (Methods in molecular biology ; v. 117) ISBN 0-89603-640-5 Expected publication date is mid March.
Contents: 1 General Preparation of Material and Staining of Sections ................= ... 1 Heather A. Davies 2 Negative Staining of Thinly Spread Biological Particulates ..............= . 13 J. Robin Harris 3 Preparation of Thin-Film Frozen-Hydrated/Vitrified Biological Specimens for Cryoelectron Microscopy .....................................= .. 31 J. Robin Harris and Marc Adrian 4 The Production of Cryosections Through Fixed and Cryoprotected Biological Material and Their Use in Immunocytochemistry .......... 49 Paul Webster 5 High-Pressure Freezing for Preservation of High Resolution Fine Structure and Antigenicity for Immunolabelling ............................= . 77 Kent McDonald 6 The Application of LR Gold Resin for Immunogold Labeling .............. = 99 J. R. Thorpe 7 Low-Temperature Embedding in Acrylic Resins .............................= . 111 Pierre Gounon 8 Quantitative Aspects of Immunogold Labeling in Embedded and Nonembedded Sections...................................................= ..... 125 Catherine Rabouille 9 Microwave Processing Techniques for Electron Microscopy: A Four-Hour Protocol ......................................................= ............. 145 Rick T. Giberson and Richard S. Demaree, Jr. 10 Electron Microscopic Enzyme Cytochemistry...............................= .... 159 Nobukazu Araki and Tanenori Hatae 11 In Situ Molecular Hybridization Techniques for Ultrathin Sections ....................................................= ................ 167 Jean-Guy Fournier and Fran=E7oise Escaig-Haye 12 Preparation of the Fission Yeast Schizosaccharomyces pombe for Ultrastructural and Immunocytochemical Study ..................... 183 M. A. Nasser Hajibagheri, Kenneth Sawin, Steve Gschmeissner, Ken Blight, and Carol Upton 13 Preparation of Double/Single-Stranded DNA and RNA Molecules for Electron Microscopy ...................................................= ............ 209 M. A. Nasser Hajibagheri 14 Applications of Electron Microscopy for Studying Protein-DNA Complexes..................................................................= .................. 229 Maria Schnos and Ross B. Inman 15 X-Ray Microanalysis Techniques .........................................= ............. 245 A. John Morgan, Carole Winters, and Stephen St=FCrzenbaum
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 nss4.cc.Lehigh.EDU (8.9.1a/8.9.1) with ESMTP id OAA158376; Mon, 1 Mar 1999 14:17:15 -0500 Message-ID: {36DAF5C8.6259645-at-lehigh.edu}
Please note the following job announcement:
ELECTRON MICROSCOPE TECHNICIAN
Lehigh University seeks an Electron Microscope Technician to perform technical duties in support of the Electron Microscopy Laboratory of the Materials Science and Engineering Department. Technician will instruct students in the operation of microscopes and other equipment; maintain and repair instruments; oversee upkeep of the lab; support research professors and students; analyze samples; give tours and demonstrations; supervise students; maintain a safe environment; and other assigned duties. Bachelors degree in physical science and/or 4+ years related work experience required. Candidates should be familiar with electron microscopes, mechanical and electronic equipment, vacuum systems, PC and/or MAC and EDS/WDS systems. Good communication and interpersonal skills are essential.
Lehigh University offers excellent benefits and vacation package. Interested candidates should forward resume to: Deanne Hoenscheid, Materials Science and Engineering, Lehigh University, 5 E. Packer Ave., Bethlehem, PA 18015. EOE. M/F/D/V.
On Mon, 1 Mar 1999 edelmare-at-casmail.muohio.edu-at-sparc5.microscopy.com wrote:
} Date: Mon, 1 Mar 1999 11:42:21 -0500 } From: edelmare-at-casmail.muohio.edu-at-sparc5.microscopy.com } To: microscopy-at-sparc5.microscopy.com } Subject: Size of TMV? } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } O.k., I've lost my cheat sheet which had the sizes of Tobbaco Mosaic Virus components, } and I can't locate the information presently. Can anyone help me out here? I know the } particles are 18.0 nm in diameter "300 nm" in length (o.k. some of them break). What } is the diameter of the central core? } } What is the spacing between the sprialing sub units? } } Thanks! } } } Richard E. Edelmann, Ph.D. } Electron Microscopy Facility Supervisor } 352 Pearson Hall } Miami University, Oxford, OH 45056 } Ph: 513.529.5712 Fax: 513.529.4243 } E-mail: edelmare-at-muohio.edu } } "RAM disk is NOT an installation procedure." }
TMV:
Pitch = 2.3 nm
Don't know about inner diameter of core; try: http://www.ncbi.nlm.nik.gov/ICTVdb/welcome.htm
Let me know the diameter if you find it. S
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've used W baskets to produce thin films of Au,Cr,Al and Cu in similar evaporators. I would think it would work with Ag too. The liquid metal seems to "hang" in the basket by what, I would guess, are surface tension effects. Any of the major EM houses should have these baskets.
I've also attempted to calculate, a priori, what the film thickness should be and never been too thrilled at the agreement with the result. A quartz crystal thickness monitor can be calibrated for most metals but if you don't have one, you might try using the change in resistance across a glass coverslip (Steere approach) or cross sectioning a series of thin films generated under known evaporation conditions and looking at them in a TEM.
cheers, John John Heckman Michigan State University
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
radsci-at-excite.com writes ... } } } In the process of trying to produce some Ag coated samples, I instead } obtained some silver beads, which would be dandy if that's } what I was after } ... } The specifics were: Denton DV502A evaporator, 4cm of 0.2mm } dia Ag wire wrapped around section of a standard carbon rod, } ...
I might suggest, rather than carbon rod, you wrap the Ag wire around tungsten wire which will be wetted. I imagine the Ag on carbon was like water on a duck's back.
BTW, why did a simple reply to this post want to send a message to {"radsci-at-excite.com"-at-sparc5.microscopy.com} ???
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/
Everett, When I have to do a quick grind on something small that I don't want to put into a mount, I use the "rubber fingers" that secretaries wear. I have found these to be much better than finger cots, which don't last one spin of the wheel. I don't know if this solution will satisfy the "safety" people but I still have finger prints. Dorrance
} ---------- } From: EVERETT RAMER } Sent: Thursday, February 1999 8:23 AM } To: Microscopy-at-Sparc5.Microscopy.Com } Subject: Finger Protection during Sample Prep } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } We have a manual metalographic grinding/polishing wheel. The sample } being prepared is held in the hand as it is pressed against the rotating } wheel. } Our safety people have asked us to provide finger protection for this } device. } Does anyone have any solution/suggestion? } Everett Ramer } }
We have been asked this question by a customer: =========================================== We are looking for a holder for the Wehnelt cap for filament adjustment. AMRAY had one when we went up there but they do not sell one nor could they tell us where to buy one. It looks like a custom job. Do you know what we are talking about? =========================================== I am sure that someone has this hidden away somewhere in their catalog but I have not been able to find it. Any help would be appreciated.
Chuck
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Look for us! ############################ WWW: www.2spi.com ############################ ==================================================
I run a very old probe in a small Geology department. I want to buy a JEOL 733 or later (eg 840A). I would be very pleased to hear from anyone who has one currently for sale or who is planning to sell one this year.
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
Dear all We do evaporation of various metals on a old Edwards evaporator. First the Tungstan (W) wire gets heated up under vacuum to clean it from all dirt. Then the requiered metal wire gets wraped around a V bent in the W wire. when heated up the wire will melt and form a melted globule. this will evaporate only after futher heating. }
} } The specifics were: Denton DV502A evaporator, 4cm of 0.2mm dia Ag wire } wrapped around ~1mm dia section of a standard carbon rod, 5x10^-7 Torr. } Melting of the wire occured abruptly at well under 20A. } } Can anyone suggest conditions or parameters to that will yield evaporation } rather than melting??? }
} } (This is probably pushing my luck, but if anyone knows any rule of thumb for } how thick the films of the above are as a function of conditions & time, } that would be super to hear....) } No it does exist.
p = density of the of the coating material in g. ( cm^-3) M = weight of coating material a source (g) R = source samle seperation, (cm) dx = film thickness.
(4 pie R^2) dx p = M
But 1) efficiency factor of evaporative method method is rarely better than 75%. thuss: (4 pie R^2) dx p = M.3/4 this will only work if the sample is directly under the evaporated source
If it is at a angle theta,
3/4.M = (4 pie R^2 dx p)/ sin theta Where theta is the angle between the gounr and the tip of the V.
since Em people do like a really thin coating (nm) for M in g/cm^3 R in cm and dx in nm
M = 4/3. ( 4pie R^2 dx p)/sin theta . 10^-7
For coating material in wire form for a diameter d cm and l lenth (cm0
M = (pie d^2 lp)/4
length of wire required for a coatint thickness of dx cm 1 = 64/3 .[( R^2 dx)/d^2 sin theta)]
hope this does help
Mr. S H Coetzee Electron Microscope Unit Private bag X3 Wits Johannesburg 2050 Tell: +27 11 716 2419 Fax : +27 11 339 3407 E-mail stephan-at-gecko.biol.wits.ac.za
Good day Listmembers A while ago there was a discussion on this list on the preparation of insect eggs for ultra-thin sectioning. Without re-opening the discussion, can somebody that collected all the replies and other experts in this regard please supply me with a general protocol if there is one?
TIA Alan N Hall Laboratory for Microscopy and Micro-Analysis NWII Building University of Pretoria Pretoria 0002 Republic of South Africa Tel: +27-12-420 3896(Office) +27-12-420 2075(Laboratory) Fax: +27-12-362 5150
Dear all, In the evaporation of metal from tungsten wire it is important not to raise the temperature of the wire too fast. If you do this, the metal will melt and drop off in a blob. The thing to do is to raise the temperature until the metal just starts to melt. Then wait until the wire is wetted - you'll see the liquid creeping up the sides of the V. The bulk of the metal will remain as a blob at the apex of the V but will not drop off. Then you can turn the heat up more until the evaporation takes place.
Eric
---------------------- Dr Eric E. Lachowski University of Aberdeen Department of Chemistry Meston Walk Old Aberdeen AB24 3UE Scotland +44 1224 272934 e.lachowski-at-abdn.ac.uk
At 03:54 PM 3/1/99, shAf wrote: } } BTW, why did a simple reply to this post want to send a } message to {"radsci-at-excite.com"-at-sparc5.microscopy.com} ??? } I am not sure about the why, but I see that too for various posts. I think it has something to do with the formating of the address, perhaps the use of the quotes.
You know, it could almost be beneficial. You can reply to the sender or the whole list by removing the unwanted part. However, about half the time, I forget to do the editing and my posts get kecked back for me to fix them.
Open Position: Microscopy Characterization Scientist
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Provide polymer morphology, particle and aesthetics characterization support at GE Plastics in Mt. Vernon, IN. In particular, TEM, SEM, optical microscopy, AFM, particle sizing, surface and other more specialized techniques are used. Leadership in effective problem solving approaches is a must.
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Interested candidates should send their resume to:
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The C.M.I./A.A.B. Descriptions of Plant Viruses No. 151 (Oct. 1975) on tobacco mosaic virus (type strain) gives the following information:
..cylindrical canal of radius c. 2 nm.....
As Sarah Miller states, a pitch of c. 2.3 nm is given.
More infromatiom about particle structure is given: If anyone would like a complete copy of this bulletin, I can mail you a copy. Dont' know if the fax machine will accept this document the way it is folded.
Maureen Petersen
************************************************************************ Maureen Petersen Department of Plant Pathology 1453 Fifield Hall University of Florida
My grandson in the states recently and proudly wrote to his Grand-Dad that he NOW has a microscope and could I send him some samples to look at. He told me that .....
"the microscope is EDU-SCIENCE and the magnifications are 100x 300x 600x."
Since my grandson is 10 years old and egar, I don't want to ask him questions like - Are the objectives aprochromats?. Is anyone on the list familiar with this microscope? I specifically want to know if it's optics are good, and if they are color corrected.
Once I know if he has a decent microscope or a toy that makes things biger and the user blind or crazy, I will better know how to encourage him, and what type of samples and reading matter to send.
Thanks for you help, from a land far away.
Shalom from Jerusalem, Azriel
+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+ Azriel Gorski, Head Fibers and Polymers Laboratory Division of Identification and Forensic Science Israel National Police Jerusalem
azrielg-at-cc.huji.ac.il ICQ User ID No. - 1750739
CHOICE - The enchanted blade, with an edge that shapes lifetimes. Richard Bach RUNNING FROM SAFETY
A friend is someone who knows the song in your heart, and can sing it back to you when you have forgotten the words. +-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+-+
Hi! I have a reference to two stains/methods for nerves: Koelle's acetyl cholinesterase technique and Winklemann's Silver Impregnation method. I have not been able to find either one. I found a "general" reference to acetyl cholinesterase from the University of Nottingham web page in which they recommend using 'DPK' mounting resin. I have never heard of it--can anyone help me out? Thanks!
Peggy Sherwood Wellman Labs of Photomedicine (224) Massachusetts General Hospital 50 Blossom Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-3192 (fax)
} -- [ From: Garber, Charles A. * EMC.Ver #3.1 ] -- } } We have been asked this question by a customer: } =========================================== } We are looking for a holder for the Wehnelt cap for filament adjustment. } AMRAY had one when we went up there but they do not sell one nor could they } tell us where to buy one. It looks like a custom job. Do you know what we } are talking about? } =========================================== } I am sure that someone has this hidden away somewhere in their catalog but I } have not been able to find it. Any help would be appreciated. } } Chuck
Chuck:
I believe these Wehnelt cap holders are available from Energy Beam Sciences.
Ken Bart
Kenneth M. Bart Director, Electron Microscopy Facility Hamilton College Clinton, New York 13323 USA kbart-at-hamilton.edu (315) 859-4715
} O.k., I've lost my cheat sheet which had the sizes of Tobbaco Mosaic Virus components, } and I can't locate the information presently. Can anyone help me out here? I know the } particles are 18.0 nm in diameter "300 nm" in length (o.k. some of them break). What } is the diameter of the central core?
The diameter of the central core is 40 A, it is packed with the nucleic acid (RNA). The protein subunits are arranged in a helix containing 16 1/3 subunits per turn ( or 49 subunits per three turns). Each subunit consists of 158 amino acids in a constant sequence.
} What is the spacing between the sprialing sub units? } It is the nucleic acid.
Best regards,
Ming
*********************************************** * 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 * ***********************************************
Charles writes ... } } We have been asked this question by a customer: } =========================================== } We are looking for a holder for the Wehnelt cap for filament } adjustment. } AMRAY had one when we went up there but they do not sell one } nor could they tell us where to buy one. ...
Just to clarify ... this sounds like a "jig" strictly for holding the wehnelt on the workbench(??) From personal experience I've never seen one of these as a necessary component for the adjustment ... or if it were, it came with the instrument, or was available from the manufacturer. Usually, its primary use is as a heat sink so that the wehnelt assy can be removed while it is hot and to allow it to cool down in a minimum amount of time. Since wehnelts vary in design and geometry I can't imagine this being available thru any other source other than the manufacturer ... which leaves you with what would seem to be a very simple job for a machine shop. (... why do I think this isn't much help? ...)
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/
} From: Mriglermas-at-aol.com Return-path: {Mriglermas-at-aol.com} To: Woody.N.White-at-mcdermott.com Cc: Mriglermas-at-aol.com
I wish to purchase a picoammeter for specimen current measurement (x-ray analysis work) and am requesting information on models, prices and vendors. Please contact me directly. Thanks.
#################################################################### 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 ####################################################################
Sonneborn, T.M. (1975), "The Paramecium aurelia complex of fourteen sibling species", Transactions of the American Microscopical Society 94: 155-178.
This is to support a high school student doing a very interesting project on individual variation in protozoa. The local library ordered a copy but estimates 3 weeks delivery.
Can someone out there help me to get it quickly? I will be happy to reimburse reasonable expenses.
Posted by our evaporater, dating from who-knows-when, but certainly long ago, is a sheet of recommended procedures for evaporation different metals. For silver it recommends using a molybdenum or tantalum wire as the filament. It says to wrap a fine wire of silver around the Ta or Mo filament, and then simply evaporate. I recently did a number of successful silver evaporations by fashioning (by hand) a Mo spiral basket and putting a small bead of Ag in it. (I had some Mo wire and Ag beads). I then evaporated the Ag just as I would have evaporated Au from a W basket. It worked fine, but, as always with evaporating from a basket in this way, the thickness uniformity over distances of 2-3 cm. was not very good.
My favorite form of fingertip protection has always (20+ years) been long fingernails. Not talons by any means, but longer than fingertip length. You can feel when a nail is touching without actually losing any skin. Granted, you sometimes generate a strange-looking manicure but an emery board or some 600 grit paper will soon put that to rights. One thing to watch out for: don't let the water to your wheel get so cold as to cause numbness in your fingers. If the water's too cold, you won't know you've ground off your fingerprints until you notice blood on the wheel. And it will really smart when the feeling comes back!
Stephan Coetzee wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Dear All } This is all fun. Lost a bit of skin myself. Latex gloves does help a } bit. Home made clamping devices I pressume will help, but I prefer } to have a "hands on" onto the sample. For large amounts of samples } automation is a option. } } } } Everett Ramer wrote: } } } } } } We have a manual metalographic grinding/polishing wheel. The sample } } } } being prepared is held in the hand as it is pressed against the rotating } } wheel. } } } } Our safety people have asked us to provide finger protection for this } } device. } } } } Does anyone have any solution/suggestion? } } } } Everett Ramer } } } } Everett, } } Mary Mager's response, while funny, is actually quite accurate. There are } } not too } } many ways to protect fingers while properly holding a small (1-1/4", } } typically) } } sample for grinding/polishing. } } } } I preface my next remarks by pointing out that I work for a manufacturer of } } } } metallographic equipment and consumables, and therefore have a financial } } interest in solving your problem: } } } } We at BUEHLER, do offer a simple grinding fixture which might help. This } } fixture } } is a squat, stainless steel, hollow cylinder with a carbide ring around } } it's base. } } The sample is clamped within another hollow cylinder seated within the } } first. } } The two cylinders are threaded, so that the inner can be raised or lowered } } with respect to the carbide 'stop' of the outer. Engraved markings allow } } material removal in increments as fine as 20microns. While this is not } } actually } } finger protection, per se, it will allow you to grasp something larger so } } that your } } fingers are not in such close proximity to the grinding wheel. We also } } offer } } a motor system which allows the fixture to rotate, in place, on the wheel} } Mr. S H Coetzee } Electron Microscope Unit } Private bag X3 } Wits } Johannesburg } 2050 } Tell: +27 11 716 2419 } Fax : +27 11 339 3407 } E-mail stephan-at-gecko.biol.wits.ac.za
Help- We are having a problem with getting nuclear autofluorescence of tissue culture cells fixed (Paraformaldehyde) and prepped for immunofluorescence. If you fix and mount the cells in glycerol-phenylene diamine with no antibody treatment, they usually look blank immediately, then over time they develop nuclear fluorescence. The nuclear stain is primarily in the fluorescein window and somewhat variable but seems to come up over time. Is it possible that this is coming from a breakdown product of the phenylene diamine we are using as an antifade? An apparently unrelated problem is nuclear background with secondary antibodies alone. If we treat with secondary and look immediately (before the other problem seems to arise), we also get nuclear staining. This seems to vary with the batch of secondary antibody so appears to be non-specific binding of the secondary. It is not blocked by serum etc. Any ideas appreciated. THanks- Dave
Dr. David Knecht Department of Molecular and Cell Biology University of Connecticut 75 N. Eagleville Rd. U-125 Storrs, CT 06269 Knecht-at-uconnvm.uconn.edu 860-486-2200 860-486-4331 (fax)
I am trying to dissolve Brefeldin A in Dulbecco's MEM for use in a cell = culture experiment for TEM study. However the drug does not seem to be = readily soluble in the MEM.=20 Can someone give me some help?
Dr. A.P. Alves de Matos Curry Cabral Hospital TEM Unit Portugal
{!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 am trying to dissolve Brefeldin A = in=20 Dulbecco's MEM for use in a cell culture experiment for TEM study. = However the=20 drug does not seem to be readily soluble in the MEM. {BR} Can someone = give me=20 some help? {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} Dr. A.P. Alves de Matos {BR} Curry = Cabral=20 Hospital {BR} TEM Unit {BR} Portugal {/FONT} {/DIV} {/BODY} {/HTML}
I have been watching what has been going on in relation to your sputter coating problems. I did not look too hard at the early postings so what = I have set out below may be covering old ground, but I hope it may help? M= ay I say that there have been many who have tried so hard to help you, my comments below in no way are critical of their stirling efforts.
Sputter coaters need a RP vacuum of just to the left of dead centre on yo= ur coater's vacuum gauge. It does not matter what the gauge has as its calibration; this setting should work for gold, gold-palladium or platinu= m.
There are three styles of sputter coater low voltage {600volts, medium voltage 1,000 to 1,500volts and a high voltage coater at 1,000 to 3,000volts.
The low voltage coaters do not have a voltage control, simply a depositio= n control which you should set at it's mid point. Set the voltage controls=
for the other two coaters in their mid range too.
As a test specimen use a 2" square piece of white paper weighted down wit= h a stub. Set the target to specimen distance at 5cms. Set the time at tw= o minutes (much too long for normal coating but fine for this test).
Pump the system down YOU DO NOT NEED ARGON! We often run with air, the difference is that argon gas gives a constant coating rate but air, wet o= r dry, gives different coatings. If you are using argon, flush to full lef= t scale once or twice to clean out the system during the pump down.
When you have a vacuum greater than 60% of the scale bleed the gas into t= he system until the pressure holds on the position mentioned above; just to the left of dead centre. When the vacuum is at the correct level, switch=
on the plasma and increase the voltage to the correct level. You should have a plasma and by adjustment of the voltage, deposition, or gas, adjus= t the current to ~20mA. Let the system run for the two minutes.
When the system completes its cycle open it up and take a look at the paper. Using a gold target - very light blue, its working but not very efficiently, pale blue, better but still not good enough, darker blue, still not good enough, blue-grey much more like it, grey to gold tint, it= s working fine.
If you do not have any of these colours the target material may be your problem. Is it one of the metals mentioned below? Even if the target is=
gold on brass and all the gold has gone you will still sputter brass! =
Chromium, aluminium and carbon will not sputter with the very simple systems used for conventional SEM. Much higher specification systems, i= n relation to current and vacuum, are required.
Well I do hope that helps? Please remember that sputter coaters are not 100% efficient when it comes to coating non-conducting specimens. Even a=
good coater will have trouble with rough and porous specimens. You shoul= d never expect to run above 15kV if you have coated this type of specimen.
Well that's it I do hope we can sort out your problem?
Regards
Steve Chapman
Senior Consultant E.M. Protrain, 16 Hedgerley, Chinnor, Oxford OX9 4TN, England. Tel & Fax 44 (0)1844 353161 Web Site - http://ourworld.compuserve.com/homepages/protrain For Consultancy and Courses in Electron Microscopy World Wide
If anyone is disposing of this Amray scope, I would appreciate knowing about it so that I could purchase some spare parts and assemblies from it before it is dumped. I am particularly interested in the main 30KV power supply and the CRT power supply, photomult assembly, and any of the circuit boards. (Specific model is 1600T without the digital controls and pushbuttons. All components and assemblies are of interest.)
I am also looking for a Robinson detector for this scope. Pls let me know if you have one and want to dispose of it.
I also have an EDAX 9100 with detector and LSI-11 console that I do not need. suggestions on its disposition are welcome.
Slightly off subject, but I am sure that someone has encountered this obstacle. We have ASCII data files on 8" floppies from various sort of data acquisision hardware/software and are seeking out a way to transfer the data to a useable media 3.5" would suffice, a 8" drive on a system w/internet access would be great, etc. etc.
Does anyone know of someone I could obtain assistance from?
__ _-==-=_,-. /--`' \_-at---at-.-- { Tim (TJ) LaFave Jr. `--'\ \ {___/. Department of Physics \ \\ " / University of North Carolina, Charlotte } =\\_/` { Charlotte, NC 28223 ____ /= | \_|/ _' `\ _/=== \___/ (704)547-3244 `___/ //\./=/~\====\ (704)509-6622 [Hm] \ // / | ===: http://www.iit.edu/~lafatim | ._/_,__|_ ==: __ \/ \\ \\`--| / \\ ---------- +*+ ---------- | _ \\: /==:-\ `.__' `-____/ |--|==: Such that the future be theirs \ \ ===\ :==:`-' to shape and direct. _} \ ===\ /==/ -----------------------------------------------------------------
Hi there there seems to be 2 versions of this book, one was pub in 1988 and one in 1995, what is the diff in them besides the ISBN number?
I have the older one I just got it today and it is a lot of fun was wondering what the newer one might offer..... The one I have has all black and white pictures in it. thanks Ed Sharpe
We are interested in quantifying the amount of sulphur in wool via EDXS analysis in the TEM.
Wool has approximately 5% sulphur within the fibre which varies for different components of the fibre. We would like to be able to mount the sample and standard in the same block and microtome them together. It is not necessary for the material to be fibrous but that would help!
To date we have tried a couple of methods without much success.
In the first instance we attempted to mix sulphur compounds in with the resin to obtain a homogenous material but found that no matter what we did there was always significant variability in the amount of sulphur throughout the resin.
The next attempt used a commercially available fibre which had a known amount of sulphur in it (0.25%) which we embedded in the resin and microtomed. This "sort of" worked OK but the level of sulphur was not quite high enough for our purpose and the uncertainties in our measurements were too high.
So, if anyone has any ideas or a source of materials we could use, I (and the rest of the team) would be most grateful.
Thanks very much in anticipation.
Colin Veitch
Instrumentation Scientist Fibre Structure & Function Group CSIRO Wool Technology PO Box 21 BELMONT Vic 3216 Australia
Tel: +61 (0)3 5246 4000 Fax: +61 (0)3 5246 4811
The information contained in this email message may be privileged or confidential information. If you are not an intended recipient, you may not copy, distribute or take any action in reliance on it. If you have received this message in error, please telephone CSIRO Wool Technology on +61 3 5246 4000.
If anyone is disposing of this Amray scope, I would appreciate knowing about it so that I could purchase some spare parts and assemblies from it before it is dumped. I am particularly interested in the main 30KV power supply and the CRT power supply, photomult assembly, and any of the circuit boards. (Specific model is 1600T without the digital controls and pushbuttons. All components and assemblies are of interest.)
I am also looking for a Robinson detector for this scope. Pls let me know if you have one and want to dispose of it.
I also have an EDAX 9100 with detector and LSI-11 console that I do not need. suggestions on its disposition are welcome.
I need information about SOPHIA process for aluminium castings ?
Best regards
Krzysztof Jan Huebner
{hubner-at-IOd.krakow.pl} :-)
FOUNDRY RESEARCH INSTITUTE Research Materials Department Manager of Structural and Mechanical Research Laboratory str. Zakopianska 73 Call (*48 12) 2618356 (after 8th march) 30-418 KRAKOW - POLAND Fax (+48 12) 2660870
Hi Lesley S. Bechtold, =20 I have read with interest your contribution from 1998-11-27 where you=20 described your treatment of mouse sperm prior to SEM. I want to=20 prepare thrombocytes in a similar way and wonder whether you always=20 apply 1% solutions of poly-L-lysine to render coverslips adhesive. As=20 I understand 0.01% solutions are suitable (and much less expensive...)= =20 for other applications. I deal with moderate amounts of "precious"=20 thrombocytes from knockout mice so I can not play around with=20 conditions. You certainly have extensive experience with this.=20 Any comments would be highly appreciated! =20 All the best, Matthias
I think DPK is a misprint for DPX which is a mounting medium consisting of polystyrene and plasticizers dissolved in xylene and should be widely available from most microscopy suppliers
Dan Hill Biochemistry Department Cambridge University Cambridge UK
On Tue, 2 Mar 1999, Peggy Sherwood wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Hi! } I have a reference to two stains/methods for nerves: Koelle's acetyl } cholinesterase technique and } Winklemann's Silver Impregnation method. I have not been able to find } either one. I found a "general" reference to acetyl cholinesterase from } the University of Nottingham web page in which they } recommend using 'DPK' mounting resin. I have never heard of it--can } anyone help me out? Thanks! } } Peggy Sherwood } Wellman Labs of Photomedicine (224) } Massachusetts General Hospital } 50 Blossom Street } Boston, MA 02114 } 617-724-4839 (voice mail) } 617-726-3192 (fax) } } } }
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There are 3-4 sulfur standards for the carbon black industry, ASTM D-1619. The concentrations I know about are 0.82%, 1.54%, and 1.93%. These are carbon black powders so combining them in block with your fibers would be a real challenge. I do know a source for these standards if you want to try them.
Chuck Butterick Engineered Carbons, Inc. ______________________________ Reply Separator _________________________________
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Hi All,
We are interested in quantifying the amount of sulphur in wool via EDXS analysis in the TEM.
Wool has approximately 5% sulphur within the fibre which varies for different components of the fibre. We would like to be able to mount the sample and standard in the same block and microtome them together. It is not necessary for the material to be fibrous but that would help!
To date we have tried a couple of methods without much success.
In the first instance we attempted to mix sulphur compounds in with the resin to obtain a homogenous material but found that no matter what we did there was always significant variability in the amount of sulphur throughout the resin.
The next attempt used a commercially available fibre which had a known amount of sulphur in it (0.25%) which we embedded in the resin and microtomed. This "sort of" worked OK but the level of sulphur was not quite high enough for our purpose and the uncertainties in our measurements were too high.
So, if anyone has any ideas or a source of materials we could use, I (and the rest of the team) would be most grateful.
Thanks very much in anticipation.
Colin Veitch
Instrumentation Scientist Fibre Structure & Function Group CSIRO Wool Technology PO Box 21 BELMONT Vic 3216 Australia
Tel: +61 (0)3 5246 4000 Fax: +61 (0)3 5246 4811
The information contained in this email message may be privileged or confidential information. If you are not an intended recipient, you may not copy, distribute or take any action in reliance on it. If you have received this message in error, please telephone CSIRO Wool Technology on +61 3 5246 4000.
by smtp.uky.edu (8.8.8/8.8.8) with ESMTP id JAA09170 for {Microscopy-at-MSA.Microscopy.Com} ; Wed, 3 Mar 1999 09:50:05 -0500 (EST) Received: from nc.gws.uky.edu ([128.163.193.169]) by pop.uky.edu (8.8.8/8.8.8) with SMTP id JAA13068 for {Microscopy-at-MSA.Microscopy.Com} ; Wed, 3 Mar 1999 09:50:05 -0500 (EST) Message-Id: {2.2.32.19990303145459.006bcec0-at-pop.uky.edu} X-Sender: jmcclin-at-pop.uky.edu X-Mailer: Windows Eudora Pro Version 2.2 (32) Mime-Version: 1.0 Content-Type: text/plain; charset="us-ascii"
Woody,
We have a JEOL 35CF for sale that was made in '82. Excellent condition and has backskatter detector. Dual supply Haskris water chiller also available that cooled a TEM as well as this SEM.
We have a stock of spare parts, vacuum tubes, manuals, tools, etc., for an old RCA EMU-3G microscope. Since the scope itself is long gone, these items are destined for that mysterious place where old scope parts go to die. UNLESS someone wants them.
Preserve history! Keep your EMU going! Read the manuals! Now's your chance. All yours for the mere cost of shipping.
Randy Tindall Electron Microscope Core College of Veterinary Medicine University of Missouri - Columbia Phone: 573-882-8304
I made a makeshift but useable holder for filament adjustment for a Amray 1830. I took one of the cardboard boxes that hold the coaters for Polaroid film (about 9"X2"X3/4") and glued two metal washers(1" outer diameter and ~7/16 inner diameter) on top of one another and to the middle of the box. The height of the washers was about 1/8". I then positioned the posts of the filament in the center of the washers to mark the spots. Then I used a needle to poke holes in the cardboard. The Wehnhelt can then be upright and it is easier to center the filament.
Mike Baxter Lehman College mykkb-at-juno.com
___________________________________________________________________ You don't need to buy Internet access to use free Internet e-mail. Get completely free e-mail from Juno at http://www.juno.com/getjuno.html or call Juno at (800) 654-JUNO [654-5866]
If this is what I think it is it should be available from E Fjeld C., Inc. 978-667-1416. CAA-FB Filament Alignment Base is item discription in my 1985 catalog.
I am posting this on behalf of nearby hospital colleagues:
1. JEOL 1200EX TEM for sale, serviced twice yearly since installation in 1985. In excellent condition. Very light use of up to 65 cases per year.
2. Reichert (now Leica) Ultracut S ultramicrotome, new in 1993, with table, drawers plus LKB Knifemaker 7800. Hardly used and in storage over the last three years.
For information, telephone Mike Sale on England (0)1872 255006. Cash offers to Keith Atkinson on England (0)1872 255006. An e-mail address: tracey.leigh-at-rcht.swest.nhs.uk
Please do not "Reply to sender" - although I would pass replies on to the correct contact!
Keith Ryan Marine Biological Association of the UK Plymouth, UK
I have been asked to research the latest methods for colloidal gold labelling in biological tissues. The researchers would first like to do some diaminobenzidine peroxidase staining on kidney tubules embedded in paraffin to see if the cell membrane antigen and distribution can be detected at the LM-peroxidase level.
Then the project would progress to preparation, sectioning, and labelling of kidney tissue at the TEM level. I have been asked to provide a beginning point of up-to-date protocols that are presently being done.
I am currently scanning Medline and Ovid search engines. Any suggestions would be greatly appreciated.
Sincerely,
Ginger Baker EM Lab Manager OSU-College of Osteopathic Medicine 918-561-8232 lizard-at-osu-com.okstate.edu
I just wanted to say a wholehearted thank you to all of you on this listserver. With your suggestions I have managed to inject an antibody which was directly conjugated to ultrasmall gold particles (thanks to Dr. Jan Leunissen and Electron Microscopy Sciences), perfuse the mouse, and embed the tissues with the specific orientation that I wanted in airtight flat embedding molds to polymerize under UV (thanks to so many people for different aspects of the embedding that I don't have space to name them all), then cut, silver enhance and examine by TEM!
The antibody in question had failed to label anything when the tissue was fixed with ANY fixative but these results were unquestionable and the morphology great!!
My most sincere thanks to you all - I'm off to try new things as our little research lab is closing down. Thanks and Good-bye!
Pat Hales McGill University Dept. of Anatomy & Cell Biology hales-at-med.mcgill.ca
hi, Brefeldin A does not dissolve in media , you need first to dissolve it in either Methanol or DMSO. We dissolve the Brefeldin A in Methanol at a conc of 5mgs/ml and use this as our stock to then be diluted into our buffer or media.
I am currently looking to upgrade my EDS system. I work in a semiconductor manufacturing facility(CMOS) and I currently have a Cambridge S200 SEM with Kevex Delta II.
I have an interest in Oxford, EDAX and a company called EVEX (which I am not familiar with)? Does anyone have experience with these companies - Good or Bad?
I am looking for a light element detector with possibly a WDS for Boron quantification.
Thanks in advance.
Jim Arnold Microelectronics and Technology Center AlliedSignal Electronics and Avionics Systems 9140 Old Annapolis Road Columbia, MD 21045
I have a requirement to process liquid and agar-bound pathogenic bacteria for SEM & TEM. I've been collecting them in cacodylate-buffered 3% glutaraldehyde where the final concentration has been 1.5% for the liquid suspended bacteria. These have been kept in the refrigerator. Does anyone have a protocol(s) that would lead me to SEM and TEM specimens from this point? Have I made a mistake already? My instruments are a Philips EM-400 and JEOL 6300V.
We have an immediate opening for a field service technician in the Los Angeles area. Responsibilities include the repair and maintenance of Scanning Electron Microscopes of a number of manufacturers including JEOL, Hitachi, Leo.
Candidate should have a minimum of three years electronic or instrumentation experience. Please e-mail or fax me at:
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Hello,
I have been asked to research the latest methods for colloidal gold labelling in biological tissues. The researchers would first like to do some diaminobenzidine peroxidase staining on kidney tubules embedded in paraffin to see if the cell membrane antigen and distribution can be detected at the LM-peroxidase level.
Then the project would progress to preparation, sectioning, and labelling of kidney tissue at the TEM level. I have been asked to provide a beginning point of up-to-date protocols that are presently being done.
I am currently scanning Medline and Ovid search engines. Any suggestions would be greatly appreciated.
Sincerely,
Ginger Baker EM Lab Manager OSU-College of Osteopathic Medicine 918-561-8232 lizard-at-osu-com.okstate.edu
We bought some of those chain meshed type gloves that protect fingers pretty well when hand grinding. Our metallographers say you can easily hold the samples and mounts. They look and feel rather clumsy to me tho. But if they work....alrighty then.
There are two special issues of journals on immunogold labeling which appeared recently. Cell Vision, Vol. 4, No. 6 (1997), and Microscopy Research and Technique, Vol. 42, No. 1 (1998). If you have trouble finding a hard copy of that issue of Cell Vision (I think it was from before it was indexed by Medline), you can download all the articles from the journal's web site - if you go to
this is the table of contents for this issue, and it links to the full text articles in pdf format.
Disclaimer: we supply immunogold reagents, and both these special issues contain several articles which feature results obtained with our products (Nanogold).
A good recent reference on post-embedding immunogold labeling is "Colloidal Gold Post-Embedding Immunocytochemistry" (Progress in Histochemistry and Cytochemistry, Vol 29, No 4) by Moise Bendayan (Gustav Fischer, 1995).
Hope this helps,
Rick Powell Nanoprobes, Incorporated.
} } Hello, } } I have been asked to research the latest methods for colloidal gold } labelling in biological tissues. The researchers would first like to do } some diaminobenzidine peroxidase staining on kidney tubules embedded in } paraffin to see if the cell membrane antigen and distribution can be } detected at the LM-peroxidase level. } } Then the project would progress to preparation, sectioning, and labelling } of kidney tissue at the TEM level. I have been asked to provide a } beginning point of up-to-date protocols that are presently being done. } } I am currently scanning Medline and Ovid search engines. Any suggestions } would be greatly appreciated. } } Sincerely, } } Ginger Baker } EM Lab Manager } OSU-College of Osteopathic Medicine } 918-561-8232 } lizard-at-osu-com.okstate.edu
****************************************************************** * NANOPROBES, Incorporated | US Toll-free: (877) 808-2101 * * 25 East Loop Road, Suite 113 | Tel: (516) 444-8815 * * Stony Brook, NY 11790-3350, | Fax: (516) 444-8816 * * USA | rpowell-at-mail.lihti.org * * * * NOW EASY TO FIND ON THE WEB: http://www.nanoprobes.com * ******************************************************************
Hello Jim., Since you mentioned the word evex, I suspect you missed the past discussions on this list server regarding them. It centered around their registering URLs with the trade names of all competitors so that any web search by manufacture's name produced the evex web page. Reports were that they were willing to sell the URLs to the principles for 10s of K$, perhaps more. Seems they (someone) did try a defense with an unsigned response. Personally wouldn't have confidence in any claim they made. You can probably find these discussion in the list server archives, ~a year ago.
Bruce Brinson Rice U.
Disclaimer: I use EDS & WDS on occasion. I have absolutely no financial interest & know no one with financial interest in the sales of equipment in this industry & no qualms about bringing this issue back to light.
Arnold, Jim 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 currently looking to upgrade my EDS system. I work in a semiconductor } manufacturing facility(CMOS) and I currently have a Cambridge S200 SEM with } Kevex Delta II. } } I have an interest in Oxford, EDAX and a company called EVEX (which I am not } familiar with)? Does anyone have experience with these companies - Good or } Bad? } } I am looking for a light element detector with possibly a WDS for Boron } quantification. } } Thanks in advance. } } Jim Arnold } Microelectronics and Technology Center } AlliedSignal Electronics and Avionics Systems } 9140 Old Annapolis Road } Columbia, MD 21045 } } email: Jim.arnold-at-alliedsignal.com } voice: (410) 964-4118 } fax: (410) 964-5046
I have to prepare pure Al foils for TEM investigation. What I need is (as usual :) a large thin area. What are the best electrolytes for the thinning of Al? Is there anything else to consider (e.g. temperature, ...)? As you see, I have not a lot of experience with this technique. Could anybody recommend a book concerning electrolytical thinning?
TIA,
Petra -------------------------------------------------------------- Dr. Petra Wahlbring Centre de Recherche Public Centre Universitaire (CRP-CU) Laboratoire d'Analyse des Materiaux (LAM) 162a, av. de la Faiencerie L-1511 Luxembourg tel. +352-466644-402 fax +352-466644-400 e-mail: petra.wahlbring-at-crpcu.lu Visit our WWW site! http://www.crpcu.lu/~wahlbrin
Don't forget our annual meeting in Gainesville Florida April 8-10
Dealines are fast approaching
For details see the latest issue of the Beam or our web site at :
http://www.biotech.ufl.edu/cbr/sems/sems.html
or cal or fax me below.
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
Hi, All We are looking for a Wenhelt Block Filament for filament ( SEM Philips 505 or 501), If you got one we're interested in it ( Free, or for sale , reasonable price)) , please contact us.
Thanks
***************************************************************** Mohamed Belhaj UFR SCIENCES Laboratoir d'Analyse des Solides Surfaces et Interfaces DTI/LASSI EP CNRS 120 BP 1039 Reims 51687 Cedex 2 Tel : 03 26 05 33 27 ou 03 26 05 33 14 Fax : 03 26 05 33 12 ******************************************************************
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Hi,
I have a usual question, which was posted before and, I believe, will be posted again and again. It is about printers for SEM images. So:
- What is the best printer for monochrome SEM/STEM images? - What is the best inexpensive printer for the same images? - What is the best printer for EDS output (color maps, maps+images)?
Thank you,
J.Meen
____________________________________________________________________ Get free e-mail and a permanent address at http://www.netaddress.com/?N=3D= 1
======================================================== FOCUS ON MICROSCOPY 1999
12th International Conference on 3D Image Processing in Microscopy 11th International Conference on Confocal Microscopy April 11th-15th, 1999 European Molecular Biology Laboratory (EMBL), Heidelberg, Germany
We wish to remind you about the upcoming deadline for abstract submission and modification.
The abstract submission and modification deadline is:
!!! Monday, 8 March 1999 --- 15.00 hours Heidelberg Time !!!
The "abstract submission part" of the database will definitely be closed at 15.00 hours, so that it is not possible to submit further abstracts or make any further modifications to the abstracts after this date.
The "registration only part" of the database will remain open until 9 April 1999!!!
Our website now also includes informations about some highlights of the commercial exhibition. This is an extract:
Leica Microsystems GmbH: Two-Photon Confocal System Leica TCS MP L.O.T. Oriel GmbH & Co KG: Nanopositioning Unit and Fiber Laser Molecular Probes: New fluorescent reagents for Cell Biology and Imaging highly - Fluorescent Alexa dyes Olympus Optical Co. GmbH: Confocal Laser-Scanning-Microscope with two-photon excitation Omicron Vakuumphysik GmbH: Scanning Near Field Optical Microscope (SNOM) Visitron Systems GmbH: 2D/3D Fluorescence Imaging System based on Cooled Digital Camera System Wallac Distribution GmbH: Confocal Microscope
12th International Conference on 3D Image Processing in Microscopy 11th International Conference on Confocal Microscopy April 11th-15th, 1999 European Molecular Biology Laboratory (EMBL), Heidelberg, Germany
Confocal microscopy, multiphoton excitation and deconvolution techniques are increasingly applied in the study of three-dimensional structures such as are encountered in biology, medicine and material sciences. Three-dimensional analysis and representation are crucial tasks in subsequent data assessment. These conferences offer a most efficient meeting point for developers and users working in these rapidly evolving fields and play an important role in the dissemination of information about new developments. Special attention will be given to the dramatic developments in live cell imaging and manipulation, such as the role of the green fluorescent protein. Further information:
Local Organizing Committee: Dr. Ernst H.K. Stelzer, EMBL, Heidelberg Prof. G.J. Brakenhoff, University of Amsterdam Dr. Andres Kriete, University of Giessen
Under the auspices of The International 3D Microscopy Society: Prof. Colin Sheppard, University of Sydney Dr. Andres Kriete, University of Giessen Prof. G.J. Brakenhoff, University of Amsterdam Prof. P-C. Cheng, SUNY at Buffalo Prof. Tony Wilson, University of Oxford Dr. Carol Cogswell, University of Sydney Dr. Vyvyan Howard, University of Liverpool Dr. Guy Cox, University of Sydney Dr. Ernst H.K. Stelzer, EMBL Prof. S. Kawata, Osaka University
I have never done any material sciences work - I've only ever done biological specimens for EM. Our engineering department has asked me to look at some ball bearings that are failing, as a favour. I know I don't need to fix or dehydrate but do I simply clean them, mount them (using double-sided tape?) and coat them as usual? Or is coating unnecessary? What whould I clean these with? I'm assuming there is grease somewhere that is not good for my vacuum!
Any help would be appreciated!! Thanks in advance.....
Lesley Bechtold
Lesley S. Bechtold Supervisor, Biological Imaging The Jackson Laboratory 600 Main St. Bar Harbor, ME 04609 207-288-6191
Ginger R Baker {lizard-at-osu-com.okstate.edu} CC: Bruce A Benjamin {benjamb-at-osu-com.okstate.edu} , Connie A Hebert {hconnie-at-osu-com.okstate.edu} , William D Meek {meekwd-at-osu-com.okstate.edu} X-Mailer: QuickMail Pro 1.5.3 (Mac) X-Priority: 3 MIME-Version: 1.0 Reply-To: Paul Webster {pwebster-at-mailhouse.hei.org} Content-Type: multipart/alternative; boundary="====50515355485450534849===1"
Reply to: RE: LM and TEM Need help locating the latest = Colloidal Gold T Dear Ginger,
The methods you are looking for can be slit into two sections; (i) = specimen preparation and (ii) colloidal gold labeling. For both, you have = many choices that will be determined by your lab equipment, lab skills = and budget. A good book to have next to you is "Fine structure = Immunocytochemistry" by G. Griffiths (published 1993 by Springer Verlag) = because it will give you advice on all your choices. First, specimen preparation: Your aim is to prepare samples that have = good morphology but also have good accessibility to antigen by the = antibodies and colloidal gold. The best way to achieve this is to obtain = sections. To section, you can either embed in resin (Lowicryl, LR White/= Gold, Unicryl, Monostep), or use cryosections. If you decide to use resin,= will you embed using the PLT method or by freeze substitution? Whatever = your choice, if you are to test the system first by light microscopy then = I would highly recommend using the same system for LM and EM. Do not try = to compare paraffin-embedded, HRP-DAB labeled material with aldehyde-fixed,= sectioned material. =
The gold labeling is also filled with many choices. It is clear that = sequential labeling (ie adding the primary antibody followed by a gold-= labeled secondary) is the best choice. However, what you choose as the = gold-labeled secondary may affect your labeleing pattern. If a = quantitative result is your aim (do you want a signal that reflects the = amount of antigen present?) then you must try not to use signal = amplification. For this either protein A-gold or immunoglobulin-gold as a = single secondary step is best. If you want to get a "strong" signal and = are not too worried about quantitative result you may want to try silver = enhancement of the gold marker.
If I were doing this study, I would fix the kidney, by perfusion, in 4% = formaldehyde (buffered in phosphate buffer). I would then cryoprotect = with sucrose, freeze in liquid nitrogen and embed part of it in Lowicryl = resin through freeze substitution. Other pieces of the cryoprotected, = frozen tissue, I would then section in a cryoultramicrotome. Both the = Lowicryl and cryo sections can be mounted onto glass substrate and labeled = with antibody and fluorescent secondary antibody. You can also label the = sections with the gold conjugated secondary you choose to use for EM and = visualize this for LM by silver enhancement. The advantage of using the = same reagents and tissues for LM is that you can easily test your system = before moving onto EM preparation. Once you have the conditions worked = out, the same blocks you used for LM can be sectioned again for EM (just = mount the sections on grids) and labeled with the same (or similar) = protocols you used for LM.
The protocols you need are all covered in the Griffiths book. Some of the = newer advances have been covered in the a book that had the contents = posted recently.
Contact me if you want more details, resources or references.
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 --====50515355485450534849===1 Content-Type: text/html; charset="US-Ascii" Content-Transfer-Encoding: quoted-printable
{HTML} {HEAD} {/HEAD} {BODY} {PRE WIDTH=3D"132"} Reply to: RE: LM and TEM Need help locating the latest = Colloidal Gold T
{/PRE} {FONT FACE=3D"Geneva" SIZE=3D3 = COLOR=3D"#000000"} Dear Ginger, {BR} {BR} The methods you = are looking for can be slit into two sections; = (i) specimen preparation and (ii) colloidal = gold labeling. For both, you have many = choices that will be determined by your = lab equipment, lab skills and budget. A = good book to have next to you is "Fine = structure Immunocytochemistry" by G. = Griffiths (published 1993 by Springer Verlag) = because it will give you advice on all your = choices. {BR} First, specimen preparation: = Your aim is to prepare samples that have = good morphology but also have good accessibility = to antigen by the antibodies and colloidal = gold. The best way to achieve this is to = obtain sections. To section, you can either = embed in resin (Lowicryl, LR White/Gold, = Unicryl, Monostep), or use cryosections. = If you decide to use resin, will you embed = using the PLT method or by freeze substitution? = Whatever your choice, if you are to test = the system first by light microscopy then = I would highly recommend using the same = system for LM and EM. Do not try to compare = paraffin-embedded, HRP-DAB labeled material = with aldehyde-fixed, sectioned material. = {BR} {BR} The gold labeling is also filled = with many choices. It is clear that sequential = labeling (ie adding the primary antibody = followed by a gold-labeled secondary) is = the best choice. However, what you choose = as the gold-labeled secondary may affect = your labeleing pattern. If a quantitative = result is your aim (do you want a signal = that reflects the amount of antigen present?) = then you must try not to use signal amplification. = For this either protein A-gold or immunoglobulin-gold = as a single secondary step is best. If = you want to get a "strong" signal = and are not too worried about quantitative = result you may want to try silver enhancement = of the gold marker. {BR} {BR} If I were doing = this study, I would fix the kidney, by perfusion, = in 4% formaldehyde (buffered in phosphate = buffer). I would then cryoprotect with = sucrose, freeze in liquid nitrogen and embed = part of it in Lowicryl resin through freeze = substitution. Other pieces of the cryoprotected, = frozen tissue, I would then section in a = cryoultramicrotome. Both the Lowicryl and = cryo sections can be mounted onto glass = substrate and labeled with antibody and fluorescent = secondary antibody. You can also label = the sections with the gold conjugated secondary = you choose to use for EM and visualize this = for LM by silver enhancement. The advantage = of using the same reagents and tissues for = LM is that you can easily test your system = before moving onto EM preparation. Once = you have the conditions worked out, the = same blocks you used for LM can be sectioned = again for EM (just mount the sections on = grids) and labeled with the same (or similar) = protocols you used for LM. {BR} {BR} The protocols = you need are all covered in the Griffiths = book. Some of the newer advances have been = covered in the a book that had the contents = posted recently. {BR} {BR} Contact me if you = want more details, resources or references. {/FONT} {FONT = FACE=3D"Monaco" SIZE=3D1 COLOR=3D"#000000"} {BR} {BR} Paul Webster, Ph.D {BR} House = Ear Institute {BR} 2100 West Third Street {BR} Los = Angeles, CA 90057 {BR} phone:213 273 8026 {BR} fax: = 213 413 6739 {BR} e-mail: pwebster-at-hei.org {BR} http://www.hei.org/htm/aemi.htm {/FONT} {/BODY} {/HTML} --====50515355485450534849===1--
Has anyone had any experience with these knives and would you like to comment on their performance relative to "standard" diamond knives?
Tom
Thomas Moninger moninger-at-emiris.iaf.uiowa.edu University of Iowa Central Microscopy Research Facility http://www.uiowa.edu/~cemrf Views expressed are mine.
On Wed, 3 Mar 1999 14:32:22 -0700, jwright-at-dugway-emh3.army.mil wrote... } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
For the TEM samples, fixation in small 1.5 mL eppendorf microfuge tubes works well since you can pellet the samples between solution changes if necessary. From the point you are at right now, you should rinse in your cacodylate-buffer (I used 0.1M cacodylate with 7.5% sucrose), then move on to post-fixation with 1% OsO4 in 0.1M cacodylate (no sucrose) until pellets turn dark brown or black (usually 30-min at room temp). If the pellets are really big, separate them before osmium treatment to make sure the entire pellet is fixed. Rinse in cacodylate buffer (no sucrose), dehydrate in an acetone or ethanol series. I use ethanol because I like to embed in LR White. After 2 changes in 100%, separate the SEM run (for critical point drying and coating) from the TEM samples- move on to resin infiltration right in the eppendorf tubes. I usually do 3:1 (solvent:resin) on a rotator for 1 hr., 1:1, 1:3, 100% times two and cure overnight. Good luck.
As promised, here is a summary of the replies I received to my enquiry about who uses the MSA (formerly EMSA) spectra file format.
Of eight replies, two didn't care for the format or never used it. "My personal preference is not to use MSA format for these purposes. It's way overcomplicated".
The rest of the replies were either reluctant or enthusiastic users who use the format to transfer spectra from one EDX analyser to another, or to the DTSA program, or into a desktop computer. Three would be just as happy to have any method to convert the spectra into Excel format, but three were quite enthusiastic about the ability to exchange spectra between different labs, different EDX programs on commercial systems, even different analysers from the same company of differnet versions.
Thanks to all who replied. I can send the text of all replies to anyone who wishes to see them.
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 best quality will probably always be from a dye sublimation printer. There are Kodak and Codonics machines at the high end for full page output at a cost of about $10K. Kodak also makes a small format (4x5) printer that does 3-color for about $600 and under $1/print. I don't know that they yet offer a black ribbon for true monochrome prints.
For general purpose work, most any of the inkjets will do decent work for both color and monochrome. You may just want to try some at your local computer stores to see what resolution you can get and how fast. We use an Epson 600 ( {$200) for pretty good work at maybe 2 minutes per full page print.
1200 dpi laser printers do a pretty good job for monochrome. We have a Lexmark Optra R. I have seen similar good results from an HP LaserJet 5. It takes a while to transmit the data to the printer, but then they whip out prints fast.
In all cases, the paper can do a lot to improve appearance. You will want some good stuff on had for the best prints.
None of what I have said is peculiar to EM work. You may want to enlist a few others in your area to get a good printer. Happy hunting.
Warren S.
At 09:13 AM 3/4/99 -0600, you wrote: } Hi, } } I have a usual question, which was posted before and, I believe, will be } posted again and again. It is about printers for SEM images. So: } } - What is the best printer for monochrome SEM/STEM images? } - What is the best inexpensive printer for the same images? } - What is the best printer for EDS output (color maps, maps+images)? } } Thank you, } } J.Meen
I am more of a TEM eprson, but I have a suggestion from watching my colleagues. Take an aluminium stub, which would normally go into an SEM, and using a 5 mm drill bit just drill a small , 1-2 mm deep, hole in top so that the ball bearing will sit in it. Before you mount the bearing, wash the bearing in organic solvents first, say two acetone washes and then two ethanol washes, and ultrasonically clean the bearing in an ultrasound bath with both solvents.
Dry it off by putting it into an oven at say 150 C for ten minutes and then mount it onto the stub using silver-dag mounting paint (used by almost everyone here) to fix the bearing onto the stub. If you have access to vacuum coating system with a Radio Frequency inductive plasma ring attachment or anything that can produce an Argon plasma, then put the stub in for ten minutes so that there are no organic residues left. Once it is finished you can put it straight into the SEM knowing that there should be a clean surface to look at. The silver dag paint should earth the bearing to the stub.
I hope this helps.
Jon
-- ***************************************** Jonathan Barnard
Microstructural Physics, H.H.Wills Physics Laboratory, University of Bristol, Tyndall Avenue, Bristol BS8 1TL.
Yes, that's right - if you are somewhere between Baltimore, Maryland and Sarasota, Florida (with a detour to visit my son in Chapel Hill, NC) I can deliver this lovely item to you in early April. I still don't want to crate it for shipment, but I am driving down to visit my mother and can drop it off on the way if it is not too far off my path. The item in question is an LKB-Huxley Ultra Microtome (model 9801-1) in nice clean condition with the knife holder and a set of chucks and collets. I have about $500 tied up in it which I would like to recover - if you can use it, get in touch and perhaps we can work something out. It just does not fit the work I am doing now, and I need both the $$ and space.
Dear Petra, We have had good success with Al alloys with the Struers Tenupol Twin-jet polisher, using their A-7 electrolyte at about -5 degrees C and 12V. The A-7 is: 100 ml. perchloric acid 700 ml. methanol 200 ml. gycerine The usual precautions with perchloric acid solutions apply. Another I have had recommended to me is: 100 ml. perchloric acid 900 ml. ethanol The solution must be kept at less than 10 degrees C. while polishing, and a voltage range of 7.5V to 20 V, depending on the alloy. I have no experience with pure Al, only commercial Al alloys. A book I recommend for these things is: "Metallography Principles and Practices" by Vander Voort, but I don't know if it is still in print.
You wrote: } } Hello Everyone, } } I have to prepare pure Al foils for TEM investigation. What I need is (as } usual :) a large thin area. What are the best electrolytes for the thinning } of Al? Is there anything else to consider (e.g. temperature, ...)? } As you see, I have not a lot of experience with this technique. Could } anybody recommend a book concerning electrolytical thinning? } } TIA, } } Petra } -------------------------------------------------------------- } Dr. Petra Wahlbring Best of luck, Mary
Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 fax: 604-822-3619 e-mail: mager-at-interchg.ubc.ca
We've been very pleased with the Epson Color Stylus 740 (approx. $280) and Epson Stylus Photo 700 (approx. $250) using Epson's Photo Quality Glossy Film. Image and text quality is very good to excellent.
James Martin Director of Analytical Services and Research Williamstown Art Conservation Center
Research Scientist, Chemistry Williams College
On Thu, 4 Mar 1999, Warren E Straszheim wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } The best quality will probably always be from a dye sublimation printer. } There are Kodak and Codonics machines at the high end for full page output } at a cost of about $10K. Kodak also makes a small format (4x5) printer that } does 3-color for about $600 and under $1/print. I don't know that they yet } offer a black ribbon for true monochrome prints. } } For general purpose work, most any of the inkjets will do decent work for } both color and monochrome. You may just want to try some at your local } computer stores to see what resolution you can get and how fast. We use an } Epson 600 ( {$200) for pretty good work at maybe 2 minutes per full page print. } } 1200 dpi laser printers do a pretty good job for monochrome. We have a } Lexmark Optra R. I have seen similar good results from an HP LaserJet 5. It } takes a while to transmit the data to the printer, but then they whip out } prints fast. } } In all cases, the paper can do a lot to improve appearance. You will want } some good stuff on had for the best prints. } } None of what I have said is peculiar to EM work. You may want to enlist a } few others in your area to get a good printer. Happy hunting. } } Warren S. } } At 09:13 AM 3/4/99 -0600, you wrote: } } Hi, } } } } I have a usual question, which was posted before and, I believe, will be } } posted again and again. It is about printers for SEM images. So: } } } } - What is the best printer for monochrome SEM/STEM images? } } - What is the best inexpensive printer for the same images? } } - What is the best printer for EDS output (color maps, maps+images)? } } } } Thank you, } } } } J.Meen } } }
Depending on the suspected failure mechanism, just give them a good soaking in acetone or some kind of degreasing detergent (local auto store) followed by a good rinse and dry. That should remove any grease. I would however talk to the Eng. folks to make sure that your cleaning would not disturb the failure mecahnism or worse.... enhance it. Yes, I would give them a light coat of Au or AuPd. Mounting on any kind of tape (carbon or otherwise) might present some drifting problems in the SEM.......perhaps Ag or C paint would do the trick....that's our solution around here anyway. Hope this helps.
John Staman LSI Logic, Process Analysis Lab, Colorado Springs, CO. 719-573-3282
} -----Original Message----- } From: Lesley S. Bechtold [SMTP:lsb-at-aretha.jax.org] } Sent: Thursday, March 04, 1999 8:53 AM } To: microscopy-at-sparc5.microscopy.com } Subject: Biologist needs help from Material Scientists!! } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Hi, } } I have never done any material sciences work - I've only ever done } biological specimens for EM. Our engineering department has asked me to } look at some ball bearings that are failing, as a favour. I know I don't } need to fix or dehydrate but do I simply clean them, mount them (using } double-sided tape?) and coat them as usual? Or is coating unnecessary? } What whould I clean these with? I'm assuming there is grease somewhere } that is not good for my vacuum! } } Any help would be appreciated!! Thanks in advance..... } } Lesley Bechtold } } } Lesley S. Bechtold } Supervisor, Biological Imaging } The Jackson Laboratory } 600 Main St. } Bar Harbor, ME 04609 } 207-288-6191 }
Assuming that you want to look at the bearings in an SEM and that the balls are made of steel, the solution is very simple:
Clean the balls to get rid of any grease. A few years back we used Trichlorethylene to degrease materials, but that is environmentally unsafe. There are other possibilities for safer degreasing.
Then you could just use a simple specimen holder and glue the balls to the holder with silver paint or carbon paint, mainly to fix them in place. I am not sure what you mean by double-sided sticky tape, but I would not use that. Who knows what the tape might do when the chamber is pumped down. Since they are steel, you don't have to worry about conductivity, just put them in the chamber, pump down and enjoy.
Michael Bode
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: info-at-soft-imaging.com
} ---------- } From: Lesley S. Bechtold[SMTP:lsb-at-aretha.jax.org] } Sent: Thursday, March 04, 1999 8:53 AM } To: microscopy-at-sparc5.microscopy.com } Subject: Biologist needs help from Material Scientists!! } } ---------------------------------------------------------------------- } -- } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------- } -. } } } Hi, } } I have never done any material sciences work - I've only ever } done } biological specimens for EM. Our engineering department has asked me } to } look at some ball bearings that are failing, as a favour. I know I } don't } need to fix or dehydrate but do I simply clean them, mount them (using } double-sided tape?) and coat them as usual? Or is coating } unnecessary? } What whould I clean these with? I'm assuming there is grease } somewhere } that is not good for my vacuum! } } Any help would be appreciated!! Thanks in advance..... } } Lesley Bechtold } } } Lesley S. Bechtold } Supervisor, Biological Imaging } The Jackson Laboratory } 600 Main St. } Bar Harbor, ME 04609 } 207-288-6191 } }
I need a razor blade holder to use in an old A/O CryoCut vintage 1978. It's a weird one, the blade is mounted to the far right of the holder to accommodate the knife mounting mechanism of the cryostat.
I could also use a razor blade holder for use in an old A/O rotary microtome. This is pretty standard, with the blade in the center of the holder.
I think they are available new, but are pretty expensive given the scope of my project. Anything old but serviceable would work for me. If you have something, maybe we can workout a deal.
Thanks
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
I have a few technical papers that deal with electropolishing aluminum using our Model 550 Single Vertical Jet Electropolisher which may be of interest:
1)" Characterization of the Microstructure, Fracture, Morphology and Toughness in Particulate and Short Fibre Reinforced Aluminum Matrix Composites", M.R. Krishnadev et al
2) "Preparation of Iron and Aluminum Samples for Ion Implantation and TEM=
Examination" J.M. McDonald
and the "Bible" of Jet Polishing:
3) "Polishing Methods for Metallic and Ceramic Transmission Electron Microscopy Specimens", B.J. Kestel
Kestel report is 60+ pages and give preparation guidelines for dozens of materials. He has recipes for pure aluminum as well as several aluminum alloys. I would recommend this report for anyone doing electropolishing.=
I have copies of all 3 of these reports as well as a bibliography of over=
200 technical reports mostly dealing with specimen preparation. I would = be pleased to mail you a copy of any or all of these if you think they will = be of use. Please let me know.
DISCLAIMER: South Bay Technology does manufacture the Model 550 Single Vertical Jet ELectropolisher and has a vested interest in promoting its use.
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy. Message text written by Petra Wahlbring } ------------------------------------------------------------------------=
The Microscopy ListServer -- Sponsor: The Microscopy Society of America =
Hello Everyone,
I have to prepare pure Al foils for TEM investigation. What I need is (as=
usual :) a large thin area. What are the best electrolytes for the thinni= ng of Al? Is there anything else to consider (e.g. temperature, ...)? As you see, I have not a lot of experience with this technique. Could anybody recommend a book concerning electrolytical thinning?
J. Meen asks ... } } } ... } } - What is the best printer for monochrome SEM/STEM images?
A dye-sublimation printer with its optional gray scale media.
} - What is the best inexpensive printer for the same images?
A good 600dpi laser printer configured for random dithering, although the next printer would work well too. I would opt for $$ being spent up front on the printer ... it will still cost pennies per page.
} - What is the best printer for EDS output (color maps, maps+images)?
A photographic quality ink jet would work well here ... inexpensive and could as well serve your second purpose, 'cept for speed.
If you're concerned with archiving, fading can be a problem with color ink jets ... moreso than with dye-subs ... so the dye-sub can serve dual purposes, gray scale and color. But, it is a hassle to constantly be changing its purposes from monochrome to color. The dye-sub's color media can be used to print gray scale, but it almost always has a tint because CMY is used, therefore I would not classify a color mode dye-sub as the best printer for your first need.
my $0.02 and 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/
Lesley writes ... } } } ... Our engineering department has asked me to } look at some ball bearings that are failing, as a favour. I } know I don't } need to fix or dehydrate but do I simply clean them, mount them (using } double-sided tape?) and coat them as usual? Or is coating } unnecessary? } What whould I clean these with? I'm assuming there is grease } somewhere } that is not good for my vacuum! } } ...
I would clean them first with acetone and then with clean ETOH and then dry to remove the small amount of absorbed water. They shouldn't need coating, but I'd mount them with conductive double-stick tape. Should be as easy as that :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/
I am using a Drukker Diamond Knive at 3 years. It is a excellent knive. I did perfect sections without compression. I cut plant tissues and pollen grains with a hard cell wall.
Rinaldo Pires dos Santos Dept of Botany - Plant Anatomy Laboratory - UFRGS Porto Alegre - RS - Brazil
A student of ours is going to the Washington, D.C. area this summer. I've already told her to stay away from the White House. What she really wants to know is: are any EM/Imaging facilities in the area? She would like to use one to help her with her work while she's there. So drop me a line and I'll pass the info on to her.
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
Warren E Straszheim says: } The best quality will probably always be from a dye sublimation printer. } There are Kodak and Codonics machines at the high end for full page output } at a cost of about $10K. Kodak also makes a small format (4x5) printer that } does 3-color for about $600 and under $1/print. I don't know that they yet } offer a black ribbon for true monochrome prints.
1. Several other manufacturers (JVC, Sony, Panasonic) offer dye sublimation printers, marketed for catching/printing video frames. [I'm planning to buy one of those, probably Sony.]
2. I doubt that a term "ribbon" can be applied to such a printer...
} For general purpose work, most any of the inkjets will do decent work for } both color and monochrome. You may just want to try some at your local } computer stores to see what resolution you can get and how fast. We use an } Epson 600 ( {$200) for pretty good work at maybe 2 minutes per full page } print.
Interesting. For monochrome of course inkjets hardly cut it. But for color - it suddenly becomes cost-effective and attractive.
} 1200 dpi laser printers do a pretty good job for monochrome. We have a } Lexmark Optra R. I have seen similar good results from an HP LaserJet 5. It } takes a while to transmit the data to the printer, but then they whip out } prints fast.
1. I have yet to see a laser printer (within $2K price range) that comes close to Optra in reproducing photographs. LaserJet-5 is good, but not that good.
2. I wonder what kind of printer connection you use, and how large your files are. My printers are on Ethernet, and 16MB files "fly in" in less than half-a-minute.
} In all cases, the paper can do a lot to improve appearance. You will want } some good stuff on head for the best prints.
(:-) -- Regards, Uri uri-at-watson.ibm.com -=-=-=-=-=-=- {Disclaimer}
Additional to the useful replies relating to the preparation of ball bearings, I would like to add a little regarding the microscopy: Small, clean and undamaged ball bearings are a good instructive aid to explain SEM functioning. This may be of some use to Jonathan when viewing those bearings and to anybody trying to explain some SEM effects. 1 At high kV (say above 20) you may have trouble seeing anything, because fine surface structure and dirt will be invisible. 2 At low kV (say 10 and below) such surface structures will be visible. 3 At low powers, regardless of kV, the ball bearing will appear like a disk, but the outer part of the disk is brighter. This nicely shows that in secondary mode, brightness almost entirely is increased with the angle of incidence. Being a sphere, tilt has no effect on the brightness distribution over that image. 4 A BS detector mounted at an angle (whereas the Robinson and some others are vertical) will make the distinction that the specimen is not a disk but a sphere, because the BS electrons directed away from the detector leave a shadow. 5 A similar effect is produced when the bias current of the secondary detector is turned off and the condenser current is turned down. This floods the secondary detector's scintillator with backscattered electrons and produces a BS image quite suitable for low powers.
Nice teaching exercise, but its useful to know about these effects when actually looking at those bearings. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes ********************** www.proscitech.com.au *****
On Friday, March 05, 1999 6:11 AM, Jonathan Barnard [SMTP:J.Barnard-at-bristol.ac.uk] wrote: } } I am more of a TEM eprson, but I have a suggestion from } watching my } colleagues. } Take an aluminium stub, which would normally go into an } SEM, and using a } 5 mm drill bit just drill a small , 1-2 mm deep, hole in } top so that the } ball bearing will sit in it. Before you mount the bearing, } wash the } bearing in organic solvents first, say two acetone washes } and then two } ethanol washes, and ultrasonically clean the bearing in an } ultrasound } bath with both solvents. } } Dry it off by putting it into an oven at say 150 C for ten } minutes and } then mount it onto the stub using silver-dag mounting } paint (used by } almost everyone here) to fix the bearing onto the stub. If } you have } access to vacuum coating system with a Radio Frequency } inductive plasma } ring attachment or anything that can produce an Argon } plasma, then put } the stub in for ten minutes so that there are no organic } residues left. } Once it is finished you can put it straight into the SEM } knowing that } there should be a clean surface to look at. The silver dag } paint should } earth the bearing to the stub. } } I hope this helps. } } Jon } } -- } ***************************************** } Jonathan Barnard } } Microstructural Physics, } H.H.Wills Physics Laboratory, } University of Bristol, } Tyndall Avenue, } Bristol BS8 1TL. } } 0117 928 9000 ext 8750 } } ***************************************** } }
The string concerning preparation of bacteria touched on delayed second fixation. This is worth a separate discussion on delayed second (usually Os) fixation: Sabatini, first to publish GA as a fixative, also published that Os fixation could be delayed by several months. That seems true for some tissues, which show no ill-effects when compared with the usual, immediate double fixation. However, we found in the lab that other tissues are sensitive to that delay. I used to run a couple of busy service labs and cannot remember specifically which tissues and what structures were affected. It would be interesting to know when delayed double fixation is acceptable and others may have experience to share. I believe that specimen preparation for SEM is never affected by delayed second fixation. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes ********************** www.proscitech.com.au *****
On Thursday, March 04, 1999 11:14 PM, Tina Schwach [SMTP:tschwach-at-tc.umn.edu] wrote: } } On Wed, 3 Mar 1999 14:32:22 -0700, } jwright-at-dugway-emh3.army.mil wrote... } } I have a requirement to process liquid and agar-bound } } pathogenic } } bacteria for SEM & TEM. I've been collecting them in } } cacodylate-buffered 3% glutaraldehyde where the final } } concentration has } } been 1.5% for the liquid suspended bacteria. These have } } been kept in } } the refrigerator. Does anyone have a protocol(s) that } } would lead me to } } SEM and TEM specimens from this point? Have I made a } } mistake already? My } } instruments are a Philips EM-400 and JEOL 6300V. } } } } John Wright } } Microbiologist } } } } West Desert Test Center } } Dugway, UT } } } } } } John, } I have stored samples in primary fixative (glut-para- } ruthenium red in } cacodylate) for several weeks, even months and they appear } to be fine. } For SEM, you may want to place (dry) your samples on some } kind of surface, } ie stainless steel chips, so you'll be able to view them. } You'll have to } do this at the end anyway. You can even place them on } nucleopore filter } membranes. Depending on what you want to see, the agar } strands can get in } the way. } } For the TEM samples, fixation in small 1.5 mL eppendorf } microfuge tubes } works well since you can pellet the samples between } solution changes if } necessary. From the point you are at right now, you } should rinse in your } cacodylate-buffer (I used 0.1M cacodylate with 7.5% } sucrose), then move on } to post-fixation with 1% OsO4 in 0.1M cacodylate (no } sucrose) until pellets } turn dark brown or black (usually 30-min at room temp). } If the pellets are } really big, separate them before osmium treatment to make } sure the entire } pellet is fixed. Rinse in cacodylate buffer (no sucrose), } dehydrate in an } acetone or ethanol series. I use ethanol because I like } to embed in LR } White. After 2 changes in 100%, separate the SEM run (for } critical point } drying and coating) from the TEM samples- move on to resin } infiltration } right in the eppendorf tubes. I usually do 3:1 } (solvent:resin) on a } rotator for 1 hr., 1:1, 1:3, 100% times two and cure } overnight. } Good luck. } } } } }
The peaks have suddenly started to move around a bit, although the resolution stays good. The problem comes and goes, in its good times the standard deviation of the position of the Co Ka peak is about 0.4 eV (10 determinations), but sometimes it's about 10 or even 20 eV. My thinking is that if it were the detector, the resolution would be degrading, but it's not, so maybe the culprit is the (analog) pulse-processor. Anyone got any thoughts on how to pin it down as being either a the detector b the subsequent signal-processing stuff, eg pulse-proc? Could I successfully test the pulse-proc with a ramp from a standard signal generator, or would that signal, being relatively clean compared with that from a detector, not really check it out rigorously enough?
cheers (well, I try)
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
======================================================== FOCUS ON MICROSCOPY 1999
12th International Conference on 3D Image Processing in Microscopy 11th International Conference on Confocal Microscopy April 11th-15th, 1999 European Molecular Biology Laboratory (EMBL), Heidelberg, Germany
We wish to remind you about the upcoming deadline for abstract submission and modification.
The abstract submission and modification deadline is:
!!! Monday, 8 March 1999 --- 15.00 hours Heidelberg Time !!!
The "abstract submission part" of the database will definitely be closed at 15.00 hours, so that it is not possible to submit further abstracts or make any further modifications to the abstracts after this date.
The "registration only part" of the database will remain open until 9 April 1999!!!
Our website now also includes informations about some highlights of the commercial exhibition. This is an extract:
Leica Microsystems GmbH: Two-Photon Confocal System Leica TCS MP L.O.T. Oriel GmbH & Co KG: Nanopositioning Unit and Fiber Laser Molecular Probes: New fluorescent reagents for Cell Biology and Imaging highly - Fluorescent Alexa dyes Olympus Optical Co. GmbH: Confocal Laser-Scanning-Microscope with two-photon excitation Omicron Vakuumphysik GmbH: Scanning Near Field Optical Microscope (SNOM) Visitron Systems GmbH: 2D/3D Fluorescence Imaging System based on Cooled Digital Camera System Wallac Distribution GmbH: Confocal Microscope
12th International Conference on 3D Image Processing in Microscopy 11th International Conference on Confocal Microscopy April 11th-15th, 1999 European Molecular Biology Laboratory (EMBL), Heidelberg, Germany
Confocal microscopy, multiphoton excitation and deconvolution techniques are increasingly applied in the study of three-dimensional structures such as are encountered in biology, medicine and material sciences. Three-dimensional analysis and representation are crucial tasks in subsequent data assessment. These conferences offer a most efficient meeting point for developers and users working in these rapidly evolving fields and play an important role in the dissemination of information about new developments. Special attention will be given to the dramatic developments in live cell imaging and manipulation, such as the role of the green fluorescent protein. Further information:
Local Organizing Committee: Dr. Ernst H.K. Stelzer, EMBL, Heidelberg Prof. G.J. Brakenhoff, University of Amsterdam Dr. Andres Kriete, University of Giessen
Under the auspices of The International 3D Microscopy Society: Prof. Colin Sheppard, University of Sydney Dr. Andres Kriete, University of Giessen Prof. G.J. Brakenhoff, University of Amsterdam Prof. P-C. Cheng, SUNY at Buffalo Prof. Tony Wilson, University of Oxford Dr. Carol Cogswell, University of Sydney Dr. Vyvyan Howard, University of Liverpool Dr. Guy Cox, University of Sydney Dr. Ernst H.K. Stelzer, EMBL Prof. S. Kawata, Osaka University
Others have offered their suggestions but may I add a friendly warning, o= r offer a bit of fun?
Ball bearings are a great eye opener for the novice SEM operator. Fix a small ball bearing to a stub with a NON conducting adhesive. Run the microscope at 20 to 25kV for a few minutes and the ball will charge like=
mad. let it charge { one of the very few reasons for using such high kV := -) }. NOW drop to 2kV and wonder of wonders you will see the inside of the=
specimen chamber. Move the ball around and change the magnification and you will be able to view the chamber in detail. Take a look at the aperture of the final lens, the EDX detector face, or the backscattered detector surface. Great fun if you know what is happening, totally confusing if you did not do it deliberately.
This "problem" may occur in other circumstances where, after using high k= V and charging a specimen, moving to a low kV the charge does not go away b= ut simply acts as a reflector of the beam. Double sided tape which is non conducting is the biggest culprit. It should never be used in SEM where = we now have available carbon double sided media which is great for a quick f= ix with lighter weight specimens.
Have fun?
Steve Chapman
Senior Consultant E.M. Protrain, 16 Hedgerley, Chinnor, Oxford OX9 4TN, England. Tel & Fax 44 (0)1844 353161 Web Site - http://ourworld.compuserve.com/homepages/protrain For Consultancy and Courses in Electron Microscopy World Wide
As an assistance to people who have day to day problems with their equipment the listserver provides a very valuable service and I am often pointing people in this direction to solve their problems.
My area of knowledge is in the operation and maintenance of scanning and transmission electron microscopes so I take an interest in and discussion= s in this area.
Of late I have noticed a tendency for replies to become very complicated,=
so much so that the person asking the question would in my mind only beco= me bemused by the reply. Should we not build the knowledge base step by ste= p rather than leap in with solutions far distant from the actual question/problem?
As a made-up example -
Q - What do I do if I do not have a good quality image on the screen in m= y TEM?
A - Change the phosphor.
Have you seen answers like this?
I like many of you reading this I would not like to inhibit anyone sendin= g in a reply to a question, but should they not place their reply on a leve= l?
Example
A - Assuming that you have prepared the specimen correctly, have the correct alignment, kV and apertures, have the condenser lenses set correctly and are in a well darkened room and have become dark adapted - = if you notice your screen is not as bright as a colleagues TEM perhaps you should consider changing the screen as they fade due to beam damage and contamination?
I look forward to being put in my place! =
Steve Chapman
Senior Consultant E.M. Protrain, 16 Hedgerley, Chinnor, Oxford OX9 4TN, England. Tel & Fax 44 (0)1844 353161 Web Site - http://ourworld.compuserve.com/homepages/protrain For Consultancy and Courses in Electron Microscopy World Wide
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
The string concerning preparation of bacteria touched on delayed second fixation. This is worth a separate discussion on delayed second (usually Os) fixation: Sabatini, first to publish GA as a fixative, also published that Os fixation could be delayed by several months. That seems true for some tissues, which show no ill-effects when compared with the usual, immediate double fixation. However, we found in the lab that other tissues are sensitive to that delay. I used to run a couple of busy service labs and cannot remember specifically which tissues and what structures were affected. It would be interesting to know when delayed double fixation is acceptable and others may have experience to share. I believe that specimen preparation for SEM is never affected by delayed second fixation. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes ********************** www.proscitech.com.au *****
On Thursday, March 04, 1999 11:14 PM, Tina Schwach [SMTP:tschwach-at-tc.umn.edu] wrote: } } On Wed, 3 Mar 1999 14:32:22 -0700, } jwright-at-dugway-emh3.army.mil wrote... } } I have a requirement to process liquid and agar-bound } } pathogenic } } bacteria for SEM & TEM. I've been collecting them in } } cacodylate-buffered 3% glutaraldehyde where the final } } concentration has } } been 1.5% for the liquid suspended bacteria. These have } } been kept in } } the refrigerator. Does anyone have a protocol(s) that } } would lead me to } } SEM and TEM specimens from this point? Have I made a } } mistake already? My } } instruments are a Philips EM-400 and JEOL 6300V. } } } } John Wright } } Microbiologist } } } } West Desert Test Center } } Dugway, UT } } } } } } John, } I have stored samples in primary fixative (glut-para- } ruthenium red in } cacodylate) for several weeks, even months and they appear } to be fine. } For SEM, you may want to place (dry) your samples on some } kind of surface, } ie stainless steel chips, so you'll be able to view them. } You'll have to } do this at the end anyway. You can even place them on } nucleopore filter } membranes. Depending on what you want to see, the agar } strands can get in } the way. } } For the TEM samples, fixation in small 1.5 mL eppendorf } microfuge tubes } works well since you can pellet the samples between } solution changes if } necessary. From the point you are at right now, you } should rinse in your } cacodylate-buffer (I used 0.1M cacodylate with 7.5% } sucrose), then move on } to post-fixation with 1% OsO4 in 0.1M cacodylate (no } sucrose) until pellets } turn dark brown or black (usually 30-min at room temp). } If the pellets are } really big, separate them before osmium treatment to make } sure the entire } pellet is fixed. Rinse in cacodylate buffer (no sucrose), } dehydrate in an } acetone or ethanol series. I use ethanol because I like } to embed in LR } White. After 2 changes in 100%, separate the SEM run (for } critical point } drying and coating) from the TEM samples- move on to resin } infiltration } right in the eppendorf tubes. I usually do 3:1 } (solvent:resin) on a } rotator for 1 hr., 1:1, 1:3, 100% times two and cure } overnight. } Good luck. } } } } }
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Additional to the useful replies relating to the preparation of ball bearings, I would like to add a little regarding the microscopy: Small, clean and undamaged ball bearings are a good instructive aid to explain SEM functioning. This may be of some use to Jonathan when viewing those bearings and to anybody trying to explain some SEM effects. 1 At high kV (say above 20) you may have trouble seeing anything, because fine surface structure and dirt will be invisible. 2 At low kV (say 10 and below) such surface structures will be visible. 3 At low powers, regardless of kV, the ball bearing will appear like a disk, but the outer part of the disk is brighter. This nicely shows that in secondary mode, brightness almost entirely is increased with the angle of incidence. Being a sphere, tilt has no effect on the brightness distribution over that image. 4 A BS detector mounted at an angle (whereas the Robinson and some others are vertical) will make the distinction that the specimen is not a disk but a sphere, because the BS electrons directed away from the detector leave a shadow. 5 A similar effect is produced when the bias current of the secondary detector is turned off and the condenser current is turned down. This floods the secondary detector's scintillator with backscattered electrons and produces a BS image quite suitable for low powers.
Nice teaching exercise, but its useful to know about these effects when actually looking at those bearings. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes ********************** www.proscitech.com.au *****
On Friday, March 05, 1999 6:11 AM, Jonathan Barnard [SMTP:J.Barnard-at-bristol.ac.uk] wrote: } } I am more of a TEM eprson, but I have a suggestion from } watching my } colleagues. } Take an aluminium stub, which would normally go into an } SEM, and using a } 5 mm drill bit just drill a small , 1-2 mm deep, hole in } top so that the } ball bearing will sit in it. Before you mount the bearing, } wash the } bearing in organic solvents first, say two acetone washes } and then two } ethanol washes, and ultrasonically clean the bearing in an } ultrasound } bath with both solvents. } } Dry it off by putting it into an oven at say 150 C for ten } minutes and } then mount it onto the stub using silver-dag mounting } paint (used by } almost everyone here) to fix the bearing onto the stub. If } you have } access to vacuum coating system with a Radio Frequency } inductive plasma } ring attachment or anything that can produce an Argon } plasma, then put } the stub in for ten minutes so that there are no organic } residues left. } Once it is finished you can put it straight into the SEM } knowing that } there should be a clean surface to look at. The silver dag } paint should } earth the bearing to the stub. } } I hope this helps. } } Jon } } -- } ***************************************** } Jonathan Barnard } } Microstructural Physics, } H.H.Wills Physics Laboratory, } University of Bristol, } Tyndall Avenue, } Bristol BS8 1TL. } } 0117 928 9000 ext 8750 } } ***************************************** } }
We lost our Ar/Kr laser very early in its life. (it was still under warranty) A bit of advice was given to us from Zeiss regarding our Ar/Kr laser. That is to run it at 50% power rather than full blast all the time. It seems to be much better now.
} Just to give an opposite example about the laser, } } I'm running an LSM 410 equipped with an Omnichrome Ar/Kr 488/568/647 laser } since 5 years (1750 hours) and i had no problem yet ! Maybe it's just luck } though... } } } At 05:01 PM 25/2/99 -0800, you wrote: } } As for my own experience, I found the 568nm Kr laser sadly unreliable, we } had to } change it twice in the past year. } } Mr. S H Coetzee Electron Microscope Unit Private bag X3 Wits Johannesburg 2050 Tell: +27 11 716 2419 Fax : +27 11 339 3407 E-mail stephan-at-gecko.biol.wits.ac.za
A number of suggestions have been made about the preparation and observation of failed ballbearings. These will certainly give good images and may well be enough to solve the problem, but if the fault lies in the alloy rather than in the physical state of the balls, it will not show up. You might have to consider embedding the bearing in resin and polishing a flat on it. A thin coat of carbon is needed unless you use a conductive resin. Then the BSE image (and EDX if possible) will reveal the internal structure of the alloy.
Eric
---------------------- Dr Eric E. Lachowski University of Aberdeen Department of Chemistry Meston Walk Old Aberdeen AB24 3UE Scotland +44 1224 272934 e.lachowski-at-abdn.ac.uk
} The peaks have suddenly started to move around a bit, although the } resolution stays good. } The problem comes and goes, in its good times the standard deviation } of the position of the Co Ka peak is about 0.4 eV (10 } determinations), but sometimes it's about 10 or even 20 eV. } My thinking is that if it were the detector, the resolution would be } degrading, but it's not, so maybe the culprit is the } (analog) pulse-processor. Anyone got any thoughts on how to pin it } down as being either } a the detector } b the subsequent signal-processing stuff, eg pulse-proc? } Could I successfully test the pulse-proc with a ramp from a standard } signal generator, or would that signal, being relatively clean } compared with that from a detector, not really check it out } rigorously enough?
One thing to note is if your movement is due to gain or offset drifts. If offset drift then both a low peak and high peak will move the same amount. If gain the low will move a little, high will move much more.
Sometimes the gain and offset pots on the pulse processor get "dirty". Run them up and down a few times or clean with electronic cleaner spray.
Disconnect and reconnect every connector. The contacts can get "dirty" over time. The physical active of disconnecting/reconnecting will clean the contacts.
You can test your MCA by using a sliding pulser. We use a Berkley Nucleonics Corp (BNC) GL-3 pulse generator to test every MCA that we ship. It's the only way to really know how well the MCA is working. It's about $5k, not cheap. You really need a very good pulser to test linearity. 10 to 20eV is a really small voltage difference.
Scott ----------------------------------------------------------------------- Scott D. Davilla Phone: 919 489-1757 (tel) 4pi Analysis, Inc. Fax: 919 489-1487 (fax) 3500 Westgate Drive, Suite 403 email: davilla-at-4pi.com Durham, North Carolina 27707-2534 web: http://www.4pi.com
Is it OK to (thin) section the Falcon cell culture inserts in cross section, and would anyone who's done this please contact me directly? I have a question or two about the best way to embed to maximize cell numbers. (I think this has been discussed before.) Thank you. Grace
In the spirit recently suggested by Steve Chapman, try disturbing the cables and processor box. If this affects the peak positions or resolution, there's a good chance you have a loose or dirty connector. Try reseating the PC boards in the processor box.
Larry Thomas Washington State University
---------- From: Ritchie Sims Sent: Friday, March 5, 1999 1:28 PM To: microscopy-at-Sparc5.Microscopy.Com Subject: EDS trouble-shooting
------------------------------------------------------------------------ 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 Experts
The peaks have suddenly started to move around a bit, although the resolution stays good. The problem comes and goes, in its good times the standard deviation of the position of the Co Ka peak is about 0.4 eV (10 determinations), but sometimes it's about 10 or even 20 eV. My thinking is that if it were the detector, the resolution would be degrading, but it's not, so maybe the culprit is the (analog) pulse-processor. Anyone got any thoughts on how to pin it down as being either a the detector b the subsequent signal-processing stuff, eg pulse-proc? Could I successfully test the pulse-proc with a ramp from a standard signal generator, or would that signal, being relatively clean compared with that from a detector, not really check it out rigorously enough?
cheers (well, I try)
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
American Soc List {microscopy-at-Sparc5.Microscopy.Com} X-Mailer: QuickMail Pro 1.5.3 (Mac) X-Priority: 3 MIME-Version: 1.0 Reply-To: Paul Webster {pwebster-at-mailhouse.hei.org} Content-Type: multipart/alternative; boundary="====56495250495150514856===1"
Reply to: RE: Complications With regard to this comment By Steve Chapman, I agree that the discussion = aspect of the listserver is slowly being erroded and would sometimes like = to see the more detailed, carfully considered answers being posted again. = I use the server as a source of assistance but also find that by reading = discussions on subjects I know little about, I can actually learn = something. This sort of passive learning is useful in a busy lab environme= nt.
I am not sure we can hold the solution providers totally responsible for = this errosion either. It is sometimes difficult to know exactly the = problem, the skill of the poster or the environmentally limiting factors = from such brief postings as in the example given by Steve ("What do I do = if I do not have a good quality image on the screen in my TEM?"). Why is = it that some questions are posted in almost annotated form from posters = who do not even provide us with an identification? =
The tendency for discussions to be carried out "off-line" in more detail = is not very beneficial either. It is true that some of the really = detailed discussions, and perhaps some of the comments about particular = products, are best covered in a more private setting, but it would be = great to see edited summaries and final results.
I know writing is tough and it takes some time to write but by putting = more effort into this skill we could improve our list. I am not getting = at non-English writers here but at anyone who writes messages which are so = brief that relevant information is omitted. The more we practice our = writing, the better we get (hopefully). =
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/aemi.htm http://www.hei.org/htm/apw.htm
Steve Chapman wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America =
{HTML} {HEAD} {/HEAD} {BODY} {PRE WIDTH=3D"132"} Reply to: RE: Complications
{/PRE} {FONT FACE=3D"Geneva" SIZE=3D3 = COLOR=3D"#000000"} With regard to this comment By Steve = Chapman, I agree that the discussion aspect = of the listserver is slowly being erroded = and would sometimes like to see the more = detailed, carfully considered answers being = posted again. I use the server as a source = of assistance but also find that by reading = discussions on subjects I know little about, = I can actually learn something. This sort = of passive learning is useful in a busy = lab environment. {BR} {BR} I am not sure we can = hold the solution providers totally responsible = for this errosion either. It is sometimes = difficult to know exactly the problem, the = skill of the poster or the environmentally = limiting factors from such brief postings = as in the example given by Steve ("What = do I do if I do not have a good quality = image on the screen in my TEM?"). Why = is it that some questions are posted in = almost annotated form from posters who do = not even provide us with an identification? = {BR} {BR} The tendency for discussions to be = carried out "off-line" in more = detail is not very beneficial either. It = is true that some of the really detailed = discussions, and perhaps some of the comments = about particular products, are best covered = in a more private setting, but it would = be great to see edited summaries and final = results. {BR} {BR} I know writing is tough and = it takes some time to write but by putting = more effort into this skill we could improve = our list. I am not getting at non-English = writers here but at anyone who writes messages = which are so brief that relevant information = is omitted. The more we practice our writing, = the better we get (hopefully). {BR} {BR} Regards, {BR} {/FONT} {FONT = FACE=3D"Monaco" SIZE=3D1 COLOR=3D"#000000"} {BR} {/FONT} {FONT FACE=3D"Geneva" SIZE=3D3 COLOR=3D"#000000"} Paul = Webster, Ph.D {BR} House Ear Institute {BR} 2100 = West Third Street {BR} Los Angeles, CA 90057 {BR} phone:213 = 273 8026 {BR} fax: 213 413 6739 {BR} e-mail: pwebster-at-hei.org {BR} http://www.hei.org/htm/aemi.htm {BR} http://www.hei.org/htm/apw.htm {/FONT} {FONT = FACE=3D"Monaco" SIZE=3D1 COLOR=3D"#000000"} {BR} {BR} {BR} {/FONT} {FONT FACE=3D"Geneva" = SIZE=3D3 COLOR=3D"#000000"} Steve Chapman wrote: {/FONT} {FONT FACE=3D"Geneva"= = SIZE=3D1 COLOR=3D"#000000"} {BR} >-----------------------------------------------------------------------= - {BR} >The = Microscopy ListServer -- Sponsor: The Microscopy = Society of America {BR}
id xma019702; Fri, 5 Mar 99 13:37:30 -0500 Received: by eastman.com id AA1958416 (5.67b/IDA-1.5 for microscopy-at-Sparc5.Microscopy.Com); Fri, 5 Mar 1999 13:37:57 -0500 Received: from ntmcon02.emn.com by eastman.com with SMTP id AA2032906 (5.67b/SMI-4.1 for {microscopy-at-Sparc5.Microscopy.Com} ); Fri, 5 Mar 1999 13:37:57 -0500 Received: by ntmcon02.emn.com with Internet Mail Service (5.5.2232.9) id {FZ4RM9PM} ; Fri, 5 Mar 1999 13:38:06 -0500 Message-Id: {E788D998AAC8D1119EC10000F8CD1E8A5B42BA-at-ntmail23.emn.com}
Regarding moving peaks in EDS, we once experienced a problem caused by the video monitor on the EDS unit being too close to the wires coming from the detector. Our problem became quite obvious when the resolution began to degrade. Moving the wires fixed the problem. Perhaps your problem is being caused by electromagnetic fields.
Dennis B. Barr (dennbarr-at-eastman.com) Physical Chemistry Research Laboratory Physical & Analytical Chemistry Research Division Eastman Chemical Company Kingsport, TN 37662-5150
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} The peaks have suddenly started to move around a bit, although the } resolution stays good. } The problem comes and goes, in its good times the standard deviation } of the position of the Co Ka peak is about 0.4 eV (10 } determinations), but sometimes it's about 10 or even 20 eV. } My thinking is that if it were the detector, the resolution would be } degrading, but it's not, so maybe the culprit is the } (analog) pulse-processor. Anyone got any thoughts on how to pin it } down as being either } a the detector } b the subsequent signal-processing stuff, eg pulse-proc? } Could I successfully test the pulse-proc with a ramp from a standard } signal generator, or would that signal, being relatively clean } compared with that from a detector, not really check it out } rigorously enough?
One thing to note is if your movement is due to gain or offset drifts. If offset drift then both a low peak and high peak will move the same amount. If gain the low will move a little, high will move much more.
Sometimes the gain and offset pots on the pulse processor get "dirty". Run them up and down a few times or clean with electronic cleaner spray.
Disconnect and reconnect every connector. The contacts can get "dirty" over time. The physical active of disconnecting/reconnecting will clean the contacts.
You can test your MCA by using a sliding pulser. We use a Berkley Nucleonics Corp (BNC) GL-3 pulse generator to test every MCA that we ship. It's the only way to really know how well the MCA is working. It's about $5k, not cheap. You really need a very good pulser to test linearity. 10 to 20eV is a really small voltage difference.
Scott ----------------------------------------------------------------------- Scott D. Davilla Phone: 919 489-1757 (tel) 4pi Analysis, Inc. Fax: 919 489-1487 (fax) 3500 Westgate Drive, Suite 403 email: davilla-at-4pi.com Durham, North Carolina 27707-2534 web: http://www.4pi.com
I have a couple of questions w/r to EELS output in other formats. I am acquiring EELS data on a GIF at another institution, but would like to take the files back to my lab. I saved the files as ASCII X,Y, tagged ASCII, and EMSA/MAS format. I can plot the X,Y data with no problem. However, in the tagged ASCII and the EMSA/MAS format, the data are written in 5 wide rows that wrap the data. I typically use Excel to plot stuff like this, but I don't know how to unwrap the data. What I am doing is going into a Wordprocessor and getting rid of the hard returns and commas and then resaving as a text file. I can automate it a little with macros, but it still takes time.
1. Are there ways to unwrap the data directly into Excel?
2. Is there a public domain EELS program that will plot the data and perhaps do some work with the EMSA/MAS format that will run on a PC or a Mac? (I would prefer PC because our Macs are old and there is not much chance of getting new ones here.)
Thanks in advance.
-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
Lesley S. Bechtold wrote: ============================================ I have never done any material sciences work - I've only ever done biological specimens for EM. Our engineering department has asked me to look at some ball bearings that are failing, as a favour. I know I don't need to fix or dehydrate but do I simply clean them, mount them (using double-sided tape?) and coat them as usual? Or is coating unnecessary? What whould I clean these with? I'm assuming there is grease somewhere that is not good for my vacuum! ================================================== It is interesting how so many different persons, skilled in the art, would approach the same kind of problem so many different ways. Of course, not much information was given either so we could all be those proverbial blind men feeling a different part of the elephant.
We ourselves approach bearing failures somewhat differently. For one thing, at the onset, it is rarely clear whether it is a straight SEM job. If corrosion is involved, for example, the ability to analyze corrosion product is lost if vigorous washing/cleaning procedures are used.
No mention was made of the examination of the raceway, but that is also something of importance in any kind of a failure analysis of this type.
Our approach is to liquid wash in cold solvent, such as acetone and/or heptane, perhaps even xylene, but you don't want a solvent that is "too good ." You want to leave some organic layer on the metal surfaces. We then remove the last vestiges of the lubricant system with exposure to an oxygen plasma such as in our Plasma Prep II plasma etcher. This is a dry process approach, corrosion product is left in situ in place, right where it is, and with EDS, sometimes complimented with Auger, one can learn information about the failure mechanism that would not be learned by straight topographical examination.
For the mounting of the bearing "balls", depending on size, we would always use one of the conductive double sided adhesive products, either carbon sheets or if larger, then Tempfix.
We ourselves believe one should use caution before washing away the information that could be contained in corrosion product.
Disclaimer: SPI Supplies manufactures the Plasma Prep II plasma etcher and supplies the carbon based adhesives. Our Structure Probe services laboratory performs failure analysis as a service on these kinds of samples.
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 ############################ ==================================================
Ritchie Sims wrote: } Dear Experts } } The peaks have suddenly started to move around a bit, although the } resolution stays good. } The problem comes and goes, in its good times the standard deviation } of the position of the Co Ka peak is about 0.4 eV (10 } determinations), but sometimes it's about 10 or even 20 eV. } My thinking is that if it were the detector, the resolution would be } degrading, but it's not, so maybe the culprit is the } (analog) pulse-processor. Anyone got any thoughts on how to pin it } down as being either } a the detector } b the subsequent signal-processing stuff, eg pulse-proc? } Could I successfully test the pulse-proc with a ramp from a standard } signal generator, or would that signal, being relatively clean } compared with that from a detector, not really check it out } rigorously enough? } } cheers (well, I try) } } 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 } }
Dear Ritchie! In similar cases the first I do is the following: I take off the signal cable from preamplifier and processor and measure its resistance, which should be near zero, and then check stability of this resistance during curving this cable. Two times in my practice I have met the case when after several working years the copper core of the cable was corroded as result of chemical interaction with material of cable insulation, and the resistance between many wires of core became rather big and unstable. After change to cable with Teflon insulation the peaks became very (!) stable. Regards.
I have run across the same problems with EMSA format and with other files that store multiple points of data per row.
I have developed macros that insert the necessary number of blank lines (5 for EMSA) following a given row, I select the data in the row, Copy-Special using the transpose function into the blank rows, delete the original row, and move down to the next row, and repeat. It isn't pretty, but it gets the job done.
If I was doing it regularly, I think I would write a program (standalone or Word Basic) that would incorporate the smarts enough to do it all automatically given the file name. As of yet, I have not been so driven. But maybe someone else has already done one.
Good luck.
At 03:54 PM 3/5/99 -0500, you wrote: } I have a couple of questions w/r to EELS output in other formats. I am } acquiring EELS data on a GIF at another institution, but would like to take } the files back to my lab. I saved the files as ASCII X,Y, tagged ASCII, and } EMSA/MAS format. I can plot the X,Y data with no problem. However, in the } tagged ASCII and the EMSA/MAS format, the data are written in 5 wide rows } that wrap the data. I typically use Excel to plot stuff like this, but I } don't know how to unwrap the data. What I am doing is going into a } Wordprocessor and getting rid of the hard returns and commas and then } resaving as a text file. I can automate it a little with macros, but it } still takes time. } } 1. Are there ways to unwrap the data directly into Excel? } } 2. Is there a public domain EELS program that will plot the data and } perhaps do some work with the EMSA/MAS format that will run on a PC or a } Mac? (I would prefer PC because our Macs are old and there is not much } chance of getting new ones here.) } } Thanks in advance. } } -Scott } } Scott D. Walck, Ph.D. } PPG Industries, Inc.
---------------------------------------------------- Warren E. Straszheim 23 Town Engineering Iowa State University Ames IA, 50011-3232
On Fri, 5 Mar 1999, Walck. Scott D. wrote: | |2. Is there a public domain EELS program that will plot the data and |perhaps do some work with the EMSA/MAS format that will run on a PC or a |Mac? (I would prefer PC because our Macs are old and there is not much |chance of getting new ones here.) |
I've been using XlispStat {http://www.xlispstat.org/} for both plotting and analysis ( Mostly EDS but some EELS ).
It's a free, cross-platform (Mac,Windows,Unix) version of Lisp enhanced with vector and matrix arithmetic, math and statistical function, linear and non-linear regression, smoothing, FFTs, and simple object oriented graphics.
It's extensible with function written either in Lisp or C: I've written functions to read & write EMSA/MAS format (roughly -- the specs are somewhat ambiguous), read spectra from a 4pi SpectraEngine on a Mac, convolution and digital filtering and other utilities. ( Haven't quite gotten it all wrapped up into a complete EDS/EELS package yet. )
---| Steven D. Majewski (804-982-0831) {sdm7g-at-Virginia.EDU} |--- ---| Department of Molecular Physiology and Biological Physics |--- ---| University of Virginia Health Sciences Center |--- ---| P.O. Box 10011 Charlottesville, VA 22906-0011 |---
Caldera Open Linux: "Powerful and easy to use!" -- Microsoft(*) (*) {http://www.pathfinder.com/fortune/1999/03/01/mic.html}
Dear Lim (S'pore), There are many infrared camera in the market. You might need to specify = what is the usage are for. By the way, how is Singapore nowadays? Still M= oney no enough?
Thank you for the replies so far. I should have included the info that it's a gain problem ie the zero peak stays in the right place. I guess it boils down to:
given that the resolution stays good, does anyone think that it could be a detector problem, or does everyone think that it's the pulse-processor?
rtch
} From: Self {GLGNOV2/RSIMS} } To: microscopy-at-sparc5.microscopy.com } Subject: EDS trouble-shooting } Date: Fri, 5 Mar 1999 21:28:27 GMT+1200
} Dear Experts } } The peaks have suddenly started to move around a bit, although the } resolution stays good. } The problem comes and goes, in its good times the standard deviation } of the position of the Co Ka peak is about 0.4 eV (10 } determinations), but sometimes it's about 10 or even 20 eV. } My thinking is that if it were the detector, the resolution would be } degrading, but it's not, so maybe the culprit is the } (analog) pulse-processor. Anyone got any thoughts on how to pin it } down as being either } a the detector } b the subsequent signal-processing stuff, eg pulse-proc? } Could I successfully test the pulse-proc with a ramp from a standard } signal generator, or would that signal, being relatively clean } compared with that from a detector, not really check it out } rigorously enough? } } cheers (well, I try) } } 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
As one of the committee members of the MSA/MAS Format I'm perplexed about your comment on the 5 wide rows. That sounds very odd indeed and I recall nothing in the format that calls out that type of coding. Perhaps there is an error somewhere or a misinterpretation of the spectral file format. .
Send me a "private" copy of the 3 files at my ANL address (Zaluzec-at-aaem.amc.anl.gov) and I'll have a look at them when I get back to the US.
Nestor
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I think that the main problem here is that we have a defined M&M=20 =46ormat (MSA/MAS in old parlance, let's face it we are a Microscopy=20 and Microanalysis community and the sooner we realize it the=20 better!), but the XEDS and EELS manufacturers don't fully support it.=20 They either write it but don't read it or read it but don't write it.=20 I think the format should be refined so that the manufacturers will=20 use it and let them have tags in the format similar to the TIFF.
Just my two cents worth (probably start a war, but just my opinion)
Jfm.
John Mansfield PhD CPhys MInstP North Campus Electron Microbeam Analysis Laboratory 417 SRB, University of Michigan 2455 Hayward, Ann Arbor MI 48109-2143 Phone: (734) 936-3352 FAX (734) 936-3352 Cellular Phone: (734) 358-7555 (Leaving a phone message at 936-3352 is preferable to 358-7555) Email: jfmjfm-at-engin.umich.edu URL: http://emalwww.engin.umich.edu/people/jfmjfm/jfmjfm.html Location: Lat. 42=B0 16' 48" Long. 83=B0 43' 48"
Ritchie Sims wrote: } Dear Experts } } The peaks have suddenly started to move around a bit, although the } resolution stays good. } The problem comes and goes, in its good times the standard deviation } of the position of the Co Ka peak is about 0.4 eV (10 } determinations), but sometimes it's about 10 or even 20 eV. } My thinking is that if it were the detector, the resolution would be } degrading, but it's not, so maybe the culprit is the } (analog) pulse-processor. Anyone got any thoughts on how to pin it } down as being either } a the detector } b the subsequent signal-processing stuff, eg pulse-proc? } Could I successfully test the pulse-proc with a ramp from a standard } signal generator, or would that signal, being relatively clean } compared with that from a detector, not really check it out } rigorously enough? } } cheers (well, I try) } } 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 } }
Dear Ritchie! In similar cases the first I do is the following: I take off the signal cable from preamplifier and processor and measure its resistance, which should be near zero, and then check stability of this resistance during curving this cable. Two times in my practice I have met the case when after several working years the copper core of the cable was corroded as result of chemical interaction with material of cable insulation, and the resistance between many wires of core became rather big and unstable. After change to cable with Teflon insulation the peaks became very (!) stable. Regards.
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Hello
Ritchie Sims wrote: } Dear Experts } } The peaks have suddenly started to move around a bit, although the } resolution stays good. } The problem comes and goes, in its good times the standard deviation } of the position of the Co Ka peak is about 0.4 eV (10 } determinations), but sometimes it's about 10 or even 20 eV. } My thinking is that if it were the detector, the resolution would be } degrading, but it's not, so maybe the culprit is the } (analog) pulse-processor. Anyone got any thoughts on how to pin it } down as being either } a the detector } b the subsequent signal-processing stuff, eg pulse-proc? } Could I successfully test the pulse-proc with a ramp from a standard } signal generator, or would that signal, being relatively clean } compared with that from a detector, not really check it out } rigorously enough? } } cheers (well, I try) } } 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 } }
Dear Ritchie! In similar cases the first I do is the following: I take off the signal cable from preamplifier and processor and measure its resistance, which should be near zero, and then check stability of this resistance during curving this cable. Two times in my practice I have met the case when after several working years the copper core of the cable was corroded as result of chemical interaction with material of cable insulation, and the resistance between many wires of core became rather big and unstable. After change to cable with Teflon insulation the peaks became very (!) stable. Regards.
Arnold, Jim 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 currently looking to upgrade my EDS system. I work in a semiconductor } manufacturing facility(CMOS) and I currently have a Cambridge S200 SEM with } Kevex Delta II. } } I have an interest in Oxford, EDAX and a company called EVEX (which I am not } familiar with)? Does anyone have experience with these companies - Good or } Bad? } } I am looking for a light element detector with possibly a WDS for Boron } quantification. } } Thanks in advance. } } Jim Arnold } Microelectronics and Technology Center } AlliedSignal Electronics and Avionics Systems } 9140 Old Annapolis Road } Columbia, MD 21045 } } email: Jim.arnold-at-alliedsignal.com } voice: (410) 964-4118 } fax: (410) 964-5046
Jim, One of my customers sent their Kevex detector to Evex for repair. They didn't complete the repair, they bent the dewar, and they never returned. A very bad bet to do business with them.
Ken Converse owner Quality Images third party SEM service Delta, PA
Lesley S. Bechtold wrote: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Hi, } } I have never done any material sciences work - I've only ever done } biological specimens for EM. Our engineering department has asked me to } look at some ball bearings that are failing, as a favour. I know I don't } need to fix or dehydrate but do I simply clean them, mount them (using } double-sided tape?) and coat them as usual? Or is coating unnecessary? } What whould I clean these with? I'm assuming there is grease somewhere } that is not good for my vacuum! } } Any help would be appreciated!! Thanks in advance..... } } Lesley Bechtold } } Lesley S. Bechtold } Supervisor, Biological Imaging } The Jackson Laboratory } 600 Main St. } Bar Harbor, ME 04609 } 207-288-6191
Lesley, All of the suggestions have been pretty much on the mark. Just two things 1) Get a degausser so you can demagnetize the sample, because if you don't, your resolution will have you swearing at your microscope for its poor performance, 2) a ball bearing that has a good surface may require up to 20kx magnification to see any surface detail. Some dirt on the surface can make finding that surface much easier. Materials failure analysis is a lot of fun!
Ken Converse owner Quality Images third party SEM service Delta, PA
Dear listserved all, I wish to know the examination dates for the next cycle of MSA certification in Electron Microscopy. I know that this question should have been sent to the business office at MSA, but I hope to have a quicky answer from any of the list members. Cheers and have a good day. Mohammed Yousuf A.Rawoof.
} } 1. Are there ways to unwrap EMSA/MAS data directly into Excel? }
Scott, it's a fairly straightforward process to unwrap the data in MSExcel. I recorded a macro to do it a while ago - although we were on Macs then, not PCs, I think I kept it somewhere... If you e-mail a file to me I could write it again, if you like. It is fairly easy to do yourself - I think it's just a matter of cutting and pasting columns. If you can work out a routine to do it once, you can 'record' what you're doing as a macro. All you have to do for subsequent files is to run the macro again.
} 2. Is there a public domain EELS program that will plot the data and } perhaps do some work with the EMSA/MAS format that will run on a PC or a } Mac? (I would prefer PC because our Macs are old and there is not much } chance of getting new ones here.) } } Thanks in advance. } } -Scott } } Scott D. Walck, Ph.D. } PPG Industries, Inc. } Glass Technology Center } Guys Run Rd. (packages) } PO Box 11472 (letters) } Pittsburgh, PA 15238-0472 } } Walck-at-PPG.com } } (412) 820-8651 (office) } (412) 820-8161 (fax) } } } } } }
I am alarmed by your statement. It is Evex Analytical's policy to "always" perform an on-site installation of "any" new detector install or detector repair.
Please be more specific on your customer's name, location, detector serial number, date of service. .
Are you absolutely sure the customer you mentioned sent a "Kevex" detector to "Evex Analytical"? Your prompt reply is appreciated.
Thank you Evex Analytical Peter Tarquinio
-----Original Message----- } From: Kenneth Converse {qualityimages-at-netrax.net} To: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com} ; Arnold, Jim {Jim.Arnold-at-alliedsignal.com}
I am working with self-assembled microstructures that are composed of a sterol and a phosphatidylcholine. I need to be able to attach these structures to glass or plastic walls of a chamber they grow in. What would you recommend? I have tried various commercially coated glass slides, super glue, jewel glue, leather glue, and almost any other glue I could think of. The problem is that my structures grow in an aqueous solution, and I was not able to find an adhesive which would not only glue the structures (super glue did the job actually), but also not dissolve in water.
I would greatly appreciate your suggestions and comments.
Thank you in advance -- Yevgeniya Zastavker.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Yevgeniya V. Zastavker Massachusetts Institute of Technology, Biophysics 77 Massachusetts Ave, Room 13-2038 Cambridge, MA 02139 (617) 253-4826
Sorry to bombard you with questions. This could be the wrong list, but I thought to try anyway. I am looking for crystallographic data of various sterols, and in particular (MAJORLY) I need information on the crystal angles of various sterols. Does anybody know of a good source for this information? I would greatly appreciate your advice.
Thank you very much in advance -- Yevgeniya Zastavker
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Yevgeniya V. Zastavker Massachusetts Institute of Technology, Biophysics 77 Massachusetts Ave, Room 13-2038 Cambridge, MA 02139 (617) 253-4826
The Electron Microscopy Laboratory at New Mexico State University has an open position for an Electron Microscopy Specialist. The laboratory provides transmission and scanning electron microscopy and some light microscopy services for the university research community and a few external organizations, in biological, physical and materials sciences fields. Qualifications: bachelor's degree minimum, master's degree desirable, with at least four years of electron microscopy experience. The preferred candidate will have experience with energy-dispersive x-ray analysis. Experience with digital image capture and analysis, fluorescence microscopy, immunocytochemistry (including immunogold labeling), and laser scanning confocal microscopy is desirable. The candidate must be competent with sample preparation techniques, including vacuum evaporation, sputter coating, critical point drying, support film production, low temperature embedding, and photographic film processing and printing. The successful candidate must be able to work well with researchers, staff, and students, and be able to train graduate and undergraduate students for independent work with relevant techniques and equipment. Duties: operations and routine maintenance of transmission and scanning electron microscopes and associated equipment; fixation, embedding, semi-thin and ultrathin sectioning, staining, and coating of samples; supervision of facility users; record keeping, including billings, budgets, maintenance of instrument and research logs, and researching and designing specimen preparation protocols as required. Salary: DOQ Website: www.nmsu.edu/~personal/postings/professional/ Screening of applicants will begin May 1, 1999 and continue until a candidate is chosen. Applications should include a resume, letter of application and three letters of recommendation.
Apply to: Dr. Reed Dasenbrock Associate Dean/Director Arts and Sciences Research Center New Mexico State University MSC RC, Box 30001 Las Cruces, NM 88003-8001 rdasenbr-at-nmsu.edu
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We have an ETEC SEM available if anyone wants it. (Stockton, CA) It is = in working condition. We bought it new in 1973 and it has been under = contract since that time until Feb. 1, 1999. We also have extra ETEC = parts (power supplies, modules, etc.) We must take it out by March 24, 1999. If anyone wants it, please let me = know or we start chopping it on that day. Thanks Judy M.
Judy Murphy Microscopy Technology Center San Joaquin Delta College 5151 Pacific Ave Stockton, CA 95207 209/954-5284 FAX 209/954-5600 e-mail; jmurphy-at-sjdccd.cc.ca.us
id {1DCJVQ86} ; Mon, 8 Mar 1999 12:10:10 -0800 Message-ID: {3D55EE50922DD21192CC00A0C9A9D8270120F8-at-cas.csudh.edu}
Dear subscribers,
I want to purchase a microtome that sections both paraffin and plastic embedded tissue. I have not purchased a rotary microtome before and I would like your suggestions as to what companies handle rotary microtomes at the present time. I have an ultramicrotome and I know that both types of media can be cut on it but I need a second microtome and have about $12,000 to spend.
Laura J. Robles
Laura J. Robles, PhD. Associate Dean, Student Academic Advancement Professor of Biology MBRS Program Director College of Arts and Sciences California State University, Dominguez Hills 1000 East Victoria Street, Carson CA 90747 310 243-3389, FAX 310 516-4268 lrobles-at-cas.csudh.edu
I think an inexpensive inkjet is the way to go. However, you must try to set the DPI of the printer to match the resolution setting of your digital images. If you capture a digital image at 1024 X800 for instance, you have about 820K of information. Now lets say you plan to print a 4X5 image similar to a Polaroid, then your printer should be no less than 300dpi {4"x5" x (300)2=1.8Mpixel} to accommodate the amount of pixel information from the capturing rate of the image. Hence, a 2048X1600 resolution setting captures 3.2 Mpixels of info, so a 400DPI setting should be used. The idea being to match the capturing info with the amount of pixels the printer can resolve to minimize interpolation...be it upwards or downwards. Not sure what the human eye can resolve tho.
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
We have an ETEC SEM available if anyone wants it. (Stockton, CA) It is = in working condition. We bought it new in 1973 and it has been under = contract since that time until Feb. 1, 1999. We also have extra ETEC parts (power supplies, modules, etc.) We must take it out by March 23, 1999. If anyone wants it, please let me = know or we start chopping it on that day. Thanks Judy M.
Judy Murphy Microscopy Technology Center San Joaquin Delta College 5151 Pacific Ave Stockton, CA 95207 209/954-5284 FAX 209/954-5600 e-mail; jmurphy-at-sjdccd.cc.ca.us
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Dear List, Can anyone explain what does "Reflective" means in Laser Terminology .
I read through a article in a Magazine regarding Laser Marker and in the = article the mentioned that Wood and Paper is 100% reflective . I am lost = and confuse
M.E.Lim Sr Regional Support Engineer QES(Asia Pacific) Sdn Bhd Tel : 603-7241188 ext 214 Fax : 603-7244488 Emails : melim-at-qes.po.my
Can anyone give me tips how to prepare micro-injected cells for TEM. Which cell culture support is best for detaching the cells? Is there any compound which we could micro-inject as a marker?
Try gap-filling super glue (crazy glue), available at any hobby store or it should be at a hardware store. No particular brand, they all work--it's the gap-filling that seems to be important. I used this to glue gelatin specimen blocks to the stage of a Vibratome which was then flooded with phosphate buffer. It holds under water if you let it set. This doesn't take long.
The problem may be the gap-filling property--it may cover your microstructures.
Phil
} I am working with self-assembled microstructures that are composed of a } sterol and a phosphatidylcholine. I need to be able to attach these } structures to glass or plastic walls of a chamber they grow in. What } would you recommend? I have tried various commercially coated glass } slides, super glue, jewel glue, leather glue, and almost any other glue I } could think of. The problem is that my structures grow in an aqueous } solution, and I was not able to find an adhesive which would not only glue } the structures (super glue did the job actually), but also not dissolve in } water. } } I would greatly appreciate your suggestions and comments. } } Thank you in advance -- Yevgeniya Zastavker. } } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ } Yevgeniya V. Zastavker } Massachusetts Institute of Technology, Biophysics } 77 Massachusetts Ave, Room 13-2038 } Cambridge, MA 02139 } (617) 253-4826
****be famous! send in a tech tip or question*** Philip Oshel Technical Editor, Microscopy Today PO Box 620068 Middleton, WI 53562 Voice: (608) 833-2885 Fax: (608) 836-1969 (please make sure my name is on any fax) oshel-at-terracom.net
May I offer a word of caution about the approach below?
While you're correct in saying you need to optimize printing conditions for your image and printer, there will probably be a little more to it than setting up comparable number of pixels. Your image is probably either a 256-level gray scale or a 16 million color (256 levels of red, green and blue). An inkjet can't print that kind of color depth in each pixel. The concepts of "halftoning" or "dithering" need to be considered. As I'm far from the expert, and we don't need a complete textbook on the listserver anyway, so I'll refer you to chapter 2 of "The Image Processing Handbook" by John Russ. As we would expect from Dr. Russ, the material is excellent.
That does bring up a question for me, though. The new HP inkjets have a "PhotoREt" technology which, I believe is supposed to be able to vary the size of the dots it produces, therefore producing better photo-printing results. Has anyone determined whether this is true, or just hype?
Jim Passmore Sr. Analytical Chemist Cryovac Division Sealed Air Corporation
---------- } From: Harry.Ekstrom } To: Microscopy } Subject: FW: Printers for SEM Images } Date: Monday, March 08, 1999 5:11PM } ------------- } } I think an inexpensive inkjet is the way to go. However, you must try to } set the DPI of the printer to match the resolution setting of your digital } images. If you capture a digital image at 1024 X800 for instance, you have } about 820K of information. Now lets say you plan to print a 4X5 image } similar to a Polaroid, then your printer should be no less than 300dpi } {4"x5" x (300)2=1.8Mpixel} to accommodate the amount of pixel information } from the capturing rate of the image. Hence, a 2048X1600 resolution setting } captures 3.2 Mpixels of info, so a 400DPI setting should be used. The idea } being to match the capturing info with the amount of pixels the printer can } resolve to minimize interpolation...be it upwards or downwards. Not sure } what the human eye can resolve tho. } } Good Luck, } Harry Ekstrom } }
Does anyone have or know of a good reference on EM lab safety? I thought there was a book called "Safety in the EM Laboratory"...but I haven't been able to find it. Probably imagined it.
FEI Company Celebrates 50 Years Philips Electron Microscopy with an Anniversary Image Contest (Calling all microscopists!)
Fifty years ago, we delivered the first Philips electron microscope. Since then, our TEMs and SEMs are used for all kinds of applications.
To celebrate the occasion, we're inviting all Philips electron microscope users all over the world to join our special Anniversary Image Contest.
To enter, simply submit prints of one or two of your best images made with a Philips electron microscope, showing the original data bar. Prints only, please! Closing date: 31 July 1999
WIN 1,000 EURO*!
Our jury panel of experts will select the best ten images, five in Life Science and five in Material Science. All ten winners will each be awarded a prize of 1000 Euro*. The winners will be announced in August 1999 on the FEI website at www.feic.com.
The following details must accompany each entry: * Name and contact address of owner * Category: Life Science or Material Science * Description of subject * Type of instrument used * Electron optic magnification * Magnification of print
Please submit your entry to: FEI Company 50 Years Philips EM Celebration P.O. Box 218 5600 MD EINDHOVEN The Netherlands
Digital images cannot be accepted for practical reasons. All submissions must be free of any legal obligations. All entries remain property of their original owners, but contestants consent to the use of their entries for promotional purposes by FEI Company without further compensation. Prizes are not transferable. Taxes are the sole responsibility of the winners. Contest rules are available on the company's website (www.feic.com) or can be requested by fax to +31 40 276 6587. FEI Company will not enter into any other correspondence regarding this contest.
*Actual prize will be the equivalent value in the winner's local currency
Yev, You might try Cell-Tak from Collaborative Biomedical Products, Two Oak Park, Bedford, Mass. 617-275-0004. This is the isolated mussel adhesion protein (map) the marine mussels and barnacles use to attach themselves to rocks and boats etc. It is commonly used by people perform atomic force microscopy to attach their molecules to a surface.
Images & Info at http://www.molbio.princeton.edu/confocal {http://www.molbio.princeton.edu/confocal}
Joseph Goodhouse Confocal / EM Core Laboratory Department of Molecular Biology Princeton University jgoodhouse-at-molbio.princeton.edu {mailto:Jgoodhouse-at-molbio.princeton.edu}
609-258-5432
-----Original Message----- From: Yevgeniya Zastavker [mailto:zhenya-at-critical.mit.edu] Sent: Monday, March 08, 1999 10:43 AM To: microscopy-at-Sparc5.Microscopy.Com Cc: Yevgeniya Zastavker Subject: adhesive for lipids
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I am working with self-assembled microstructures that are composed of a sterol and a phosphatidylcholine. I need to be able to attach these structures to glass or plastic walls of a chamber they grow in. What would you recommend? I have tried various commercially coated glass slides, super glue, jewel glue, leather glue, and almost any other glue I could think of. The problem is that my structures grow in an aqueous solution, and I was not able to find an adhesive which would not only glue the structures (super glue did the job actually), but also not dissolve in water.
I would greatly appreciate your suggestions and comments.
Thank you in advance -- Yevgeniya Zastavker.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Yevgeniya V. Zastavker Massachusetts Institute of Technology, Biophysics 77 Massachusetts Ave, Room 13-2038 Cambridge, MA 02139 (617) 253-4826
I think an inexpensive inkjet is the way to go. However, you must try to set the DPI of the printer to match the resolution setting of your digital images. If you capture a digital image at 1024 X800 for instance, you have about 820K of information. Now lets say you plan to print a 4X5 image similar to a Polaroid, then your printer should be no less than 300dpi {4"x5" x (300)2=1.8Mpixel} to accommodate the amount of pixel information from the capturing rate of the image. Hence, a 2048X1600 resolution setting captures 3.2 Mpixels of info, so a 400DPI setting should be used. The idea being to match the capturing info with the amount of pixels the printer can resolve to minimize interpolation...be it upwards or downwards. Not sure what the human eye can resolve tho.
I just scanned through postings about processing tissue cultures and coverslips for TEM, but I didn't see anything on processing cells grown on 8 well Permanox slides. I will be embedding in PolyBed 812, with a transition through propylene oxide. Unfortunately, the wells don't survive the p.o. step. I would appreciate hearing about any experiences with Permanox slides. Thank you very much.
Sandy Perkins
Laboratory for Neurotoxicity Studies Virginia-Maryland Regional College of Veterinary Medicine Virginia Tech
I agree that you must match the print resolution to the image resolution, but take some exception with your math.
At the crux of the issue is how many printer pixels are required to represent a single image pixel with a satisfactory level of gray or color scale resolution. While a dye sub printer conceptually requires only a single pixel to render the whole range of colors or gray scales, an inkjet printer may require multiple pixels based on the technology used. If an ink jet can only be tunred on and off (like a laser printer), then dithering will be required over a number of pixels to give the appearance of shades of color. More pixels will be needed for smoother or finer gradations. Now I think I heard that new inkjet printers can control the amount of ink at each pixel so that they approach the dye-subs in using only one printer pixel per one image pixel. However, I think better results would be had by allowing something like an 6x6 printer pixel pattern for each image pixel.
Using those assumptions, a 1024x800 image would require 6144x4800 pixels in a 5x4 inch space which requires 1200 dpi printer resolution. Doubling the image resolution to 2048 requires doubling the printer resolution to 2400 dpi. However, the limited resolution (both spatial and color) of the human eye may permit decent images with much less printer resolution, but there would be some loss of image detail.
At 03:11 PM 3/8/99 -0700, Ekstrom, Harry wrote: } } I think an inexpensive inkjet is the way to go. However, you must try to } set the DPI of the printer to match the resolution setting of your digital } images. If you capture a digital image at 1024 X800 for instance, you have } about 820K of information. Now lets say you plan to print a 4X5 image } similar to a Polaroid, then your printer should be no less than 300dpi } {4"x5" x (300)2=1.8Mpixel} to accommodate the amount of pixel information } from the capturing rate of the image. Hence, a 2048X1600 resolution setting } captures 3.2 Mpixels of info, so a 400DPI setting should be used. The idea } being to match the capturing info with the amount of pixels the printer can } resolve to minimize interpolation...be it upwards or downwards. Not sure } what the human eye can resolve tho. } } Good Luck, } Harry Ekstrom
---------------------------------------------------- Warren E. Straszheim 23 Town Engineering Iowa State University Ames IA, 50011-3232
I have an address for Electron Microscopy Safety Handbook. 2nd Edition. 1994. $15.00 Barber, V.C. and J.A. Mascorro (Eds)
San Francisco Press, Box 428600 San Francisco, CA 94142-6800
Hopefully it is still in print, Sally -- Sally Burns Center for Electron Optics B5 Center for Integrated Plant Systems E. Lansing, MI 48824 (517) 355-5004 burnssal-at-pilot.msu.edu
We are electropolishing a two-phase alloy containing a Laves phase and a bcc solid solution. The bcc phase is mainly Vanadium and the Laves phase is mainly HfV2. We are experiencing preferential polishing of the bcc phase leading to marginal TEM specimen quality.
Reply to: RE: Electropolishing of Hf alloy Try adding about 5% acetic acid, polish at -40 C, 100 volts.
Alternate polish: (worked on V-20Ti) = 5.3 g lithium chloride Temp=3D -50 C 11.1 g magnesium perchlorate voltage=3D 190-200 500 ml methanol current=3D 40-50 mA 100 ml butyl cellosolve
Above done with a South Bay 550-B single jet polisher in 1985. A notation mentions even grain boundaries. =
Bernie Kestel Materials Science Division Argonne National Laboratory Argonne, Il., 60439 E-mail {kestel-at-anl.gov} 100 ml butyl cellosolve David E. Luzzi wrote: } We are electropolishing a two-phase alloy containing a Laves phase and a = bcc } solid solution. The bcc phase is mainly Vanadium and the Laves phase is } mainly HfV2. We are experiencing preferential polishing of the bcc phase } leading to marginal TEM specimen quality. } } Our solution is H2SO4 / Methanol / HF / butyl cellusolve } } Does anyone have any suggestions? Thanks in advance. } } David E. Luzzi } Department of Materials Science } University of Pennsylvania } 3231 Walnut Street } Philadelphia, PA 19104-6272 } } 215-898-8366 } 215-573-2128 - fax } luzzi-at-lrsm.upenn.edu {mailto:luzzi-at-lrsm.upenn.edu} } } } } RFC822 header } ----------------------------------- } } Received: from dns2.anl.gov (dns2.anl.gov [146.139.254.3]) by = } horus.et.anl.gov (8.6.11/8.6.11) with ESMTP id NAA23584 for {kestel-at-horus.= et.anl.gov} ; Tue, 9 = } Mar 1999 13:42:50 -0600 } Received: from Sparc5.Microscopy.Com (sparc5.microscopy.com = } [206.69.208.10]) by dns2.anl.gov (8.9.1a/8.6.11) with SMTP id NAA21456; = Tue, 9 Mar 1999 = } 13:42:50 -0600 (CST) } Received: (from daemon-at-localhost) by Sparc5.Microscopy.Com (8.6.11/8.6.= 11) = } id NAA12717 for dist-Microscopy; Tue, 9 Mar 1999 13:07:10 -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 NAA12703 for = } "MicroscopyFilteredEmail-at-msa.microscopy.com"; Tue, 9 Mar 1999 13:06:39 -= 0600 } Received: from sol1.lrsm.upenn.edu (SOL1.LRSM.UPENN.EDU [130.91.56.35]) = by = } Sparc5.Microscopy.Com (8.6.11/8.6.11) with ESMTP id NAA12692 for = } {Microscopy-at-sparc5.microscopy.com} ; Tue, 9 Mar 1999 13:06:24 -0600 } Received: from Luzzi.sol1.lrsm.upenn.edu (LRSM221PC.LRSM.UPENN.EDU = } [130.91.56.249]) } by sol1.lrsm.upenn.edu (8.8.5/8.8.4) with SMTP } id OAA08509 for {Microscopy-at-sparc5.microscopy.com} ; Tue, 9 Mar 1999 = } 14:23:08 -0500 (EST) } From: "David E. Luzzi" {luzzi-at-sol1.lrsm.upenn.edu} } To: {Microscopy-at-sparc5.microscopy.com} } Subject: Electropolictropoliing of Hf alloy } Date: Tue, 9 Mar 1999 14:22:45 -0500 } Message-ID: {001001be6a62$36779b20$f9385b82-at-Luzzi.sol1.lrsm.upenn.edu} } MIME-Version: 1.0 } boundary=3D"----=3D_NextPart_000_0011_01BE6A38.4DAABAE0" } X-Priority: 3 (Normal) } X-MSMail-Priority: Normal } X-Mailer: Microsoft Outlook 8.5, Build 4.71.2232.26 } X-MimeOLE: Produced By Microsoft MimeOLE V4.72.2106.4 } Importance: Normal } X-MS-TNEF-Correlator: 00000000CB8AF9838A20D111B1C200C04FDF38E4A42E5600 } Errors-to: Microscopy-request-at-sparc5.microscopy.com } Content-Type: multipart/mixed; } boundary=3D"----=3D_NextPart_000_0011_01BE6A38.4DAABAE0" } Content-Length: 3968 } Status: = } --====48555149575755524855===1 Content-Type: text/html; charset="US-Ascii" Content-Transfer-Encoding: quoted-printable
{HTML} {HEAD} {/HEAD} {BODY} {PRE = WIDTH=3D"132"} Reply to: RE: Electropolishing of Hf alloy
{/PRE} {FONT FACE=3D"Geneva" SIZE=3D3 COLOR=3D"#000000"} Try = adding about 5% acetic acid, polish at -40 = C, 100 volts. {BR} {BR} Alternate polish: = (worked on V-20Ti) {BR} {BR} 5.3 = g lithium chloride = Temp=3D -50 C {BR} 11.1 g magnesium perchlorate = voltage=3D 190-200 {BR} 500 ml methanol = current=3D 40-50 = mA {BR} 100 ml butyl cellosolve {BR} {BR} = Above done with a South Bay 550-B single = jet polisher in 1985. {BR} A notation mentions = even grain boundaries. {BR} {BR} Bernie = Kestel {BR} Materials Science Division {BR} = Argonne National Laboratory {BR} Argonne, = Il., 60439 E-mail = <kestel-at-anl.gov> {BR} 100 ml = butyl cellosolve {BR} David E. Luzzi wrote: {/FONT} {FONT = FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#000000"} {BR} >We are electropolishing = a two-phase alloy containing a Laves phase = and a bcc {BR} >solid solution. The bcc = phase is mainly Vanadium and the Laves phase = is {BR} >mainly HfV2. We are experiencing = preferential polishing of the bcc phase {BR} >leading = to marginal TEM specimen quality. {BR} > {BR} >Our = solution is H2SO4 / Methanol / HF / butyl = cellusolve {BR} > {BR} >Does anyone have = any suggestions? Thanks in advance. {BR} > {BR} >David = E. Luzzi {BR} >Department of Materials Science {BR} >University = of Pennsylvania {BR} >3231 Walnut Street {BR} >Philadelphia, = PA 19104-6272 {BR} > {BR} >215-898-8366 {BR} >215-573-2128 = - fax {BR} > {/FONT} {FONT FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#0000FF"} {U} luzzi-at-lrsm.= upenn.edu {/U} {/FONT} {FONT = FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#000000"} < {/FONT} {FONT FACE=3D"= Geneva" SIZE=3D1 = COLOR=3D"#0000FF"} {U} mailto:luzzi-at-lrsm.upenn.edu {/U} {/FONT} {FONT = FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#000000"} > {BR} > {BR} > {BR} > {BR} >RFC822 = header {BR} >----------------------------------- {BR} > {BR} > = Received: from dns2.anl.gov (dns2.anl.gov = [146.139.254.3]) by {BR} >horus.et.anl.gov = (8.6.11/8.6.11) with ESMTP id NAA23584 for = < {/FONT} {FONT FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#0000FF"} {U} kestel-at-= horus.et.anl.gov {/U} {/FONT} {FONT = FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#000000"} >; Tue, 9 {BR} >Mar = 1999 13:42:50 -0600 {BR} > Received: from = Sparc5.Microscopy.Com (sparc5.microscopy.com = {BR} >[206.69.208.10]) by dns2.anl.gov = (8.9.1a/8.6.11) with SMTP id NAA21456; Tue, = 9 Mar 1999 {BR} >13:42:50 -0600 (CST) {BR} > = Received: (from {/FONT} {FONT FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#0000FF"} {U} = daemon-at-localhost {/U} {/FONT} {FONT = FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#000000"} ) by Sparc5.Microscopy.Com = (8.6.11/8.6.11) {BR} >id NAA12717 for dist-Microscopy; = Tue, 9 Mar 1999 13:07:10 -0600 {BR} > Received: = from no_more_spam.com (Sparc5 [206.69.208.10]) = by {BR} >Sparc5.Microscopy.Com (8.6.11/8.6.11) =
Arizona Imaging and Microanalysis Society Annual Spring Meeting Thursday, March 11, 1999 University of Arizona Student Union Sr. Ballroom
There is no registration fee.
8:30 - 9:00 Registration
9:00 - 9:15 Welcome Dr. Clark Lantz, President AIMS
9:15 - 10:15 Microanalysis Society Tour Speaker - Applications of SEM/EDX to forensic cases and research related to food product and pharmaceutical tampering and counterfeiting. Dr. Frank Platek, US Food and Drug Administration
10:15 - 10:30 Break
10:30 - 10:55 Metals as documents: some uses of imaging and microanalysis in African history. Dr. David Killick, Anthropology, University of Arizona
10:55 - 11:20 Visualizing surfactant aggregation with atomic force microscopy Jon Wolgemuth, Physics, University of Arizona
11:20 - 11:45 Multi-parametric analysis of cell function in 2- and 3- dimensions by spectral imaging microscopy Dr. Ron Lynch, Physiology, University of Arizona
11:35 - 1:30 Lunch AIMS Business Meeting
1:30 - 1:55 In situ molecular imaging of stress proteins and oxidative damage Dr. Claire Payne, Microbiology & Immunology University of Arizona
1:55 - 2:20 Determining the functional signficance of cytoskeletal proteins using microinjection and transfection techniques Dr. Carol Gregorio, Cell Biology and Anatomy University of Arizona
2:20 - 2:45 Fiberoptic Confocal Microscope for In Vivo Imaging Dr. Art Gmitro, Radiology, University of Arizona
2:45 - 3:00 Break
3:00 - 5:00 Student Presentations
5:30 - 7:30 Banquet ($12 for dinner, contact Suzanne Kelly {suekelly-at-ag.arizona.edu} to reserve dinner)
Microscopy Society of America Tour Speaker - Digital manipulation of acquired images: What is possible vs what is ethical Dr. Jack Kinnamon, University of Denver
.................................................................... : 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"
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Reply to: RE: Processing of micro-injected cells for TEM Dear Raija,
If you are to microinject cells, then the best support is one that gives = access for your needles. I am sure you would prefer to use coverslips. = The material you use will depend on your cells but glass usually works = well. Get the coverslips that have locator grids etched onto them. It is = important to know if you plan to examine the cells for morphology or if = you want to immunolabel them. I will assume you only want to look at = their morphology.
Once you have micro-injected cells and know where they are on the grid (so = don't plate then at too high a density), fix with aldehyde, post fix with = osmium tetroxide, dehydrate and infiltrate with resin (as you would any = piece of tissue for TEM). During the final stages of infiltration (= acetone or propylene oxide) it is wise to transfer the cells, still on the = glass coverslip, to a glass or metal dish. Plastic will dissolve. =
Embed the glass in resin, cells up, at the bottom of an aluminum weigh = boat. Push the glass to the bottom of the dish and pour unpolymerized = resin over. Polymerize by heat and remove from the aluminum dish. Cut = the thin layer of resin away from the back of the coverslip to expose the = glass. Now you can remove the glass by plunging into liquid nitrogen and = rapidly warming few times. It will cause the block to crack but it does = work. Alteratively you can heat the resin, glass-down, on a hot plate and = slide the glass of.. Either way the cells will remain in the resin, as = will the grid locator lines. You should then be able to locate your cells = and cut sections. This is not as easy as I make it seem but it all has = been done before.
If you don't like the idea of embedding at the bottom of a dish, it is = also possible to embed the glass coverslip, cells to the resin, over BEEM = or gelatin capsules, or over flat embedding molds. You will still have to = remove the glass. You could use plastic coverslips and section them too, = but finding your cells will not be easy.
The second part of your question - is there anything you can inject to = identify the cells by EM. Yes, colloidal gold particles can be prepared = that can be microinjected into cells. However, if the gold suspension is = too concentrated it will block the needle. If it is too dilute, it will be= more difficult to detect by EM. =
Immunocytochemistry by request.
Regards,
Paul Webster
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 http://www.hei.org/htm/apw.htm
Raija Sormunen wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America =
by arl-img-10.compuserve.com (8.8.6/8.8.6/2.18) id DAA22626; Wed, 10 Mar 1999 03:04:50 -0500 (EST)
Hi,
The Royal Microscopical Society (Oxford England) produced a series of safety notes for its members that included - the microscopes, embedding a= nd fixation.
I have had use of some of these recently so I do know that they are still=
available.
Steve Chapman
Senior Consultant E.M. Protrain, 16 Hedgerley, Chinnor, Oxford OX9 4TN, England. Tel & Fax 44 (0)1844 353161 Web Site - http://ourworld.compuserve.com/homepages/protrain For Consultancy and Courses in Electron Microscopy World Wide
Dear all, thanks for the tips preparing liposomes for TEM observations. In summary, most answers dealed with negativ staining of the liposomes. We' ve tried so and got good results for an overview. I added all replies to this meeages for all who're interested in this topic. Bernward 1.Don Gantz wrotes: To all who desired more info on fixing and staining of liposomes-- I apologize for the delay in my response. We had a major snowstorm and I lost a day. The initial query by Bernward Laube was about evaluating homogeneity of liposome suspensions ( {100nm diam) and estimating particle volumes. We have been using a osmium fix/negative stain technique for a number of years to look at size distribution of VLDL, chylomicrons, and synthetic lipid emulsions. The osmium fix firms up these particles nicely so that flattening is greatly reduced. In fact, based upon metal shadowing exps. the slight enlargement of diameter due to flattening after fixation is negated by slight shrinkage of particles during processing. The result is an excellent estimate of diameter and volume.
In regard to lipid vesicles/liposomes, our experience with smaller ones of 50nm or so prepared by sonication and processed with only negative stain is that they are preserved pretty well. We would expect some flattening although we have not shadowed these. However, with larger liposomes of 100nm+ diam, we believe OsO4 will aid in preservation and stability and presumably reduce flattening.
If one requires differentiation of lamellar structure (uni vs multi) among particles within a population, negative staining is unreliable because of variability of stain penetration. For this purpose, we use vitreous ice cryomicroscopy as we are fortunate to have it available.
In general our fixation/negative staining procedure is as follows:
1) Add 1 volume of 2% OsO4 in Cacodylate buffer to 2 volumes of emulsion or liposome prep. Fix for 30 mins.+
2) Place small aliquot on freshly flow-discharged carbon formvar-coated grid for a few seconds.
3) Remove and blot off excess fluid and immediately stain with 1% NaPT.
4) After a few seconds, remove and blot excess fluid. Air dry.
Optimum concentration is trial and error. Fixed suspension can be diluted as needed. Hope this helps. 2.Sheila Garcia wrotes:I've just finished my Doctorate Thesis, and I worked preparing liposome. I used phosphotungstic acid in 2% aq solution, neutralized with KOH 1M (PTK). But, before it, I used bacitracin 0,1mg/ml aq sol., as a wetting agent ( D.W.Gregory and B.J.S. Pirie-Journal of Microscopy, v.99,pt.3, dec. 1973,.261-265). Take a coated grid (carbon, formvar), put a drop of bacitracin sol. for 2 min. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the liposome (a dilueted solution), draw off again and add a drop of PTK for 2 min. I hope it could help you. Profa. Dra. Sheila Garcia
Check out the following paper:
"Sequential treatment by phosphotungstic acid and uranyl acetate enhances the adherence of lipid membranes and membrane proteins to hydrophobic EM grids", A. N. Barnakov, Journal of Microscopy. Vol 175, Pt 2, August 1994, pp. 171-174. 3.Joe Neilly:Don,
Could you elaborate on how you would fix liposomes w/ Osmium tetroxide. We have tried the negative stain approach with some success but have to live with the obvious artifacts such as flattening. There are likely other artifacts caused by ionic or chemical changes of the stain. The chemical fixation sounds promising but I'm not sure how this could be done on a liquid sample. 4.Charles Garber:For your information, we have never been successful in quenching a sample fast enough (the larger sample used for SEM work) in order to keep the vesicles from rupturing. So we do this solely by freeze fracture TEM.
Do you know the name of Richard Banfield in the UK? He pretty much was the first person to really commercialize a cryo-SEM fracture system, when he owned the former Hexland Ltd. company and which was purchased by Oxford Instruments.
As of about three or four years ago, he confirmed to me that he too, had never been able to visualize liposomes via cryo SEM because of the quench rate and rupturing problem.
Should you figure out a way to quench and see the structures by SEM, a lot of us would like to know the secret! 5.Ming Chen:The easiest way to do is by negative staining technique. A 1-2% PTA (phosphotungstic acid) soultion is commonly used. It only take a few minutes to do and you can examine it under TEM to see the distribution of liposomes right away. 6. L.R. Melsen:We have looked at liposome using routine negative staining protocol. The vesicles will flatten upon drying, but simple math can reconstruct the volume of the sphere. 7. Charles Butterick: Try negative staining with a 2% ammonium molybdate (aq). Take a coated grid (carbon, formvar, etc.), put a drop of liposomes on top of the grid for a minute. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the ammonium molybdate. After a minute, draw off drop as before and allow the grid(s) to dry. Take it to the TEM. The above procedure is only a starting point. The concentration, stain, and times can all be varied to achieve optimum results. You might check out some EM texts on negative staining for other ideas. Good luck. 8. Maria Ericsson: When I've prepared liposome suspensions for TEM I've just done a simple negative staining and it's worked fine. I adsorb the suspension to a carbon coated copper grid for about 1 minute, blot off excess liquid with a filterpaper and stain with 1-2% aqueous Uranyl Acetate for 1 minute and blot on a filterpaper again. (If there is phosphate in your buffer, you will need to wash in a drop of water before the Uranyl Acetate staining) 9. Marc Schmutz:To observe liposomes without cryo systems is quit a difficult purpose. Pure lipid systems can not be easily seen in negative staining (try always Uranyl acetate and PTA or other stains). And they generaly undergo severe structural changes during the staining process. So to calculate the volume I will not try to do it and furthermore I will not believe in a volume calculate from negative staining images. Myself I'm observing routinely liposomes pure or with proteins or DNA associated and I always use cryo TEM. It's really a easy approach and also very rapid. (You don't need more than half a day per specimen) As you said you don't have access to such a apparatus but why you don't consider collaborating with a lab equiped in cryo ? If you need some more infos about cryo you can ask me and I will try to help you.
Bernward Laube University of Bielefeld Faculty of Biology Department Plant Morphology and Cell Ultrastructure Universittsstrasse 25 Germany 33615 Bielefeld phone: +49 521 1065592 fax: +49 521 1066039 e-mail: b.laube-at-biologie.uni-bielefeld.de http://www.uni-bielefeld.de/biologie/Pflanzenmorphologie
This is a multi-part message in MIME format. --------------B73E5AA6C975623FC057F918 Content-Type: text/plain; charset=us-ascii Content-Transfer-Encoding: 7bit
"...reflector may be either a highly polish metal surface or ...coated. The coating consists of a highly brilliant metallic deposit or a dielectric material." Principles & Practice of Laser Technology.
While both wood & paper are dielectric, their fine porosity would preclude 100% reflective. Typical light reflective number for paper would be 70%. Special paper grades can be much higher. In the case of a coherent laser beam, a porous surface or a surface comprised of fine particles would cause significant scattering.
'Laser marker' refers to a family of products that are used to put bar codes and other identifying markings on products. Since they mark by burning/evaporating away the surface, a marking laser would not work on a 100% reflective surface.
J. Roy Nelson Material Testing Laboratory jrnelson-at-nj1.aae.com
"melim-at-qes.po.my"-at-sparc5.microscopy.com wrote: } } Dear List, } Can anyone explain what does "Reflective" means in Laser Terminology . } } I read through a article in a Magazine regarding Laser Marker and in the article the mentioned that Wood and Paper is 100% reflective . I am lost and confuse } } M.E.Lim } Sr Regional Support Engineer } QES(Asia Pacific) Sdn Bhd } Tel : 603-7241188 ext 214 } Fax : 603-7244488 } Emails : melim-at-qes.po.my --------------B73E5AA6C975623FC057F918 Content-Type: text/x-vcard; charset=us-ascii; name="jrnelson.vcf" Content-Transfer-Encoding: 7bit Content-Description: Card for jrnelson Content-Disposition: attachment; filename="jrnelson.vcf"
Dear all, thanks for the tips preparing liposomes for TEM observations. In summary, most answers dealed with negativ staining of the liposomes. We' ve tried so and got good results for an overview. I added all replies to this message for all those who're interested in this topic. Bernward 1.Don Gantz wrotes: To all who desired more info on fixing and staining of liposomes-- I apologize for the delay in my response. We had a major snowstorm and I lost a day. The initial query by Bernward Laube was about evaluating homogeneity of liposome suspensions ( {100nm diam) and estimating particle volumes. We have been using a osmium fix/negative stain technique for a number of years to look at size distribution of VLDL, chylomicrons, and synthetic lipid emulsions. The osmium fix firms up these particles nicely so that flattening is greatly reduced. In fact, based upon metal shadowing exps. the slight enlargement of diameter due to flattening after fixation is negated by slight shrinkage of particles during processing. The result is an excellent estimate of diameter and volume.
In regard to lipid vesicles/liposomes, our experience with smaller ones of 50nm or so prepared by sonication and processed with only negative stain is that they are preserved pretty well. We would expect some flattening although we have not shadowed these. However, with larger liposomes of 100nm+ diam, we believe OsO4 will aid in preservation and stability and presumably reduce flattening.
If one requires differentiation of lamellar structure (uni vs multi) among particles within a population, negative staining is unreliable because of variability of stain penetration. For this purpose, we use vitreous ice cryomicroscopy as we are fortunate to have it available.
In general our fixation/negative staining procedure is as follows:
1) Add 1 volume of 2% OsO4 in Cacodylate buffer to 2 volumes of emulsion or liposome prep. Fix for 30 mins.+
2) Place small aliquot on freshly flow-discharged carbon formvar-coated grid for a few seconds.
3) Remove and blot off excess fluid and immediately stain with 1% NaPT.
4) After a few seconds, remove and blot excess fluid. Air dry.
Optimum concentration is trial and error. Fixed suspension can be diluted as needed. Hope this helps. 2.Sheila Garcia wrotes:I've just finished my Doctorate Thesis, and I worked preparing liposome. I used phosphotungstic acid in 2% aq solution, neutralized with KOH 1M (PTK). But, before it, I used bacitracin 0,1mg/ml aq sol., as a wetting agent ( D.W.Gregory and B.J.S. Pirie-Journal of Microscopy, v.99,pt.3, dec. 1973,.261-265). Take a coated grid (carbon, formvar), put a drop of bacitracin sol. for 2 min. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the liposome (a dilueted solution), draw off again and add a drop of PTK for 2 min. I hope it could help you. Profa. Dra. Sheila Garcia
Check out the following paper:
"Sequential treatment by phosphotungstic acid and uranyl acetate enhances the adherence of lipid membranes and membrane proteins to hydrophobic EM grids", A. N. Barnakov, Journal of Microscopy. Vol 175, Pt 2, August 1994, pp. 171-174. 3.Joe Neilly:Don,
Could you elaborate on how you would fix liposomes w/ Osmium tetroxide. We have tried the negative stain approach with some success but have to live with the obvious artifacts such as flattening. There are likely other artifacts caused by ionic or chemical changes of the stain. The chemical fixation sounds promising but I'm not sure how this could be done on a liquid sample. 4.Charles Garber:For your information, we have never been successful in quenching a sample fast enough (the larger sample used for SEM work) in order to keep the vesicles from rupturing. So we do this solely by freeze fracture TEM.
Do you know the name of Richard Banfield in the UK? He pretty much was the first person to really commercialize a cryo-SEM fracture system, when he owned the former Hexland Ltd. company and which was purchased by Oxford Instruments.
As of about three or four years ago, he confirmed to me that he too, had never been able to visualize liposomes via cryo SEM because of the quench rate and rupturing problem.
Should you figure out a way to quench and see the structures by SEM, a lot of us would like to know the secret! 5.Ming Chen:The easiest way to do is by negative staining technique. A 1-2% PTA (phosphotungstic acid) soultion is commonly used. It only take a few minutes to do and you can examine it under TEM to see the distribution of liposomes right away. 6. L.R. Melsen:We have looked at liposome using routine negative staining protocol. The vesicles will flatten upon drying, but simple math can reconstruct the volume of the sphere. 7. Charles Butterick: Try negative staining with a 2% ammonium molybdate (aq). Take a coated grid (carbon, formvar, etc.), put a drop of liposomes on top of the grid for a minute. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the ammonium molybdate. After a minute, draw off drop as before and allow the grid(s) to dry. Take it to the TEM. The above procedure is only a starting point. The concentration, stain, and times can all be varied to achieve optimum results. You might check out some EM texts on negative staining for other ideas. Good luck. 8. Maria Ericsson: When I've prepared liposome suspensions for TEM I've just done a simple negative staining and it's worked fine. I adsorb the suspension to a carbon coated copper grid for about 1 minute, blot off excess liquid with a filterpaper and stain with 1-2% aqueous Uranyl Acetate for 1 minute and blot on a filterpaper again. (If there is phosphate in your buffer, you will need to wash in a drop of water before the Uranyl Acetate staining) 9. Marc Schmutz:To observe liposomes without cryo systems is quit a difficult purpose. Pure lipid systems can not be easily seen in negative staining (try always Uranyl acetate and PTA or other stains). And they generaly undergo severe structural changes during the staining process. So to calculate the volume I will not try to do it and furthermore I will not believe in a volume calculate from negative staining images. Myself I'm observing routinely liposomes pure or with proteins or DNA associated and I always use cryo TEM. It's really a easy approach and also very rapid. (You don't need more than half a day per specimen) As you said you don't have access to such a apparatus but why you don't consider collaborating with a lab equiped in cryo ? If you need some more infos about cryo you can ask me and I will try to help you.
Bernward Laube University of Bielefeld =46aculty of Biology Department Plant Morphology and Cell Ultrastructure Universit=D1tsstrasse 25 Germany 33615 Bielefeld phone: +49 521 1065592 fax: +49 521 1066039 e-mail: b.laube-at-biologie.uni-bielefeld.de http://www.uni-bielefeld.de/biologie/Pflanzenmorphologie
We use 4 and 8 well Labtek=A9 Chamber Slides (permanox slides).... substituting Ethanol for the Propylene Oxide. They are my favorite slides for processing cell cultures.
See the following publication for more information:
"Subcellular Localization of SV2 and Other Secretory Vesicle Components in PC12 Cells by an Efficient Method of Preembedding EM Immunocytochemistry for Cell Cultures", Tanner, Ploug and Tao-Cheng, Journal of Histochemistry and Cyrtochemistry, Vol. 44, No. 12, pp. 1481-1488, 1996.
If you have any questions.. feel free to contact me.
Virginia Tanner Crocker
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******************************************************************* Virginia Tanner Crocker Biologist NIH, NINDS EM Facility, Bldg 36, Room 3B24 Bethesda, MD 20892
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Thanks to all who replied. The ETEC has been spoken for. If for some = reason it does not leave as scheduled I will keep the names of those who = responded, just in case, since it MUST GO.
Thanks again, Judy Murphy
Judy Murphy Microscopy Technology Center San Joaquin Delta College 5151 Pacific Ave Stockton, CA 95207 209/954-5284 FAX 209/954-5600 e-mail; jmurphy-at-sjdccd.cc.ca.us
by mail-ewr-3.pilot.net (Pilot/8.8.8) with ESMTP id MAA19999; Wed, 10 Mar 1999 12:27:37 -0500 (EST) Received: from ridexch1.rid.com ([148.189.116.16]) by mailgw.bi-pharm.com with ESMTP id MAA24037; Wed, 10 Mar 1999 12:28:05 -0500 (EST) Received: by RIDEXCH1 with Internet Mail Service (5.5.2232.9) id {F437PDQP} ; Wed, 10 Mar 1999 12:27:36 -0500 Message-ID: {5063A0AB7328D211BCAA0008C7A4467704758B-at-RIDMSG05}
Sandy:
The p.o. step is not necessary. Dehydration with any of the 812 replacements can be done through EtOH alone. I use at least 3x 100%, then grade thru the EtOH:epoxy at 2:1, 1:1 and 1:2, then into pure resin. The only caveat is that the EtOH and resin must be very carefully mixed--both to ensure complete mixing and to avoid the creation of bubbles in the mix. I also use only freshly prepared resin once I reach the 1:1 stage, but have had no problem using older batches that were stored at -20 C and thawed just prior to use.
Roger Moretz Toxicology & Safety Assessment
} -----Original Message----- } From: Sandy Perkins [SMTP:skperkin-at-vt.edu] } Sent: Tuesday, March 09, 1999 11:01 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: TEM-cells on Permanox slides } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Hi- } } I just scanned through postings about processing tissue cultures and } coverslips for TEM, but I didn't see anything on processing cells grown on } 8 well Permanox slides. I will be embedding in PolyBed 812, with a } transition through propylene oxide. Unfortunately, the wells don't } survive } the p.o. step. I would appreciate hearing about any experiences with } Permanox slides. Thank you very much. } } Sandy Perkins } } Laboratory for Neurotoxicity Studies } Virginia-Maryland Regional College } of Veterinary Medicine } Virginia Tech } }
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Dear all, thanks for the tips preparing liposomes for TEM observations. I= n summary, most answers dealed with negativ staining of the liposomes. We' ve tried so and got good results for an overview. I added all replies to this message for all those who're interested in this topic. Bernward 1.Don Gantz wrotes: To all who desired more info on fixing and staining of liposomes-= - I apologize for the delay in my response. We had a major snowstorm and I lost a day. The initial query by Bernward Laube was about evaluating homogeneity of liposome suspensions ( {100nm diam) and estimating particle volumes. We have been using a osmium fix/negative stain technique for a number of years to look at size distribution of VLDL, chylomicrons, and syntheti= c lipid emulsions. The osmium fix firms up these particles nicely so that flattening is greatly reduced. In fact, based upon metal shadowing exps. the slight enlargement of diameter due to flattening after fixation is negated by slight shrinkage of particles during processing. The result i= s an excellent estimate of diameter and volume.
In regard to lipid vesicles/liposomes, our experience with smaller ones of 50nm or so prepared by sonication and processed with only negative stain is that they are preserved pretty well. We would expect some flattening although we have not shadowed these. However, wit= h larger liposomes of 100nm+ diam, we believe OsO4 will aid in preservation and stability and presumably reduce flattening.
If one requires differentiation of lamellar structure (uni vs multi) among particles within a population, negative staining is unreliable because of variability of stain penetration. For this purpose, we use vitreous ice cryomicroscopy as we are fortunate to have it available.
In general our fixation/negative staining procedure is as follows:
1) Add 1 volume of 2% OsO4 in Cacodylate buffer to 2 volumes of emulsion or liposome prep. Fix for 30 mins.+
2) Place small aliquot on freshly flow-discharged carbon formvar-coated grid for a few seconds.
3) Remove and blot off excess fluid and immediately stain with 1% NaPT.
4) After a few seconds, remove and blot excess fluid. Air dry.
Optimum concentration is trial and error. Fixed suspension can be diluted as needed. Hope this helps. 2.Sheila Garcia wrotes:I've just finished my Doctorate Thesis, and I worked preparing liposome. I used phosphotungstic acid in 2% aq solution, neutralized with KOH 1M (PTK). But, before it, I used bacitracin 0,1mg/ml aq sol., as a wetting agent ( D.W.Gregory and B.J.S. Pirie-Journal of Microscopy, v.99,pt.3, dec. 1973,.261-265). Take a coated grid (carbon, formvar), put a drop of bacitracin sol. for 2 min. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the liposome (a dilueted solution), draw off again and add a drop of PTK for = 2 min. I hope it could help you. Profa. Dra. Sheila Garcia
Check out the following paper:
"Sequential treatment by phosphotungstic acid and uranyl acetate enhances the adherence of lipid membranes and membrane proteins to hydrophobic EM grids", A. N. Barnakov, Journal of Microscopy. Vol 175, Pt 2, August 1994, pp. 171-174. 3.Joe Neilly:Don,
Could you elaborate on how you would fix liposomes w/ Osmium tetroxide. W= e have tried the negative stain approach with some success but have to live with the obvious artifacts such as flattening. There are likely other artifacts caused by ionic or chemical changes of the stain. The chemical fixation sounds promising but I'm not sure how this could be done on a liquid sample. 4.Charles Garber:For your information, we have never been successful in quenching a sample fast enough (the larger sample used for SEM work) in order to keep the vesicles from rupturing. So we do this solely by freeze fracture TEM.
Do you know the name of Richard Banfield in the UK? He pretty much was the first person to really commercialize a cryo-SEM fracture system, when he owned the former Hexland Ltd. company and which was purchased by Oxfor= d Instruments.
As of about three or four years ago, he confirmed to me that he too, had never been able to visualize liposomes via cryo SEM because of the quench rate and rupturing problem.
Should you figure out a way to quench and see the structures by SEM, a lo= t of us would like to know the secret! 5.Ming Chen:The easiest way to do is by negative staining technique. A 1-2% PTA (phosphotungstic acid) soultio= n is commonly used. It only take a few minutes to do and you can examine it under TEM to see the distribution of liposomes right away. 6. L.R. Melsen:We have looked at liposome using routine negative staining protocol. The vesicles will flatten upon drying, but simple math can reconstruct the volume of the sphere. 7. Charles Butterick: Try negative staining with a 2% ammonium molybdate (aq). Take a coated grid (carbon, formvar, etc.), put a drop of liposomes on top of the grid for a minute. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the ammonium molybdate. After a minute, draw off drop as before and allow the grid(s) to dry. Take it to the TEM. The above procedure is only a starting point. The concentration, stain, and times can all be varied to achieve optimum results. You might check out some EM text= s on negative staining for other ideas. Good luck. 8. Maria Ericsson: When I've prepared liposome suspensions for TEM I've just done a simple negative staining and it's worked fine. I adsorb the suspension to a carbon coated copper grid for about 1 minute, blot off excess liquid with a filterpaper and stain with 1-2% aqueous Uranyl Acetate for 1 minute and blot on a filterpaper again. (If there is phosphate in your buffer, you will need to wash in a drop of water before the Uranyl Acetate staining) 9. Marc Schmutz:To observe liposomes without cryo systems is quit a difficult purpose. Pure lipid systems can not be easily seen in negative staining (try always Uranyl acetate and PTA or other stains). And they generaly undergo severe structural changes during the staining process. So to calculate the volume I will not try to do it and furthermore I will not believe in a volume calculate from negative staining images. Myself I'm observing routinely liposomes pure or with proteins or DNA associated and I always use cryo TEM. It's really a easy approach and also very rapid. (You don't need more than half a day per specimen) As you said you don't have access to such a apparatus but why you don't consider collaborating with a lab equiped in cryo ? If you need some more infos about cryo you can ask me and I will try to help you.
Bernward Laube University of Bielefeld Faculty of Biology Department Plant Morphology and Cell Ultrastructure Universit=D1tsstrasse 25 Germany 33615 Bielefeld phone: +49 521 1065592 fax: +49 521 1066039 e-mail: b.laube-at-biologie.uni-bielefeld.de http://www.uni-bielefeld.de/biologie/Pflanzenmorphologie
Hello, Does anyone have a suggesstion on how to fix/reset the advance mechanism to a reichert e ultramicrotome. The mechanism on the left side (0.5 um to 2 um advance) does not work, I have to get close with the coarse knife advance then use the electronic advance which makes alignment tedious. Thanks
We are thinking of purchasing a negative scanner for use with TEM negatives from our Jeol 2000-FX, and also for SEM negatives. A scanner has been recommended to us: the Agfa Duoscan T2500, which has a resolution (hardware) of 1250 x 2500 dpi and a density of 3.4D. (The scans would be output to a Kodak DS 8650 PS printer)
We need the quality of the scans to match the quality of the standard darkroom enlarger if possible, as we would like to 'go digital' at least for routine work. Does anyone have experience of routine negative scanning for TEM prints, with this or other scanners, and if so, is it realistic to expect such high quality?
Also, what additional image processing software would people recommend we got to go along with this?
Any advice would be appreciated.
Caspar
Caspar McConville, Ph.D. Technical Specialist New York State College of Ceramics Alfred University
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America=20
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Dear all, thanks for the tips preparing liposomes for TEM observations. I= n summary, most answers dealed with negativ staining of the liposomes. We' ve tried so and got good results for an overview. I added all replies to this message for all those who're interested in this topic. Bernward 1.Don Gantz wrotes: To all who desired more info on fixing and staining of liposomes-= - I apologize for the delay in my response. We had a major snowstorm and I lost a day. The initial query by Bernward Laube was about evaluating homogeneity of liposome suspensions ( {100nm diam) and estimating particle volumes. We have been using a osmium fix/negative stain technique for a number of years to look at size distribution of VLDL, chylomicrons, and syntheti= c lipid emulsions. The osmium fix firms up these particles nicely so that flattening is greatly reduced. In fact, based upon metal shadowing exps. the slight enlargement of diameter due to flattening after fixation is negated by slight shrinkage of particles during processing. The result i= s an excellent estimate of diameter and volume.
In regard to lipid vesicles/liposomes, our experience with smaller ones of 50nm or so prepared by sonication and processed with only negative stain is that they are preserved pretty well. We would expect some flattening although we have not shadowed these. However, wit= h larger liposomes of 100nm+ diam, we believe OsO4 will aid in preservation and stability and presumably reduce flattening.
If one requires differentiation of lamellar structure (uni vs multi) among particles within a population, negative staining is unreliable because of variability of stain penetration. For this purpose, we use vitreous ice cryomicroscopy as we are fortunate to have it available.
In general our fixation/negative staining procedure is as follows:
1) Add 1 volume of 2% OsO4 in Cacodylate buffer to 2 volumes of emulsion or liposome prep. Fix for 30 mins.+
2) Place small aliquot on freshly flow-discharged carbon formvar-coated grid for a few seconds.
3) Remove and blot off excess fluid and immediately stain with 1% NaPT.
4) After a few seconds, remove and blot excess fluid. Air dry.
Optimum concentration is trial and error. Fixed suspension can be diluted as needed. Hope this helps. 2.Sheila Garcia wrotes:I've just finished my Doctorate Thesis, and I worked preparing liposome. I used phosphotungstic acid in 2% aq solution, neutralized with KOH 1M (PTK). But, before it, I used bacitracin 0,1mg/ml aq sol., as a wetting agent ( D.W.Gregory and B.J.S. Pirie-Journal of Microscopy, v.99,pt.3, dec. 1973,.261-265). Take a coated grid (carbon, formvar), put a drop of bacitracin sol. for 2 min. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the liposome (a dilueted solution), draw off again and add a drop of PTK for = 2 min. I hope it could help you. Profa. Dra. Sheila Garcia
Check out the following paper:
"Sequential treatment by phosphotungstic acid and uranyl acetate enhances the adherence of lipid membranes and membrane proteins to hydrophobic EM grids", A. N. Barnakov, Journal of Microscopy. Vol 175, Pt 2, August 1994, pp. 171-174. 3.Joe Neilly:Don,
Could you elaborate on how you would fix liposomes w/ Osmium tetroxide. W= e have tried the negative stain approach with some success but have to live with the obvious artifacts such as flattening. There are likely other artifacts caused by ionic or chemical changes of the stain. The chemical fixation sounds promising but I'm not sure how this could be done on a liquid sample. 4.Charles Garber:For your information, we have never been successful in quenching a sample fast enough (the larger sample used for SEM work) in order to keep the vesicles from rupturing. So we do this solely by freeze fracture TEM.
Do you know the name of Richard Banfield in the UK? He pretty much was the first person to really commercialize a cryo-SEM fracture system, when he owned the former Hexland Ltd. company and which was purchased by Oxfor= d Instruments.
As of about three or four years ago, he confirmed to me that he too, had never been able to visualize liposomes via cryo SEM because of the quench rate and rupturing problem.
Should you figure out a way to quench and see the structures by SEM, a lo= t of us would like to know the secret! 5.Ming Chen:The easiest way to do is by negative staining technique. A 1-2% PTA (phosphotungstic acid) soultio= n is commonly used. It only take a few minutes to do and you can examine it under TEM to see the distribution of liposomes right away. 6. L.R. Melsen:We have looked at liposome using routine negative staining protocol. The vesicles will flatten upon drying, but simple math can reconstruct the volume of the sphere. 7. Charles Butterick: Try negative staining with a 2% ammonium molybdate (aq). Take a coated grid (carbon, formvar, etc.), put a drop of liposomes on top of the grid for a minute. Draw off drop with with a piece of torn filter paper. Before the grid dries, add a drop of the ammonium molybdate. After a minute, draw off drop as before and allow the grid(s) to dry. Take it to the TEM. The above procedure is only a starting point. The concentration, stain, and times can all be varied to achieve optimum results. You might check out some EM text= s on negative staining for other ideas. Good luck. 8. Maria Ericsson: When I've prepared liposome suspensions for TEM I've just done a simple negative staining and it's worked fine. I adsorb the suspension to a carbon coated copper grid for about 1 minute, blot off excess liquid with a filterpaper and stain with 1-2% aqueous Uranyl Acetate for 1 minute and blot on a filterpaper again. (If there is phosphate in your buffer, you will need to wash in a drop of water before the Uranyl Acetate staining) 9. Marc Schmutz:To observe liposomes without cryo systems is quit a difficult purpose. Pure lipid systems can not be easily seen in negative staining (try always Uranyl acetate and PTA or other stains). And they generaly undergo severe structural changes during the staining process. So to calculate the volume I will not try to do it and furthermore I will not believe in a volume calculate from negative staining images. Myself I'm observing routinely liposomes pure or with proteins or DNA associated and I always use cryo TEM. It's really a easy approach and also very rapid. (You don't need more than half a day per specimen) As you said you don't have access to such a apparatus but why you don't consider collaborating with a lab equiped in cryo ? If you need some more infos about cryo you can ask me and I will try to help you.
Bernward Laube University of Bielefeld Faculty of Biology Department Plant Morphology and Cell Ultrastructure Universit=D1tsstrasse 25 Germany 33615 Bielefeld phone: +49 521 1065592 fax: +49 521 1066039 e-mail: b.laube-at-biologie.uni-bielefeld.de http://www.uni-bielefeld.de/biologie/Pflanzenmorphologie
I'm trying to cobble together a new probe. So far, I have one WDS spectrometer which used to be on an 840. Can anyone tell me exactly which JEOL SEMs it will fit on to? I am pretty sure it will go onto 6300 and 6400, and maybe also the 733. So far I've had no luck getting this info from JEOL, but if someone within their organisation can help, great. Also, does anyone know whether a two-crystal spectro can be transformed into a 4-crystal type? I've been told that it can't be done in the field, but can the factory do it?
Anybody got any more of these spectros that they'd be willing to sell? And maybe also an 840 or 840A?
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
I have been very happy with the results from my Polaroid Sprintscan 45 (~$8500). For a TEM neg, it will digitize at 2000dpi and it will output 12 bits. I can't remember the optical density but I think it is similar to what you are quoting here, but you can look it up on their web site. For a 35 mm slide, it will do 4000 dpi. You have to ask them for a special TEM negative carrier that fits into their 4x5 holder. I use 300 dpi as the standard for what kind of enlargement I can get because my HP 890C (which does a great job on photo deluxe paper) is 300 dpi and the two sub-dye printers that I have access to are both 300 dpi. This gives a usable enlargement factor of 2000/300 = 6.7x printing to these printers. It is also very quick. I think that Polaroid could definitely improve their user control interface a bit, but for the most part it works well. I have had some problems with the computer recognizing the scanner on two systems, (one with a Sprintscan 35 and the other with a Sprintscan 45) that have flatbed scanners attached and that are on. The solution is simply to turn the flatbed power off and reopen the program.
I think that the Duoscan is a flatbed. You have to be careful with putting the negatives on the glass because you can get Newton rings in you images. I have a Umax Powerlook II flatbed that is 600 x 1200 dpi that I sometimes use and you can sometimes see them. You need a mask to lift it off the glass. I have asked the listserver in the past if that defocuses the scanned image, but I did not get a satisfactory answer. I use the flatbed as a contact printer by scanning 6 images at once held in Neg-a-file sheets and digitizing to 150 dpi. This is the minimum value that I can still read the numbers on the JEOL negatives. This is actually better than making contacts prints in the darkroom, because I can select areas to adjust the contrast and brightness independently. This makes BF, DF, and diffraction images come out well even if they are on one sheet.
I highly recommend Adobe Photoshop (version 4 is what I have) coupled with John Russ' Image Processing Toolkit Plug-ins. I Import the images as 12 bits (16bit), adjust the levels to what I want, and then convert the images to 8 bits. You can then use all of Photoshop's features.
-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: Caspar McConville To: Microscopy List Server -----------------------------------------------------------------------.
We are thinking of purchasing a negative scanner for use with TEM negatives from our Jeol 2000-FX, and also for SEM negatives. A scanner has been recommended to us: the Agfa Duoscan T2500, which has a resolution (hardware) of 1250 x 2500 dpi and a density of 3.4D. (The scans would be output to a Kodak DS 8650 PS printer)
We need the quality of the scans to match the quality of the standard darkroom enlarger if possible, as we would like to 'go digital' at least for routine work. Does anyone have experience of routine negative scanning for TEM prints, with this or other scanners, and if so, is it realistic to expect such high quality?
Also, what additional image processing software would people recommend we got to go along with this?
Any advice would be appreciated.
Caspar
Caspar McConville, Ph.D. Technical Specialist New York State College of Ceramics Alfred University
One approach might be to plunge freeze on filmed grids and then clamp these to a standard cryoSEM specimen support. The support could have a hole drilled through it for STEM. Because of the water film thickness, it could be sublimed by freeze drying, although any dissolved salts would be left behind.
A similar approach would be to quench the liposomes on something like aluminium foil, in a size & shape which could then be clamped under a thin ring which is drilled to screw to a standard suppport. These attachments can be done quite easily under a shallow depth of liquid nitrogen in a polystyrene container.
Hoping these ideas are useful to someone
Keith Ryan Marine Biological Association Plymouth, UK
Hello, The Newtonian Ring problem might be corrected the way it was done on anti-Newton glass slides, in days gone by. They used glass which was slightly etched on the side which went next to the film. I am not suggesting you etch the glass face of your scanner but it might be worth experimenting on a spare piece of glass. The etch is ever so slight; maybe just acid fumes would do the trick. This didn't seem to degrade projected slides. Just a suggestion. Good luck.
Alex Greene SCIENTIFIC INSTRUMENTATION SERVICES, INC. Number 499, Post Office Box 19400 Austin, Texas 78760 Phone 512/282-5507 Fax 512/280-0702
TEM & SEM Maintenance -----Original Message----- } From: Walck. Scott D. {walck-at-ppg.com} To: Caspar McConville {mcconville-at-olsen.alfred.edu} ; Micro {microscopy-at-Sparc5.Microscopy.Com}
Tamara, The copy I have of "Safety in the EM Lab..." is from San Francisco Press, ISBN 0-911302-56-5, 1985. You should be able to get it ordered with that info. Good luck.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Winston W. Wiggins, Supervisor 09 Mar 1999 4:30 PM CRC-Electron Microscopy Lab Ofc: 704-355-1267 Carolinas Medical Center Lab: 704-355-7220 P.O. Box 32861 Fax: 704-355-7648 Charlotte, NC 28232-2861 USA Eml: WWiggins-at-Carolinas.org ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
} -----Original Message----- } From: Tamara Howard [SMTP:howard-at-cshl.org] } Sent: Tuesday, March 09, 1999 9:49 AM } To: Microscopy Listserver } Subject: EM safety book? } } ------------------------------------------------------------------------ } 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 or know of a good reference on EM lab safety? I thought } there was a book called "Safety in the EM Laboratory"...but I haven't been } able to find it. Probably imagined it. } } Thanks for any help anyone can give! } } Tamara Howard } CSHL } }
I think you are making an excellent choice with the new Agfa DuoScan T2500. You will appreciate the 3.4 D on this unique flatbed scanner. The glass less negative carriers fit into the lower tray, which slides into a precise position. The negatives will not contact any surface. I have just ordered also a DuoScan T2500. You will learn more about it at:
One of the most versatile imaging software is CorelDraw8 and CorelPHOTO-PAINT8. It offers the highest image format manipulation with Plug-ins, for most applications. Highly recommended.
We are looking for commercially available options for being able to transfer a metallographic type sample from a sample preparation workstation and into an SEM while under vacuum or inert atmosphere. We have a number of make and model SEMs therefore just any general information is fine on this subject.
thanks,
Mark A. Wall
Mr. Mark A. Wall Sr. Scientific Assoc. L-350 Chemistry & Materials Science Directorate Lawrence Livermore National Laboratory Livermore, CA USA 94550
I think you are making an excellent choice with the new Agfa DuoScan T2500. You will appreciate the 3.4 D on this unique flatbed scanner. The glass less
negative carriers fit into the lower tray, which slides into a precise position. The negatives will not contact any surface. I have just ordered also a DuoScan T2500. You will learn more about it at:
One of the most versatile imaging software is CorelDraw8 and CorelPHOTO-PAINT8. It offers the highest image format manipulation with Plug-ins, for most applications. Highly recommended.
Laszlo J. Veto
Caspar McConville wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } We are thinking of purchasing a negative scanner for use with TEM } negatives from our Jeol 2000-FX, and also for SEM negatives. A } scanner has been recommended to us: the Agfa Duoscan T2500, which } has a resolution (hardware) of 1250 x 2500 dpi and a density of 3.4D. } (The scans would be output to a Kodak DS 8650 PS printer) } } We need the quality of the scans to match the quality of the standard } darkroom enlarger if possible, as we would like to 'go digital' at } least for routine work. Does anyone have experience of routine } negative scanning for TEM prints, with this or other scanners, and if } so, is it realistic to expect such high quality? } } Also, what additional image processing software would people } recommend we got to go along with this? } } Any advice would be appreciated. } } Caspar } } Caspar McConville, Ph.D. } Technical Specialist } New York State College of Ceramics } Alfred University
I just want to say THANK YOU to all who responded to my question regarding the electrolytical thinning of Al. Now I own a collection of serveral different recipes! If anybody else is interested in it, just let me know.
Cheers,
Petra -------------------------------------------------------------- Dr. Petra Wahlbring Centre de Recherche Public Centre Universitaire (CRP-CU) Laboratoire d'Analyse des Materiaux (LAM) 162a, av. de la Faiencerie L-1511 Luxembourg tel. +352-466644-402 fax +352-466644-400 e-mail: petra.wahlbring-at-crpcu.lu Visit our WWW site! http://www.crpcu.lu/~wahlbrin
My first posting was placed whilst away from the office, now I am back I can give you the full details of the safety data mentioned.
1. Safety in the Electron Microscope Room - S. K. Chapman, Microscopy &=
Analysis, March 89, 27-29, (only two pages of text)
2. Routine Handling of Resins, RMS E.M. Safety Committee, Single sheet handout
OR from one of the same group
3. Resins: Toxicity, Hazards and Safe Handling - B. E. Causton, Proceedings RMS, Vol 16/4, June 81, 265-269
4. Routine Handling of Fixatives, RMS E.M. Safety Committee, Single shee= t handout.
I am prepared to scan in part of 1, plus 2 and 3 and send as attachments direct to those who ask.
As you can see I was not quite correct they are not all Royal Microscopic= al Society publications. Their address is 37/38 St. Clements, Oxford OX4 1AJ= , England
Steve Chapman
Senior Consultant E.M. Protrain, 16 Hedgerley, Chinnor, Oxford OX9 4TN, England. Tel & Fax 44 (0)1844 353161 Web Site - http://ourworld.compuserve.com/homepages/protrain For Consultancy and Courses in Electron Microscopy World Wide
I believe the two scanners are both excellent, although I have a preference for the Polaroid, that I am also going to buy (if can get the money).
As far as being capable of being able to use the scanner instead ot the darkroom, I strongly believe that both scanner technology, and computer technology is not yet capable of completely replacing a darkroom, unfortunately, for all its applications.
Would appreciate very much a feedback on this subject.
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
May I offer a word of caution about the approach below?
While you're correct in saying you need to optimize printing conditions for your image and printer, there will probably be a little more to it than setting up comparable number of pixels. Your image is probably either a 256-level gray scale or a 16 million color (256 levels of red, green and blue). An inkjet can't print that kind of color depth in each pixel. The concepts of "halftoning" or "dithering" need to be considered. As I'm far from the expert, and we don't need a complete textbook on the listserver anyway, so I'll refer you to chapter 2 of "The Image Processing Handbook" by John Russ. As we would expect from Dr. Russ, the material is excellent.
That does bring up a question for me, though. The new HP inkjets have a "PhotoREt" technology which, I believe is supposed to be able to vary the size of the dots it produces, therefore producing better photo-printing results. Has anyone determined whether this is true, or just hype?
Jim Passmore Sr. Analytical Chemist Cryovac Division Sealed Air Corporation
---------- } From: Harry.Ekstrom } To: Microscopy } Subject: FW: Printers for SEM Images } Date: Monday, March 08, 1999 5:11PM } ------------- } } I think an inexpensive inkjet is the way to go. However, you must try to } set the DPI of the printer to match the resolution setting of your digital } images. If you capture a digital image at 1024 X800 for instance, you have } about 820K of information. Now lets say you plan to print a 4X5 image } similar to a Polaroid, then your printer should be no less than 300dpi } {4"x5" x (300)2=1.8Mpixel} to accommodate the amount of pixel information } from the capturing rate of the image. Hence, a 2048X1600 resolution setting } captures 3.2 Mpixels of info, so a 400DPI setting should be used. The idea } being to match the capturing info with the amount of pixels the printer can } resolve to minimize interpolation...be it upwards or downwards. Not sure } what the human eye can resolve tho. } } Good Luck, } Harry Ekstrom } }
The M & M '99 Golf tournament will be Sunday, Aug. 1, 1999 just outside Portland, OR. Hole sponsorships are available for $65.00 each on a first come, first serve basis. Prizes for longest drive, longest putt etc. can also be donated. In addition, Logo gifts to be distributed to all participants will also be most welcome. To reserve your holes or to supply gifts/prizes etc. please notify me as soon as possible.
Thank you,
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.ladd.cc
We have an aging BioRad Confocal (1991!)with the associated computer and software (COMOS ver.7.0a). Has anyone replaced/upgraded their computer on this system other than just purchasing an upgrade from BioRad? What problems were encountered and what were some solutions? Thanks in advance for any info.
History of unit is below: Like an old car, different parts of the computer are beginning to need replacement, but I understand that several of the boards are specialized propietary boards (control of scan head and frame grabber) and cannot be replaced with conventional boards. We want to "upgrade" the computer, ie new mother board, more memory, etc... The existing mother board is not compatible with any device drivers other than SCSIs.
******************************************** John P. Shields Center for Ultrastructural Research 151 Barrow Hall University of Georgia Athens, GA 30602-2403 (706)542-4080 jpshield-at-arches.uga.edu ********************************************
I need to know the wavelength of the main ~ 441.6nm emission line from our He-Cd laser source to at least 5 significant figures, if possible (wavelength in air atmosphere). Any references containing other related data (emission lines) would be useful too.
-- ***************************************** Jonathan Barnard
Microstructural Physics, H.H.Wills Physics Laboratory, University of Bristol, Tyndall Avenue, Bristol BS8 1TL.
I have recently taken over an SEM with an AN 10000 analyser and I was wondering if anyone out there kows if it is possible to get the results files out to a PC ?
Any help greatly appreciated.
******************************************* Robert McDonald EPMA & SEM Laboratories Dept Geography - Earth Sciences Division Gregory Building LilyBank Gardens University of Glasgow Glasgow G12 8QQ Scotland, UK email: robert-at-earthsci.gla.ac.uk Tel:- +44 (0)141 330 5505/5442 FAX:- +44 (0)141 330 4817 ********************************************
Reply to: RE: SEM vacuum transfer = For many years I used a simple permanent magnet to lift and transfer = samples mounted on metallographic mounts with a steel washer epoxied onto = the back surface. A hole in the circular magnet allowed a push rod to = release the mount from the magnet. Possibly a small electromagnet could = be made that would be easier to operate in your system-or even a vacuum = powered suction cup to eliminate magnetic fields. Most likely you will = need to make one. = Bernard Kestel Materials Science Division Argonne National Laboratory Argonne, Il., 60439
Phone: (630) 252-4945 E-mail {kestel-at-anl.gov} Mark Wall wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America =
{HTML} {HEAD} {/HEAD} {BODY} {PRE = WIDTH=3D"132"} Reply to: RE: SEM vacuum transfer =
{/PRE} {FONT FACE=3D"Geneva" SIZE=3D3 COLOR=3D"#000000"} For = many years I used a simple permanent magnet = to lift and transfer samples mounted on = metallographic mounts with a steel washer = epoxied onto the back surface. A hole in = the circular magnet allowed a push rod to = release the mount from the magnet. Possibly = a small electromagnet could be made that = would be easier to operate in your system-or = even a vacuum powered suction cup to eliminate = magnetic fields. Most likely you will need = to make one. {BR} {BR} Bernard Kestel {BR} = Materials Science Division {BR} Argonne = National Laboratory {BR} Argonne, Il., = 60439 {BR} {BR} Phone: (630) 252-4945 = E-mail <kestel-at-anl.gov> {BR} Mark = Wall wrote: {/FONT} {FONT FACE=3D"Geneva" SIZE=3D1 COLOR=3D"#000000"} {BR} >-----------------------------------------------------------------------= - {BR} >The = Microscopy ListServer -- Sponsor: The Microscopy = Society of America {BR}
Peggy, Can you please contact me with your address, at your convenience? My computer crashed and I lost your e-mail, and snail mail addresses. I'll send the nerve staining info out when you reply. Best wishes Ronnie Houston Cytochemistry & Molecular Pathology Texas Scottish Rite Hospital for Children 2222 Welborn Street Dallas, TX 75219
I am looking for any references that can give the emission wavelength of a HeCd laser to five significant figures (in air/vacuum). The line inparticular is the 441.6 nm line, but a table of up to date measurements would be useful too.
-- ***************************************** Jonathan Barnard
Microstructural Physics, H.H.Wills Physics Laboratory, University of Bristol, Tyndall Avenue, Bristol BS8 1TL.
} We are thinking of purchasing a negative scanner...
Check out the Imacon FlexTight II. This is an affordable drum scanner with magnetic carriers for various size media (2x2 skides up to at least 8x10 sheeets) that make it as easy to use as a flatbed scanner. The unit has 5,760 dpi optical resolution (although this may drop to 4,800 dpi for something the size of an EM negative), 12-bit grayscale, 24, 32 and 48-bit color, and 14 bits (4.1 OD) of dynamic range, and it is fast. For image editing, Photoshop is the best. Period.
} That does bring up a question for me, though. The new HP } inkjets have a "PhotoREt" technology which, I believe is } supposed to be able to vary the size of the dots it produces, } therefore producing better photo-printing results. Has anyone } determined whether this is true, or just hype?
HP's statement, in itself, is explicid, so I doubt they could say it if it weren't true. I own one of these printers (720C) and looking closely it is hard to tell just how small the dots are, or when and where there being used. Appearance is very close to "random dithering". A couple of clues which would lead you to believe the technology is not just hype: (1) the suggested DPI for the printer is 300dpi which is beyond what would be calculated for a normal 600dpi dithering printer, and (2) dithering can not be seen at all for the primary colors reb, green & blue as created with cyan, magenta and yellow.
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/
A researcher would like to be able to tell the difference between apototic cells and necrotic and/or other degenerating cells in an invertebrate nervous system, using TEM. So far the only general statement I've come across is that apototic cells will undergo autophagy within their plasma membranes whereas necrotic cells tend to spill their contents and get cleaned up by other cells.
Any additional tips will be appreciated!
Aloha, Tina
http://www.pbrc.hawaii.edu/bemf/microangela **************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
I've just found a neat screen capture (and more) utility called HyperSnap-DX, a free trial version can be had from www.hyperionics.com, and registration is only $25!
cheers
rtch
} Have a couple of avi movies here and we need some stills printed from them. } Anyone have any ideas/shareware/freeware? } } } Scott D. Whittaker 218 Carr Hall } EM Technician Gainesville, FL 32610 } University Of Florida ph 352-392-1184 } ICBR EM Core Lab fax 352-846-0251 } sdw-at-biotech.ufl.edu http://www.biotech.ufl.edu/~emcl/ } The home of " Tips & Tricks " }
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
I'm not a microscopy expert, but a retired computer systems engineer.
An inkjet can produce high color if it uses CMYK inks (vs RGB). Photo-realistic inkjet printers don't dither (screen) or halftone such as with b&w printing. They attempt to produce a 1-1 relationship with the input color data. } } That does bring up a question for me, though. The new HP } inkjets have a "PhotoREt" technology which,
H-P's RET technology has been around for quite some time. I find it introduces distracting patterns into the hardcopy. H-P sat on its laurels for quite some time, and most experts say that they have fallen behind. Unless they've changed, H-Ps use RGB ink.
I find that the Epson and newer Cannon printers do a credible job of photo-color, both are CMYK.
However, you will find that the proper software will make a bigger difference in the results than the printer. If you want high color fidelity, then I recommend Adobe Photoshop, (even if you only use it for printing), calibrated for the printers ink that you use. (Epson and Cannon ink files are provided for Photoshop.) If you expect to manipulate color on the computer screen and then print the same colors on the printer, your computer has to have a "color system" such as Kodak ColorSync (adjusts for your scanner, screen and printer). Color Systems are pretty much free on the Macintosh, but you are may be out of luck if you have a Windows box (I've not found one that works very well).
Your Color System has to "know" your hardware unless you have a means of calibrating. (UMAX scanners provide a means to calibrate from a Kodak target.)
The confusion arises out of the difference between the well established printing industry and the newer technology of photorealistic inkjet printing.
In short: make sure you can calibrate your scanner for color and that it comes with a color system,
that ink files are available for your printer's ink for use by photoshop, (even if you don't use Photoshop, other programs use the PS as a standard),
I will agree wholehearted that Adobe PhotoShop is best, but I'll bet that most people never learn it properly to get their $600 worth. A very impressive PhotoShop clone is Ulead Photo Impact ( http://www.ulead.com {http://www.ulead.com} ) for ~$90.00 which does what most people use PhotoShop for (TWAIN compliant, Image resizing and level adjustments), plus Web-based image production is built in, not an add-on.) Try their 15-day fully functional demo. Enough said on image editors.
Just passing along some unbiased information.
Walt Bobrowski Subcellular Pathology Parke-Davis Research 2800 Plymouth Road Ann Arbor, MI 48105
Over the years our workforce has dwindled to the point that I find myself = working alone in the lab (metalographic sample prep/optical microscopy + = image analysis) almost all the time. The lab doesn't have windows to the = hallway, so it is difficult for people walking by to see if I am in the = lab, and if I am OK. My manager is concerned about my safety and is asking = me for suggestions. What do other labs do to make working alone safe? Everett Ramer Federal Energy Technology Center
I have replaced the original Compaq 386 with a Dell 486. I had no problems with the BioRad MRC 600 system and comos software. BioRad people have told of problems with some computers but I haven't seen any.
Larry D. Ackerman Lily & Yuh Nung Jan Laboratories Howard Hughes Medical Institute UCSF, Box 0725, Rm U226 533 Parnassus Ave. San Francisco, CA 94143
This announcement is for colleagues of Don W. Fawcett, M.D. ("A textbook = of histology", Bloom & Fawcett and "The Cell"), Professor Emeritus, = Harvard University. =
This coming Sunday (3/14/99) he will be celebrating his 82nd birthday. A = few e-mail greetings might surprize and please him. His e-mail address is = {DFawc20586-at-aol.com} . =
I am sure he would appreciate good wishes from anyone inspired by his = books and papers too.
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 --====48525552525549495651===1 Content-Type: text/html; charset="US-Ascii" Content-Transfer-Encoding: quoted-printable
{HTML} {HEAD} {/HEAD} {BODY} {FONT FACE=3D"Monaco" = SIZE=3D1 COLOR=3D"#000000"} Dear Microscopists and Histologists, {BR} {BR} This = announcement is for colleagues of Don W. = Fawcett, M.D. ("A textbook of histology", = Bloom & Fawcett and "The Cell"), = Professor Emeritus, Harvard University. = {BR} {BR} This coming Sunday (3/14/99) he will = be celebrating his 82nd birthday. A few = e-mail greetings might surprize and please = him. His e-mail address is < {/FONT} {FONT FACE=3D"Monaco" = SIZE=3D1 COLOR=3D"#0000FF"} {U} DFawc20586-at-aol.com {/U} {/FONT} {FONT FACE=3D"= Monaco" = SIZE=3D1 COLOR=3D"#000000"} >. {BR} {BR} I am sure he would = appreciate good wishes from anyone inspired = by his books and papers too. {BR} {BR} Paul Webster, = Ph.D {BR} House Ear Institute {BR} 2100 West Third = Street {BR} Los Angeles, CA 90057 {BR} phone:213 = 273 8026 {BR} fax: 213 413 6739 {BR} e-mail: {/FONT} {FONT = FACE=3D"Monaco" SIZE=3D1 COLOR=3D"#0000FF"} {U} pwebster-at-hei.org {/U} {/FONT} {= FONT = FACE=3D"Monaco" SIZE=3D1 COLOR=3D"#000000"} {BR} {/FONT} {FONT FACE=3D"Monaco" SIZE=3D1 COLOR=3D"#0000FF"} {U} http://www.hei.= org/htm/aemi.htm {/U} {/FONT} {/BODY} {/HTML} --====48525552525549495651===1 Content-Type: application/quickmail Content-Transfer-Encoding: base64
by newton.wadsworth.org (8.8.8/8.8.8) with SMTP id QAA21360 for {microscopy-at-msa.microscopy.com} ; Thu, 11 Mar 1999 16:59:34 -0500 (EST) Sender: tivol-at-wadsworth.org Message-ID: {36E839DF.41C6-at-wadsworth.org}
Jonathan Barnard wrote: Dear Johnathan,
} I need to know the wavelength of the main ~ 441.6nm emission } line from our He-Cd laser source to at least 5 significant figures, } if possible (wavelength in air atmosphere). Any references } containing other related data (emission lines) would be useful too. } Do such variables as pressure and composition (e.g., humidity) of the air allow the wavelength to be determined to 5 sig figs? The refractive index of air is related to pressure, and I think the cor- rect expression is n-1 = kP. Since P can easily vary by ~3% at sea level, depending on k, n could vary by more than 10^-5. The humidity might be even more important (especially for any other emission lines where water has n very different from that of air). Yours, Bill Tivol
We're trying to visualize meiotic spindle microtubules using LM and TEM. = Our methods for fixing and embedding insect testes are fairly = standard, however we are using 8% tannic acid in our fixative. This = appears to be what others have used, but we were wondering if anyone has = any warnings, hints or suggestions regarding microtubule preservation = and TEM.
We are also experimenting with different methods of preparing insect = testes for immunocytochemistry and LM. We are using various antibodies = to microtubules and nucleoproteins. We would like to optimize our = methods for the preservation of these structures. Is "live" tissue best = or will fixed tissue suffice? What is the best way to get the cells = spread onto slides? We've tried thumb "squashing" and cytospinning, but = are not satisfied with the results. Any suggestions would be greatly = appreciated!
Thanks,
Laura K. Garvey Dept. of Molecular and Cell Biology University of Connecticut
} } I have recently taken over an SEM with an AN 10000 analyser and I was } wondering if anyone out there kows if it is possible to get the results } files out to a PC ? } Bob-
Several years ago I wrote a nunber of utilities that would allow you to do just what you describe. You can write the files to an AN 10000 3.5" floppy (I hope yours *is* a 3.5" system? - if not, my programs won't help), which my utility will then read on a PC, copying the files to the PC in a byte-for-byte format. Then I wrote other utilities that would convert spectra and linescan files into tab-separated text files, and would extract images from studies and convert them, or individual image files, to baseline TIFF 6.0 files. I still use these utilities on a daily basis.
For a long while these were available on an FTP server here, but after aeons of no hits, and in the general progression of computer hardware, this went by the wayside. It would be the work of a few minutes to re-establish this, if there is interest. In the meantime, Bob, you may try contacting Pat Nicholson in the Dept. of Physics and Astronomy at Glasgow. I'm not sure, but I may well have given him copies of these files. In any case, I'll post the URL when I have re-established it.
If you have Windows, you should have everything you need already.
In DOS, the PrintScrn key used to dump a copy of the screen on the dot-matrix printer. In Windows nothing apparently happens. However, if you check the clipboard, PrintScrn snaps a copy of the desktop and stores it for pasting. Try pressing PrintScrn and then try paste into Word, or elsewhere. You will get a bitmap of the screen. Perhaps you don't want the whole screen - then Alt-PrintScrn copies just the active application window to the clipboard.
Using this process you end up with a lot or a little extra window junk around the sides. Then I use MS Imager that came with Office 6.0 (and was available on the MS website) to crop the image down to what I want.
I tried this using the AVI player from MS. I stopped the video, pulled the slider to the frame I wanted, and pressed Alt-PrintScrn. I opened up MS Imager, selected the File, New, Clipboard option and up came my AVI viewer window as a bitmap. I saved copies of it before and after cropping. I will send those to you directly. They are from the PICTURE.AVI movie that came on the Windows 98 disk.
You can use your imagination to apply this technique for other things, like producing your own instructions with snapshots showing what your EDS screen looks like at various stages.
Hope this Tip and Trick helps.
Warren
At 01:03 PM 3/11/99 +0000, you wrote: } } Have a couple of avi movies here and we need some stills printed from them. } Anyone have any ideas/shareware/freeware? } } } } } } {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} { } GO GATORS } Scott D. Whittaker 218 Carr Hall } EM Technician Gainesville, FL 32610 } University Of Florida ph 352-392-1184 } ICBR EM Core Lab fax 352-846-0251 } sdw-at-biotech.ufl.edu http://www.biotech.ufl.edu/~emcl/ } The home of " Tips & Tricks "
A Mac application that delivers the most commonly used tools in Photoshop (e.g. adjusting levels, assembling RGB images and montages) is Color-It!, from MicroFrontiers. It retails for about $50.
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 Thu, 11 Mar 1999, Bobrowski, Walter 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 will agree wholehearted that Adobe PhotoShop is best, but I'll bet that } most people never learn it properly to get their $600 worth. A very } impressive PhotoShop clone is Ulead Photo Impact ( http://www.ulead.com } {http://www.ulead.com} ) for ~$90.00 which does what most people use } PhotoShop for (TWAIN compliant, Image resizing and level adjustments), plus } Web-based image production is built in, not an add-on.) Try their 15-day } fully functional demo. Enough said on image editors. } } Just passing along some unbiased information. } } Walt Bobrowski } Subcellular Pathology } Parke-Davis Research } 2800 Plymouth Road } Ann Arbor, MI 48105 } } TEL: (734) 622-7814 } FAX: (734) 622-3478 } Mailto:Walter.Bobrowski-at-WL.COM {mailto:Walter.Bobrowski-at-WL.COM} } } } }
The nice thing about HyperSnap is that you can copy just any selected rectangular portion from your screen, and then either print it, fool around with it, or save it in an astonishing number of formats.
Ritchie
} } If you have Windows, you should have everything you need already. } } In DOS, the PrintScrn key used to dump a copy of the screen on the } dot-matrix printer. In Windows nothing apparently happens. However, if you } check the clipboard, PrintScrn snaps a copy of the desktop and stores it } for pasting. Try pressing PrintScrn and then try paste into Word, or } elsewhere. You will get a bitmap of the screen. Perhaps you don't want the } whole screen - then Alt-PrintScrn copies just the active application window } to the clipboard. } } Using this process you end up with a lot or a little extra window junk } around the sides. Then I use MS Imager that came with Office 6.0 (and was } available on the MS website) to crop the image down to what I want. } } I tried this using the AVI player from MS. I stopped the video, pulled the } slider to the frame I wanted, and pressed Alt-PrintScrn. I opened up MS } Imager, selected the File, New, Clipboard option and up came my AVI viewer } window as a bitmap. I saved copies of it before and after cropping. I will } send those to you directly. They are from the PICTURE.AVI movie that came } on the Windows 98 disk. } } You can use your imagination to apply this technique for other things, like } producing your own instructions with snapshots showing what your EDS screen } looks like at various stages.
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
Everett, if the door to your lab is of a "simple" configuration, it would be easy to replace it with a "windowed" door as long as your boss is willing to accept the cost as a price towards improved safety. Obviously, not a complete answer to the problem but a first step of common sense.
Mike Bucker Feed Microscopy Consolidated Labs of Va
} } } "EVERETT RAMER" {Everett.Ramer-at-fetc.doe.gov} 03/11 4:23 PM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Over the years our workforce has dwindled to the point that I find myself working alone in the lab (metalographic sample prep/optical microscopy + image analysis) almost all the time. The lab doesn't have windows to the hallway, so it is difficult for people walking by to see if I am in the lab, and if I am OK. My manager is concerned about my safety and is asking me for suggestions. What do other labs do to make working alone safe? Everett Ramer Federal Energy Technology Center
Everett and all: I work at Dow Chemical and we have a fairly rigorous "lone operator" system. If we are working in an isolated area or at a time when there are few people in the building (evenings, weekends, etc.) we carry little alert transmitters which call out to plant security (as well as within the building). The building receivers are location sensitive and we have a "lone-operator" login book at the building entrance with a building map. The lone operator marks the map as s/he signs in. Between the map and the alert location, they can find us pretty fast.
This is pretty elaborate, but maybe a mini version using something like the "First Alert" products would work ("Help, I've fallen and I can't get up") - have it tied into a siren or flashing light outside your lab so that anyone along the corridor would know there was a problem. Maybe even carry a cordless phone with an autodial button to your site security - caller ID would get them to you pretty quick.
This is definitely NOT a trivial matter. It is always disconcerting to me when I am working in an area with nobody else around - I hope you get an effective solution soon.
there's lots of stuff on t.e,m, of apoptosis in vertebrate cells (it's very popular in HIV, cancer, inflammation response etc) and much of it indicates visible nuclear changes but relative stability of cytoplasm compared with necrosis. There was a review article (as a good starting point): Microscopical Study of Cell Death via Apoptosis by S. Verhaegen in MIcroscopy and Analysis, January 1998 pp5-7
but you could do a reference or citation 'trawl' on the authors: Kerr, J.F.; Wyllie, A.H. or Currie
Malcolm Haswell Electron Microscopy School of 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: Tina Carvalho To: Microscopy Listserver
Hi, All-
A researcher would like to be able to tell the difference between apototic cells and necrotic and/or other degenerating cells in an invertebrate nervous system, using TEM. So far the only general statement I've come across is that apototic cells will undergo autophagy within their plasma membranes whereas necrotic cells tend to spill their contents and get cleaned up by other cells.
Any additional tips will be appreciated!
Aloha, Tina
http://www.pbrc.hawaii.edu/bemf/microangela **************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
by ahmler1.mail.eds.com (8.9.1/8.9.1) with ESMTP id JAA30032; Fri, 12 Mar 1999 09:51:43 -0500 (EST) Received: from SYS1.BSCO.COM (sys1.bsco.com [198.132.68.84]) by nnsa.eds.com (8.9.1/8.9.1) with SMTP id JAA27069; Fri, 12 Mar 1999 09:51:12 -0500 (EST) Received: from rfurdanowiczw [10.224.0.46] by SYS1.BSCO.COM (IBM VM SMTP Level 310) via TCP with SMTP ; Fri, 12 Mar 1999 09:51:21 EST Reply-To: {rwafu-at-bsco.com} {microscopy-at-Sparc5.Microscopy.Com}
Yes, on my AN10000 there is a program called MSDOSCV.SV which converts files between the Link (now Oxford) and PC DOS operating systems. It writes files to 720kB 3.5" floppies that previously have been formatted to that density on a PC.
An alternative that I sometimes resort to is to capture on a PC output meant to go to the Facit printer over the serial connection.
If you need more details, contact me directly or seek support from Oxford (they are very helpful), for example Ruth Murray ( ruth-at-oxford.usa.com ).
Valdemar Furdanowicz Research Labs Bethlehem Steel Co. valdemar-at-fast.net or rwafu-at-bsco.com
-----Original Message----- } From: Robert McDonald [mailto:R.McDonald-at-geology.gla.ac.uk] Sent: Thursday, March 11, 1999 10:08 AM To: microscopy-at-sparc5.microscopy.com
Hi All:
I have recently taken over an SEM with an AN 10000 analyser and I was wondering if anyone out there kows if it is possible to get the results files out to a PC ?
Any help greatly appreciated.
******************************************* Robert McDonald EPMA & SEM Laboratories Dept Geography - Earth Sciences Division Gregory Building LilyBank Gardens University of Glasgow Glasgow G12 8QQ Scotland, UK email: robert-at-earthsci.gla.ac.uk Tel:- +44 (0)141 330 5505/5442 FAX:- +44 (0)141 330 4817 ********************************************
Differentiating apoptosis and necrosis morphologically is based primari= ly upon nuclear changes, although there are characteristic cytoplasmic cha= nges as well. In general (note the wiggle words), necrotic cells swell and = lyse, whereas apoptotic cells shrink and fragment. Chromatin in apoptotic ce= lls forms electron-dense crescents at the nuclear envelope, then breaks up.=
Apoptotic cells fragment into "apoptotic bodies" that may contain bits = of chromatin. A good place to see characteristic ultrastructural changes = of apoptosis is in lymphoid tissues, where lymphocytes die via apoptosis a= nd then are phagocytosed by resident macrophages (the ones that are someti= mes called "tingible body macrophages" because of the staining properties o= f the apoptotic cell remnants in them.)
Differenting apoptosis from necrosis is a tricky deal. Apoptotic cells= may undergo secondary necrosis, during which they swell and lyse. So, just=
because you see necrotic cells doesn't mean that they didn't die apoptotically. Like everything else, it's complicated; there is a cont= inuum of change with apoptosis and necrosis at opposite poles and a lot of st= uff in between!
There are lots of good reviews on this topic. The Aug 28, 1998 issue o= f Science had a special section on apoptosis, and on page 1302, there is = a series of three electron micrographs of neurons undergoing apoptosis. = One of the first reviews of the subject contains the best collection of micrographs I've found - "Cell death: the significance of apoptosis" in= the International Review of Cytology 68:251-306, 1980. Another good revie= w was in Amer J of Pathol 146:3-15, 1995. The title is "Apoptosis, oncosis, = and necrosis: an overview of cell death."
Hope this helps!
Jane A. Fagerland, Ph.D. Dept. Microscopy and Microanalysis Abbott Laboratories Abbott Park IL 60064-6202 =
In the past I have been on jobs where a great deal of the time was spent isolated. Now days I think it is a huge safety liability. One time I had appendicitis and had to drive myself to 50 miles on back roads to a clinic with my knees up on the stearing wheel. The point here is, even though you work in a laboratory complex, if something happened you probably wouldn't get help until the cleaning crew found you. Bad news! the other issue is: Life is short and work is long, and working alone sucks. It's not emotionally healthy. Make a change. Consolidate in with other workers.
Bob Derm Imaging Center Microscopist Ex-wood cutter
On Thu, 11 Mar 1999, EVERETT RAMER 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 } -----------------------------------------------------------------------. } } } Over the years our workforce has dwindled to the point that I find myself working alone in the lab (metalographic sample prep/optical microscopy + image analysis) almost all the time. The lab doesn't have windows to the hallway, so it is difficult for people walking by to see if I am in the lab, and if I am OK. My manager is concerned about my safety and is asking me for suggestions. What do other labs do to make working alone safe? } Everett Ramer } Federal Energy Technology Center } }
The New York State Center for Advanced Thin Film Technology University at Albany - State University at New York announces the following positions:
Senior Analytical Specialist (Two Positions Available) The New York State Center for Advanced Thin Film Technology at the University at Albany - SUNY is a fast growing, high technology research and development program with a mission of supporting industry and creating new jobs. This position will serve as a primary, materials characterization, support person for the Center's scientific and technical staff in our advanced surface science facilities.
Job responsibilities include: performing day to day operation and support of the advanced surface science laboratories; assisting in data acquisition and analysis; participating in selected research projects to ensure contractual deliverables are met; providing materials characterization of microelectronic, optoelectronic and photonic samples for faculty and staff; providing instrumentation training to staff and student; and working with students on the advanced analytical tools.
The position requires: a Ph.D. in materials science or related field such as physics or chemistry and a minimum of three years experience in the characterization of microelectronic, optoelectronic or photonic materials or a bachelors degree with 10 years of relevant work experience; demonstrated ability to work in a high energy, team oriented environment; and excellent communication and analytical skills. Preference will be given to those candidates with the required work experience with Auger Electron Spectroscopy, X-Ray Photoelectron Spectroscopy, Scanning Electron Microscopy, X-Ray diffractometry, Atomic and Scanning Tunneling Microscopy, or Transmission Electron Microscopy. Salary and Benefits are highly competitive and dependent upon experience.
Please submit a resume and cover letter to:
Jacqueline DiStefano NYS Center for Advanced Thin Film Technology CESTM B110 251 Fuller Road Albany, NY 12203
The SUNY/Research Foundation is an Equal Opportunity/Affirmative Action Employer.
Well, the best method for presevering wonderfully empheral / delicate structures is cryo-preservation and freeze subsitution. I see you are a little far from us to come and try some freezing, but maybe you have some ready access to some rapid freezing.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 352 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu
I'll second the opinion about getting your money's worth out of Adobe Photoshop. I've used PhotoImpact (we have v 3 here at work; I've not used v 4 which is out already) and agree it's good. My favorite, though, is Paintshop Pro (Jasc, Inc.) which I bought for home after trying a download version. I find it is a little more intuitive than PhotoImpact (at least for me!). It handles layered images like Photoshop, and runs most Photoshop plugins. I even use it with the Image Processing Toolkit (Dr. John Russ); most of the plugins run without problem, although a few tend to crash. PhotoImpact also handles layers, I believe, but with an object-oriented approach. I haven't tried the IP Toolkit plugins in PhotoImpact.
Paintshop Pro can probably be had for a little less than Photoimpact. List price is probably $90 or $100, but I've seen it on some of the on-line computer stores for much less. I think one may have even had it for something like $58 (?). Check out info & demo at http://www.jasc.com
Disclaimer: I have no ties to any of these software packages!
Jim Passmore Cryovac Division Sealed Air Corp.
---------- } From: Walter.Bobrowski } To: Microscopy } Subject: RE: Photo Editors } Date: Thursday, March 11, 1999 3:52PM } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I think you are making an excellent choice with the new Agfa DuoScan T2500. You will appreciate the 3.4 D on this unique flatbed scanner. The glass less
negative carriers fit into the lower tray, which slides into a precise position. The negatives will not contact any surface. I have just ordered also a DuoScan T2500. You will learn more about it at:
One of the most versatile imaging software is CorelDraw8 and CorelPHOTO-PAINT8. It offers the highest image format manipulation with Plug-ins, for most applications. Highly recommended.
Laszlo J. Veto
Caspar McConville wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } We are thinking of purchasing a negative scanner for use with TEM } negatives from our Jeol 2000-FX, and also for SEM negatives. A } scanner has been recommended to us: the Agfa Duoscan T2500, which } has a resolution (hardware) of 1250 x 2500 dpi and a density of 3.4D. } (The scans would be output to a Kodak DS 8650 PS printer) } } We need the quality of the scans to match the quality of the standard } darkroom enlarger if possible, as we would like to 'go digital' at } least for routine work. Does anyone have experience of routine } negative scanning for TEM prints, with this or other scanners, and if } so, is it realistic to expect such high quality? } } Also, what additional image processing software would people } recommend we got to go along with this? } } Any advice would be appreciated. } } Caspar } } Caspar McConville, Ph.D. } Technical Specialist } New York State College of Ceramics } Alfred University
I am pseudocolorizing some data we have of cellular responses from striatum of rats learning a T-maze from Ann Graybiel's tetrode recording project. What I am looking for is any "standard" pseudocolor tables used by imagers to colorize 0-255 grey levels into RGB values.
I can apply a standard spectrum from 0 to 255 running from magenta (255 0 255 RGB) through blue, cyan, green, yellow to red (255 0 0) or start with blue and go to red in a linear way. What this produces is colors that are mostly in the middle. What I think imagers must be using are tables weighted toward the red and blue, so transitions show up better. I can play with my tables in Excel to boost the red and blue ends and then apply them using Paint Shop Pro - what I was wondering was if you have or know where I can find any standard tables, preferably of numerical RGB values used by the pros in the field.
Hi, A student in the lab is looking at museum samples of dinosaur bones and teeth on the SEM. He could get access to more samples if he could restore them to their original condition ( ie, remove the gold). Is there a good nondestructive way to do this?
Kim DeRuyter Histology and Electron Microscopy Labs University of Alaska Fairbanks
In a message dated 3/12/99 9:55:46 AM Hawaiian Standard Time, underwoo-at-u.washington.edu writes:
{ { Bad news! the other issue is: Life is short and work is long, and working alone sucks. It's not emotionally healthy. } }
Perhaps it is true for some that working alone is not emotionally healthy. However, if you ARE emotionally healthy, then working alone can be emotionally healthy and spiritually healthy also. It can be a time for focussed energy, contemplative thought, creative thought - all without interruption. Personally I love working alone and after the work I enjoy the company of my family and friends.
} From: Robert Underwood {underwoo-at-u.washington.edu}
} } In the past I have been on jobs where a great deal of the time was spent } isolated. Now days I think it is a huge safety liability. One time I had } appendicitis and had to drive myself to 50 miles on back roads to a clinic } with my knees up on the stearing wheel. The point here is, even though } you work in a laboratory complex, if something happened you probably } wouldn't get help until the cleaning crew found you. Bad news! the other } issue is: Life is short and work is long, and working alone sucks. It's } not emotionally healthy. Make a change. Consolidate in with other workers.
I spent most of my life farming and ranching. Both rather dangerous occupations. You spend almost all your time alone and in the busy season I might be alone 48 hours at a time so no one would even start looking for me for a couple of days.
Not many people are killed because of the isolation. Your best protection' is to think before you do something dangerous.
Gordon
Gordon Couger gcouger-at-couger.com Owner PRAG-L PRactical AGriculture List www.couger.com/prag-l Stillwater, OK 405 624-2855 GMT -6:00
Can anyone help me acquire an instruction manual for a Reichert-Jung FC4E cyroultramicrotome attachment for a Reichert-Jung Ultracut E ultramicrotome? I would be able to pay copy and mailing expenses.
Thanks
Damian Neuberger Research Scientist damian_neuberger-at-baxter.com
There are some two way pagers that have a man down feature. If the pager is in a horozonal postion for 10 seconds it beeps. If the wearer doesn't acknoledge the beep it sends an alarm to a central station. These pagers have a panic button as well and will serve as regular alpha numeric pagers that allow yes/no acknoledgement from the wearer.
Disclaimer: I own a substantial share of Datalink System that manufactures and selling these. See www.rfdata.net. Gordon
Gordon Couger gcouger-at-rfdata.net Datalink Systems www.rfdata.net Stillwater, OK 405 624-2855 GMT -6:00 =================================
Everett Ramer's question regarding working-alone has ellicited thoughtful and poignant reply. Useful reply as well, by Jerry Heeschen.
To level this solemn feeling, here is mine originally just shared with Everett.
Nathan Haese - at work, alone in a garage, on a new microscope, perhaps too alone.
Everett,
At a large chemical company that I once worked for, we had a badge with a radio alert button that we wore when we worked alone on weekends. Pressing the button would alert the gate guard to come find us in the lab=
Does any one have an instruction/operations manual for an American Optical microtome knife sharpener? Or if it's basic enough to enlighten me just put it in an email? Thanks in advance Steve D'Angelo
Let's join and pay our tribute to Drs Richard Henderson & Nigel Unwin for their Aminoff Award.. Cheers!
The Royal Swedish Academy of Sciences has decided to award the Gregori Aminoff prize in crystallography for 1999 to dr. Richard Henderson and dr. Nigel Unwin, MRC Laboratory of Molecular Biology, Cambridge, England, for their development of methods for structure determination of biological macromolecules using electron diffraction. The prize is presented at the Annual Meeting of the Academy 31. March 1999. The Aminoff symposium 29 - 30. March is organised to the honour of the prize-winners.
The symposium is supported by the Academy through its Nobel Institute for Chemistry.
Aminoff Symposium Structure Determination of Macromolecules with Electron Diffraction 29 - 30. March 1999
Monday 29. March
13.00 - 13.15 Opening of the symposium: Erling Norrby Introduction: Ivar Olovsson
General and non-biological Systems
13.15 - 14.15 Atomic Resolution Electron Microscopy in Biology Richard Henderson
14.15 - 15.00 Imaging Individual Atoms by Electron Microscopy Sven Hovmller, Stockholm
15.00 - 15.30 Coffee/Tea
Diffraction Studies of Membrane Proteins
15.30 - 16.30 Different methods for the study of membrane structures. Matti Saraste, EMBL, Heidelberg
16.30 - 17.15 Electron Crystallography of Membrane-bound Enzymes Hans Hebert, Stockholm
17.15 - 18.00 Structural Studies on the Cytochrome bc1 Complex So Iwata, Uppsala
18.00 - 18.30 General discussion
18.30 Dinner in the Club House of the Academy
Tuesday 30. March
Studies of Virus Structures
09.00 - 10.00 Combination of Different Methods in Virus Studies Michael Rossmann, Purdue
10.00- 10.45 Virus Studies, Essence of Supermolecular Symmetry Holland Cheng, Stockholm
10.45 - 11.15 Coffee/Tea
Non-crystalline Materials
11.15 - 12.00 Visualization of Single Protein Molecules by Electron tomography. Ulf Skoglund, Stockholm
12.00 - 13.00 0 - dimensional Crystallography Marin van Heel, Imperial College
13.00 - 14.30 Lunch in the Club House
Concluding talks
14.30 - 15.30 Making Light Work: The Membrane Proteins of Plant Photosynthesis Werner Khlbrandt, Frankfurt
Getting gold off museum fossil pieces is nay impossible. Al would be easier but basically its the same problem. C coating is good enough for low powers, but again, Curators do not like that dark coating, To view these "hard, dry and non-conducting" specimens uncoated, the best solution is a poor vacuum SEM; a fully fledged Environmental SEM would also do well, but its a more expensive instrument for that job. Poor vacuum SEM's (I believe at least a couple of the major manufacturers make instruments with that facility) use only mechanical pump vacuum in the specimen chamber and because of a vacuum limiting aperture retain high vacuum in the gun chamber. Secondary mode is impossible, but a Robinson detector gives excellent images for this type of work. Magnifications under these conditions are limited to about 2000x, but details in fossils do not warrant higher magnifications; its the SEM's superior depths of field that wins out over light microscopy. Kim - all you require now is one of those scopes! Years ago I modified an Etec Autoscan to function reversibly as a poor vacuum instruments. It worked well but it was a fair bit of trouble to accomplish the required modifications. Our online contain a link to an archive collated by Scott Wight. This contains listserver contributions concerned with Environmental and Poor Vacuum SEM. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes ********************** www.proscitech.com.au *****
On Saturday, March 13, 1999 6:55 AM, Kim DeRuyter [SMTP:fnksd1-at-uaf.edu] wrote: } } Hi, } A student in the lab is looking at museum samples of } dinosaur bones and } teeth on the SEM. He could get access to more samples if } he could restore } them to their original condition ( ie, remove the gold). } Is there a good } nondestructive way to do this? } } Kim DeRuyter } Histology and Electron Microscopy Labs } University of Alaska Fairbanks }
Due to the response (and fun) of our contest at last years MSA/MAS Conference, we will repeat the contest this year in Portland. The concept of the contest is based on composite micrographs, each made up from two or more images - one of which must be microscopical in nature. Prizes of value will be awarded and one will not have to be present to win. If of interest, kindly advise by return email and I will see that you receive full contest detail. Don Grimes, Microscopy Today
Many years ago I worked in a hospital basement EM lab. One day my boss and I came out of the inner TEM room and wondered why the hallways were deserted. Then a fire marshall came by demanding to know why we hadn't vacated the building during the fire drill!
The following email notice arrived from our university safety office: ------------
Kim
I seem to remember a story - a long time ago - about about dipping the sample in liquid mercury - the gold is taken into the liquid as an amalgam and leaves the specimen clean. I have never tried it. I do not know what any Safety person would say about that these days!
Keith Ryan Marine Biological Association of the UK Citadel Hill Plymouth Devon PL1 2PB England
In a message dated 3/12/99 9:45:34 PM Mid-Atlantic Standard Time, fnksd1-at-uaf.edu writes:
{ { Hi, A student in the lab is looking at museum samples of dinosaur bones and teeth on the SEM. He could get access to more samples if he could restore them to their original condition ( ie, remove the gold). Is there a good nondestructive way to do this?
Kim DeRuyter Histology and Electron Microscopy Labs University of Alaska Fairbanks } } Good Morning! A less destructive way to analyze these samples would be to coat them with carbon instead of gold, and then ashing the carbon off with O2. Gold is tricky to remove on most materials (I work mostly with semiconductors), but would be very difficult to remove on dinosaur bones. (This is assuming your samples are small enough to fit into an available asher or RIE tool). There are several labs that could perform both the coating and ashing - let me know if you have trouble locating one close to your area. Lisa Montanaro Consultant, MME
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im, A good way to handle specimens of this sort is to make casts of the surface and examine the casts in the SEM. This way there is no damage to the original artifact. I had an anthropology grad student do this with human teeth for his thesis research. He worked out a very inexpensive, low tech, but reliable method to do the casting. He can be reached at the following for full details of his method: Dr. Chris Schmidt Indianapolis University cschmidt-at-indy.edu
Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
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Hi, A student in the lab is looking at museum samples of dinosaur bones and teeth on the SEM. He could get access to more samples if he could restore them to their original condition ( ie, remove the gold). Is there a good nondestructive way to do this?
Kim DeRuyter Histology and Electron Microscopy Labs University of Alaska Fairbanks
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by dj.stud.ntnu.no (8.9.1/8.9.3) with ESMTP id QAA03224; Mon, 15 Mar 1999 16:01:30 +0100 (MET)
I have the same problem with the Microm-Heidelberg, Heavy duty Microtome HM 350. We would also be able to pay copy and mailing expenses.
Gary. NTNU Norway. } } } Can anyone help me acquire an instruction manual for a Reichert-Jung FC4E } cyroultramicrotome attachment for a Reichert-Jung Ultracut E } ultramicrotome? I would be able to pay copy and mailing expenses. } } Thanks } } Damian Neuberger } Research Scientist } damian_neuberger-at-baxter.com } } }
Two postdoctoral fellowships are available immediately in the Supramolecular Structure and Function group in the Bioengineering & Physical Science Program at the NIH. Our laboratory is looking for a physical scientist (e.g., physics or materials) and biological scientist (e.g., biophysics) to help develop and apply new methods based on electron microscopy and spectroscopy. There is considerable flexibility in the scope of research which includes the following:
(i) development of EELS spectrum-imaging in the STEM to map phosphorus, calcium and other elements in macromolecular assemblies and cells.
(ii) development of energy-filtered TEM to establish elemental detection limits and to map elemental distributions.
(iii) development of x-ray microanalysis and cryo-preparation techniques for studies in cell biology.
(iv) image processing techniques to determine the structures of large macromolecular assemblies using cryo-EM and STEM.
(v) applications of any of the above methods to biomedical research in collaboration with investigators in other NIH laboratories.
Our laboratory is equipped with field-emission scanning transmission electron microscopy (STEM), electron energy loss spectroscopy (EELS), energy-filtered electron microscopy (EFTEM), x-ray microanalysis (EDXS), cryo-electron microscopy, and UNIX-based image processing.
Preference will be given to candidates with less than five years of relevant postdoctoral experience. Candidates from the United States or from overseas are welcome to apply.
For additional information see: http://www.nih.gov/od/ors/beps/ssfr/
Please send curriculum vitae and bibliography to:
Dr. Richard Leapman
Biomedical Engineering & Physical Sciences Program
- Eye on Imaging - MSC/SMC Conference May 26-28, 1999
Sponsored by the Microscopical Society of Canada
We are pleased to annouce the 26th annual meeting of the Microscopical Society of Canada. This spring meeting and exhibition will be taking place for three days, May 26-28, 1999, on the campus of the University of Guelph in Guelph, Ontario. Many interesting speakers have agreed to participate including Dr. John Russ (author of the Image Processing Handbook, Dr. P.C. Cheng (multi-photon microscopy and microscope construction) , Dr. Chris Yip (AFM), Dr. Nestor Zaluzec (Tele-Presence microscopy), Dr. Nick White (3-D quantitative analysis and multi-photon microscopy) and Dr. Brian Kaye (Fractal analysis) amongst others. We are also offering a variety of afternoon workshops as well as a commercial exhibition offering a full range of Microscopy and Imaging equipment and supplies. Please visit our web site for information and registration packages:
http://www.uoguelph.ca/botany/rootlab/msc99.htm
Please pass this link along to anybody that may be interested. Deadline for submission of abstracts and pre-registration is April 6, 1999. Hope you can make it,
George Harauz Microscopical Society of Canada Chairman, Local Organizing Committee University of Guelph Guelph, Ontario gharauz-at-uoguelph.ca
You might look into utilities that allow you to adjust the color LUT and data separately of each other. Then you should be able to stretch the colors to fit like you want. If that is not easily possible with your software, then you might try playing with the gamma, contrast and brightness on your image before applying the pseudo-color and you should be able to get your weighting as you like it.
If you absolutely need help, I might be able to fabricate a color table here if you can send me a typical image and your color table.
At 03:46 PM 3/12/99 -0500, you wrote: } Hi computer imagers,, } } I am pseudocolorizing some data we have of cellular responses from striatum of } rats learning a T-maze from Ann Graybiel's tetrode recording project. What I } am looking for is any "standard" pseudocolor tables used by imagers to colorize } 0-255 grey levels into RGB values. } } I can apply a standard spectrum from 0 to 255 } running from magenta (255 0 255 RGB) through blue, cyan, green, yellow to red } (255 0 0) or start with blue and go to red in a linear way. What this produces } is colors that are mostly in the middle. What I think imagers must be using } are tables weighted toward the red and blue, so transitions show up better. I } can play with my tables in Excel to boost the red and blue ends and then apply } them using Paint Shop Pro - what I was wondering was if you have or know where } I can find any standard tables, preferably of numerical RGB values used by the } pros in the field. } } } ------------------------------------------------------------------ } |Glenn Holm {mailto:karuzis-at-wccf.mit.edu} | } |Graybiel Lab (617)253-5780;fax (617)253-1599 | } |M.I.T Dept. of Brain + Cog. Sci. Cambridge, MA 02139 | } ------------------------------------------------------------------
Kim DeRuyter 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, } A student in the lab is looking at museum samples of dinosaur bones and } teeth on the SEM. He could get access to more samples if he could restore } them to their original condition ( ie, remove the gold). Is there a good } nondestructive way to do this? } } Kim DeRuyter } Histology and Electron Microscopy Labs } University of Alaska Fairbanks Kim, One thing that no one has mentioned, yet, is low kV operation. If your instrument will operate at 5kV or lower (perferably around 1kV) and you limit your beam current, you should be able to view uncoated specimens at at least a couple of kX. Some intruments will go much higher at that voltage range. Grains of quartz may still present a problem because SiO2 is such a good insulator, but many other minerals will work fine under those conditions.
Ken Converse owner Quality Images third party SEM service Delta, PA
I have set up an FTP site at prism.mit.edu, port 2101, with the files I mentioned that will convert LINK/Oxford AN10/eX/L files. Apologies if the documentation is sparse!
To access a non-standard port in FTP you will need to know how your FTP program works. In Netscape, use the following URL:
ftp://prism.mit.edu:2101
In WS_FTP you have to change the port number in the Advanced tab. From a command line ftp program (e.g. Unix, or DOS from Win 95/98/NT), first invoke the program without a server name, i.e. just enter
ftp
The program responds with the ftp prompt ftp} . Then you enter:
open prism.mit.edu 2101
The user is anonymous, and the password is unimportant. I don't know how you would do it with other FTP programs. Sorry about the difficulty, but I am already using the standard FTP port for another, non-public, purpose!
I think you are making an excellent choice with the new Agfa DuoScan T2500. You will appreciate the 3.4 D on this unique flatbed scanner. The glass less
negative carriers fit into the lower tray, which slides into a precise position. The negatives will not contact any surface. I have just ordered also a DuoScan T2500. You will learn more about it at:
One of the most versatile imaging software is CorelDraw8 and CorelPHOTO-PAINT8. It offers the highest image format manipulation with Plug-ins, for most applications. Highly recommended.
Laszlo J. Veto
Caspar McConville wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } We are thinking of purchasing a negative scanner for use with TEM } negatives from our Jeol 2000-FX, and also for SEM negatives. A } scanner has been recommended to us: the Agfa Duoscan T2500, which } has a resolution (hardware) of 1250 x 2500 dpi and a density of 3.4D. } (The scans would be output to a Kodak DS 8650 PS printer) } } We need the quality of the scans to match the quality of the standard } darkroom enlarger if possible, as we would like to 'go digital' at } least for routine work. Does anyone have experience of routine } negative scanning for TEM prints, with this or other scanners, and if } so, is it realistic to expect such high quality? } } Also, what additional image processing software would people } recommend we got to go along with this? } } Any advice would be appreciated. } } Caspar } } Caspar McConville, Ph.D. } Technical Specialist } New York State College of Ceramics } Alfred University
} From: Self {miller.TEX.TAFA} To: Microscopy-at-MSA.Microscopy.Com
Howdy all, There is a graduate student here who is trying to look at a possible stem cell line under TEM. The problem is that they are quite small, don't seem to pellet very well, and he has only been able to get me a few thousand cells at a time. We have tried to embed the cells in 2% Agar after fixation but this hasn't worked. Is it possible to filter the media and cells, and then process the filter with the cells stuck onto it? Maybe use a cytospin to spin the cells into a filter? Is there anyone with experience with this? Any suggestions or ideas would be greatly appreciated.
Thanks in advance Frank Herbert Technician Integrated Microscopy Core Department of Cell Biology Baylor College of Medicine
Somewhere in the deep receses of my mind I recall a cytochemical test for arsenic. I think it was for EM but I am not sure. Does anyone out there know it??
Or was it all just a bad dream?
Greg 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
We are trying to equip our lab with a reflected light BF/DF microscope (+ trinocular head, B&W video system, 4X5" Polaroid system), for two basic needs: 1. viewing and photographing our fine metal, ceramic, and carbide powders for QA purposes (e.g. visual standards), and 2. viewing and photographing mounted/polished thermal spray powder and coating samples prepared at our parent company's facility.
Our budget allows for a used (reconditioned or demo) top-name microscope (e.g. Nikon, Olympus, Leitz, Zeiss) or a new lesser- name scope (e.g. Meiji...) with the same basic features. Not being professional microscopists ourselves, we ask you to comment on aspects of the above tradeoff, based on your experience. Our main concern is flatness of field and sharpness of image at magnifications up to 500X. Thanks for your help!
Sincerely,
Robert A. Miller TAFA Material Technologies, Inc. 1702 Mykawa Road Pearland TX 77581 USA PHONE: 281-485-7765 FAX: 281-485-0211 EMAIL: miller-at-tafa.com
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Hi, I'm trying to find out if the elemental imaging system on my scope (a Zeiss 902A TEM) is working...or if it is a problem with the sample. Does anyone have a sample they could spare for elemental imaging?
In October, I had a great sample grid (30nm liver sections with no stain). We were able to get nice images of iron. The scope had its annual service in November and the elemental imaging system has not worked correctly since that time. The serviceman is suggesting that the sample is fried (no pun intended). I think the scope is whacked. So I'm trying to get another liver sample (my source for the first sample retired) or I'm willing to try something else. I need a "known" sample, cut 30 nm thick, no support film (600 mesh grids help) or staining.
Any help or suggestions would be GREATLY appreciated!
Sincerely, Beth Richardson
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
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I'm interested. Elaine
Dr. Elaine Humphrey Biosciences Electron Microscopy Facility University of British Columbia 6270 University Blvd, mail-stop Botany Vancouver, BC CANADA, V6T 1Z4 Phone: 604-822-3354 FAX: 604-822-6089 e-mail: ech-at-unixg.ubc.ca
hello members. here is a probable dumb question. Maybe an impossible situation.
Suppose that I have a prepared microscope slide-- 1"x3" glass slide with specimen under cover slip. Ordinary LM analysis works fine. Is there some other analysis method besides confocal that would offer better resolution and increased depth of field? The idea is to not have to prepare TEM specimens.
Is this possible? Gary Gaugler, Ph.D. ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Modern surfers use PC boards. You can too at http://photoweb.net ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~