I'm a student now considering studies in basics of microscopy to consider a possible career change.
I'm seeking information on flourescence microscopy, why would someone use this over other methods, the difference between single photon and two photon fluorescence . Anyone's experience on laser scanning confocal microscopes ( 1photon and 2 photon), is it worth the expense? Although I don't know what they cost, I am told that they are expensive, does anyone know?
Thank You Paul Bitetto
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 06:42:33 2004
Silvia, There are indeed flat, enclosable embedding cylindrical molds, standard size #00, with a flat end....on both sides! Just like BEEM capsules. I get ours from Electron Microscopy Sciences (#70021):
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From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 08:33:47 2004
I don't know a good way to remove the hairs, but for counting and measuring stomata, it would be easier if any waxy cuticle was removed during processing. I have done this by using acetone as a dehydrant, rather than ethanol. In my experience, it cleans up the surface of the leaf quite well and makes the stomata and other surface features stand out nicely.
Good luck.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
-----Original Message----- } From: by way of Ask-A-Microscopist [mailto:kn77-at-uwyo.edu] Sent: Tuesday, August 31, 2004 6:34 PM To: microscopy-at-microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kn77-at-uwyo.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Tuesday, August 31, 2004 at 14:43:11 ------------------------------------------------------------------------ ---
Email: kn77-at-uwyo.edu Name: Kusum Naithani
Organization: University of Wyoming
Education: Graduate College
Location: Laramie, Wyoming, USA
Question: Hi! I'm working on plant leaf material (sagebrush). Leaf is covered by minute white hairs and due to this reason I'm not able to find the distribution of stomatal cells. Could you please suggest a way to remove these hairs so that I can see stomatal cells. 2nd question.. I want to measure the size and depth of stomatal cells. Could you please tell me the way to fix leaf in its living conditions. I would greatly appreciate your help. Thanks! Kusum
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mingram-at-rohmhaas.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 1, 2004 at 05:24:13 ---------------------------------------------------------------------------
Email: mingram-at-rohmhaas.com Name: Mike Ingram
Organization: Rohm and Haas
Title-Subject: [Microscopy] [Filtered] MListserver: Registration of SEM's due to X-rays
Question: In Delaware it appears we are required to register our SEM as a X-ray source. Does any know this to be true?
You can try removing the trichomes by very gently shaving the leaf. Hold a razor blade almost vertical, just not touching the leaf surface, and pull in the direction of the tilt. This requires a steady hand, and doesn't work on robust trichomes, but might on sagebrush -- I don't know that plant. The other approach would be to go at the leaf with the blade near horizontal and again just not touching the leaf, like shaving your face. But I haven't done that in so long, I've forgotten how. You don't need to remove the entire trichome, just most of it, so the surface is revealed. The best way to measure the size of the stomatal cells would be with a light microscope on freshly picked leafs in a room (or chamber) of the appropriate humidity. Or, same conditions, but make a replica with dental silicone or one of the replica materials the EM companies sell. The replica could then be examined in the SEM, or with a light microscope.
Phil
} Email: kn77-at-uwyo.edu } Name: Kusum Naithani } } Organization: University of Wyoming } } Education: Graduate College } } Location: Laramie, Wyoming, USA } } Question: Hi! } I'm working on plant leaf material (sagebrush). Leaf is covered by } minute white hairs and due to this reason I'm not able to find the } distribution of stomatal cells. Could you please suggest a way to } remove these hairs so that I can see stomatal cells. } 2nd question.. } I want to measure the size and depth of stomatal cells. Could you } please tell me the way to fix leaf in its living conditions. } I would greatly appreciate your help. } Thanks! } Kusum } } ---------------------------------------------------------------------------
-- Philip Oshel Supervisor, BBPIC microscopy facility Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 - 1284 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 11:16:27 2004
Hi Paul, Fluorescence microscopy when you have access to confocal microscope is awesome. For career, every experimentals fields, particulary with bio tag on the name is not an easy career from money perspective, but from scientific point of view it is great.
All the best,
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } Hello All } } I'm a student now considering studies in basics of microscopy to consider a } possible career change. } } I'm seeking information on flourescence microscopy, why would someone use } this over other methods, the difference between single photon and two photon } fluorescence . Anyone's experience on laser scanning confocal microscopes ( } 1photon and 2 photon), is it worth the expense? Although I don't know what } they cost, I am told that they are expensive, does anyone know? } } Thank You } Paul Bitetto } } } } } } } }
-- Antoine Blanc, Research Associate Ecole Polytechnique de Montreal Chemical Engineering, Mike Buschmann Laboratory CP 6079, succ. Centre-ville Montréal Qc, Canada H3C 3A7 Tel.:514-340-4711 ext.:3212,3336,3337 FAX:514-340-4159 secrétariat: 514-340-4711 ext.:4984
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 11:44:29 2004
I'm not sure about Delaware, but this is true for the State of Michigan. You may want to check your State's website in the registration or health section for information.
In the last conversation I had with the state inspector, the State is considering dropping registration for scanning electron microscopes because of the negligible danger from SEMs leaking X-rays. Since the SEM by all practical purposes cannot operate unless everything is buttoned up for the vacuum, the chance for X-ray leakage is just about nil.
Stu Smalinskas, P.E. SKF USA Plymouth, Michigan
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Mike wrote:
Question: In Delaware it appears we are required to register our SEM as a X-ray source. Does any know this to be true?
Thanks
Email: mingram-at-rohmhaas.com Name: Mike Ingram Organization: Rohm and Haas
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From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 12:27:45 2004
O.k., here is my suggestion for removing the root hairs: Follow any standard fixation protocol (I know you asked about them as well) but you'll find that sagebrush does not "wet" very well, so you'll need to add a surfactant like Kodak Photo-flo or Tween (1 drop / 10ml fixative is generally enough) - if you are looking to follow up with light microscopy then FAA might work very nicely for you [FAA = Formalin-Acedic acid-Alcohol Formulation: 50% (or 70%) ethyl alcohol - 90ml, Glacial Acetic Acid - 5ml, Formalin - 5ml]. In which the alcohol works well to "wet" the material.
In any case, dehydrate the samples to 50%-100% solvent (EtOH), plunge freeze in liquid nitrogen, rub the surface of the leaves with a wood stick, pre- cooled razor blade, plastic bar, etc. This should break all the tricomes off the leaf surface revealing the stomata. Transfer samples back from liquid nitrogen and continue with sample prep.
To accurately measure somata depth you will have to section the samples and look at the cells in cross-section.
Standard fixatation
1) Primary Fixation: 1-2% paraformaldehyde, 2-4% glutaraldehyde, in a suitable pH buffer (i.e. 6.8-7.2 pH 0.2 M Sodium Phosphate, 0.05 M Sodium Cacodylate, HEPES, PIPES, etc. ). Fixation for 5-120 min. at room temp. (20- 22 C), normal growth temp (37 C?) or on ice (0-4 C). [50 min. -at- room temp]. Specimens should be cut as small as possible.
2) Rinse: 4 times -at- 10-15 min. each with the above buffer without aldehyde fixatives. (Residual aldehydes will bind with OsO4 in secondary fixation if used.)
*3) Secondary Fixation: 1-2% Osmium tetroxide (OsO4) in full to half strength buffer used above. Fixation for 2-6 hours at room temperature. (OsO4 fixation generally not used if immunological staining procedures will be used.)
*4) Rinse: 4 times -at- 15-20 min. each with distilled water.
5) Dehydration: Generally either absolute ethanol (200 proof) or glass distilled acetone is used.
% Solvent in water Time
25% 20-30 min 50% 20-30 min 75% 20-30 min 95% 30-60 min 100% 60+ min 100% 60+ min
SEM: CPD samples
TEM: resin embbedd samples
} } Email: kn77-at-uwyo.edu } Name: Kusum Naithani } } Organization: University of Wyoming } } Education: Graduate College } } Location: Laramie, Wyoming, USA } } Question: Hi! } I'm working on plant leaf material (sagebrush). Leaf is covered by minute white } hairs and due to this reason I'm not able to find the distribution of stomatal } cells. Could you please suggest a way to remove these hairs so that I can see } stomatal cells. 2nd question.. I want to measure the size and depth of stomatal } cells. Could you please tell me the way to fix leaf in its living conditions. I } would greatly appreciate your help. Thanks! Kusum } } --------------------------------------------------------------------------- }
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Director 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 12:57:24 2004
This is also true for Illinois. We used to have routine inspections by the IL Dept of Nuclear Safety, but they felt it was no longer justified for SEMs.
Alan Stone ASTON
At 12:06 PM 9/1/2004, you wrote:
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Alan Stone ASTON Metallurgical Services Co., Inc. 200 Larkin Drive Ste A Wheeling, IL 60090 847/353-8100 www.astonmet.com
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 14:02:18 2004
Mike, As of July 2000 it was still required. Try contacting
Delaware Health and Social Services Division of Public Health Office of Radiation Control P.O. Box 637 Dover, DE 19903 302-739-3787
Perhaps 4 years later you might be able to find a website for them. From Stu's reply it sounds like some sanity might be returning concerning SEMs and x-rays, but I'm not going to go there today.
Good luck, Ken Converse
owner QUALITY IMAGES Servicing Scanning Electron Microscopes Since 1981 16 Creek Rd. Delta, PA 17314 717-456-5491 Fax 717-456-7996 kenconverse-at-qualityimages.biz qualityimages.biz
-----Original Message----- } From: by way of MicroscopyListserver [mailto:mingram-at-rohmhaas.com] Sent: Wednesday, September 01, 2004 10:21 AM To: microscopy-at-microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mingram-at-rohmhaas.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 1, 2004 at 05:24:13 ---------------------------------------------------------------------------
Email: mingram-at-rohmhaas.com Name: Mike Ingram
Organization: Rohm and Haas
Title-Subject: [Microscopy] [Filtered] MListserver: Registration of SEM's due to X-rays
Question: In Delaware it appears we are required to register our SEM as a X-ray source. Does any know this to be true?
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From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 14:55:47 2004
Just some thoughts on your breaking grids: 1. Make sure the Pioloform is absolutely fresh. 2. Can you use a regular mesh grid for additional support? 3. You could obtain carbon coated slot grids from Ladd or some other EM supplier.
John Arnott
Disclaimer: Ladd Research sells grids, custom coated grids, and the supplies needed to make them yourself.
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: ladres-at-att.net
----- Original Message ----- } From: "Bill Tivol" {tivol-at-caltech.edu} To: {microscopy-at-msa.microscopy.com} Sent: Tuesday, August 31, 2004 12:57 PM
The fall meeting of the Texas Society for Microscopy will be Oct. 21-23 at the Hilton Garden Inn in Allen, TX. We have an exciting schedule planned including the Thursday workshop "EBSD and FESEM - A Formidable Combination for Characterization of Semiconductor Materials" to be presented by Dr. Keith Dicks from Oxford Instruments. The workshop will take place at Microtech Analytical Labs, Inc., 538 Haggard Street, Suite 402 Plano, TX 75074. In addition to the Thursday workshop, Oxford Instruments, JEOL and Microtech Analytical Labs are making the Inca Energy EDX and Inca Crystal EBSD system on a JEOL 6500F available for demo Tues, Wed and Fri (Oct. 19-20, 22). This is an opportunity to bring your own samples and evaluate this technique with one on one contact with the industry experts. To schedule demo time, contact Mike Crowley with Oxford Instruments at 512-246-7551 or by email at crowley-at-ma.oxinst.com. Space is limited so sign up early.
All registration forms and hotel information are available on our web site: http://www.texasmicroscopy.org/. We look forward to seeing you in the fall.
Regards, Jodi Roepsch Program Chair 972-952-3228, j-roepsch1-at-raytheon.com
-- ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Becky Holdford (r-holdford-at-ti.com) 972-995-2360 972-648-8743 (pager) SC Packaging FA Development Texas Instruments, Inc. Dallas, TX ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 17:00:06 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (nbauer-at-paulstra.com) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Wednesday, September 1, 2004 at 10:11:08 ---------------------------------------------------------------------------
Email: nbauer-at-paulstra.com Name: Nathan Bauer
Organization: Paulstra
Education: Graduate College
Location: Grand Rapids, Mi, USA
Question: Can carbon nano tubes : 1) Carry an electrical current (and if so, how much)?
This is not strictly true, Michigan are thinking of stopping the registration if the instrument is "unmodified" from the manufacturer. If you have added anything that was not supplied by the manufacturer, i.e an XEDS system, CL system, viewport or similar then they still want it registered and tested. We just had our health and safety people check all our stuff because the Michigan inspector came round and checked all of our machines (TEMs, SEMs FIBs and XPS).
-- John Mansfield PhD MInstP North Campus Electron Microbeam Analysis Laboratory 417 SRB, University of Michigan 2455 Hayward, Ann Arbor MI 48109-2143 USA Phone: (734) 936-3352 FAX (734) 763-2282 Cell. Phone: (734) 834-3913 (Leaving a phone message at 936-3352 is preferable to 834-3913) Email: jfmjfm-at-engin.umich.edu URL: http://emalwww.engin.umich.edu/people/jfmjfm/jfmjfm.html Location: Lat. 42° 16' 48" Long. 83° 43' 48" AIM: thejfmjfm
Home address: 4304 Spring Lake Boulevard Ann Arbor MI 48108-9657 Phone (734) 994-3096
On Sep 1, 2004, at 1:06 PM, Kestutis Smalinskas wrote:
} } } ----------------------------------------------------------------------- } ------- } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ----------------------------------------------------------------------- } -------- } } I'm not sure about Delaware, but this is true for the } State of Michigan. You may want to check your State's } website in the registration or health section for } information. } } In the last conversation I had with the state } inspector, the State is considering dropping } registration for scanning electron microscopes because } of the negligible danger from SEMs leaking X-rays. } Since the SEM by all practical purposes cannot operate } unless everything is buttoned up for the vacuum, the } chance for X-ray leakage is just about nil. } } Stu Smalinskas, P.E. } SKF USA } Plymouth, Michigan } } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ } } Mike wrote: } } Question: In Delaware it appears we are required to } register our SEM as a X-ray source. Does any know } this to be true? } } Thanks } } Email: mingram-at-rohmhaas.com } Name: Mike Ingram } Organization: Rohm and Haas } } } } } } __________________________________ } Do you Yahoo!? } Yahoo! Mail is new and improved - Check it out! } http://promotions.yahoo.com/new_mail }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 1 20:52:22 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (lotocka-at-acn.waw.pl) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 1, 2004 at 14:49:49 ---------------------------------------------------------------------------
Email: lotocka-at-acn.waw.pl Name: Barbara Lotocka
Organization: Department of Botany, Warsaw Agricultural University
Title-Subject: [Microscopy] MListserver: fixative for lichen
Question: Hello Everyone,
I would be most grateful for any suggestions on a fixation protocol (for transmission electron microscope) optimized for lichens.
I fixed some samples of Cladonia in a fixative that is routinely used for plant samples in my department (paraformaldehyde + glutaraldehyde in sodium cacodylate buffer), but after embedding in epoxy resin the thallus looked shrunken and the section were "scratched" as if the thallus was extremely hard. Perhaps the problem was in dehydration? I used the usual graded series of ethanol and acetone.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (pzou-at-feico.com) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 1, 2004 at 20:19:52 ---------------------------------------------------------------------------
Question: Can anybody comment on the image darkening effect as one progressively scans the surface of a sample with a SEM? What are the possible physical causes, and how to reduce the effect?
Any articles that can provide an overview of this phenomenon?
One possible way to remove hairs that should work is to make a surface replica of the surface using nail varnish or a mounting medium such as Shur Mount and stripping off when dry. On most of the leaf tissues I've worked with it removes hairs, fungi, surface debris but does not damage the surface itself.
Ian
Ian Hallett HortResearch Mt Albert Research Centre, Private Bag 92 169 Auckland, New Zealand Fax +64 9 815 4201 Telephone +64 9 815 4200 EMail ihallett-at-hortresearch.co.nz
} } } by way of Ask-A-Microscopist {kn77-at-uwyo.edu} 1/09/2004 11:34:02 a.m. } } }
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kn77-at-uwyo.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Tuesday, August 31, 2004 at 14:43:11 ---------------------------------------------------------------------------
Email: kn77-at-uwyo.edu Name: Kusum Naithani
Organization: University of Wyoming
Education: Graduate College
Location: Laramie, Wyoming, USA
Question: Hi! I'm working on plant leaf material (sagebrush). Leaf is covered by minute white hairs and due to this reason I'm not able to find the distribution of stomatal cells. Could you please suggest a way to remove these hairs so that I can see stomatal cells. 2nd question.. I want to measure the size and depth of stomatal cells. Could you please tell me the way to fix leaf in its living conditions. I would greatly appreciate your help. Thanks! Kusum
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From MicroscopyL-request-at-ns.microscopy.com Thu Sep 2 01:26:55 2004
Gareth Morgan MPhil MSc FIBMS, Department of Laboratory Medicine (Labmed), Karolinska Institute, Karolinska University Hospital at Huddinge, F46 SE 141 86 Stockholm Sweden
OBS! Besöksadress: F-Huset, Forskningsgatan 2 F52, Rum 2.10. Laboratoriet för klinisk patologi och cytologi.
NB! Visiting address: Building F, Research Corridor 2 F52, Room 2.10. Clinical Histo- and Cytopathology Laboratory.
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 2 01:54:31 2004
See section 9.10.6 in Goldstein et al.: Scanning electron microscopy and X-ray analysis" Plenum Press 1992. It begins: "A sample subjected to electron bombardment in a diffusion-pumped vacuum gradually becomes ccovered with a contamination layer due to polymerization, under the action of the beam, of organic matter adsorbed on the surface".
Ways to reduce the effect are: clean vacuum, clean sample, cold finger.
-----Original Message----- } From: by way of MicroscopyListserver [mailto:pzou-at-feico.com] Sent: 2. september 2004 04:16 To: microscopy-at-microscopy.com
--------------------------- Dr Wally H. Müller Senior University Research University of Utrecht, Faculty of Biology Molecular Cell Biology - Electron Microscopy Kruyt building, Room West 510 Padualaan 8, 3584 CH Utrecht, The Netherlands Phone +31 30 2533588 Fax +31 30 2513655 E-mail W.H.Muller-at-bio.uu.nl
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 2 05:16:35 2004
In a message dated 9/2/04 3:45:16 AM, Gareth.Morgan-at-labmed.ki.se writes:
} Anyone out there know of any courses in microscopic morphometry/digital } image analysis - preferably in Europe.
Upcoming courses on quantitative image analysis that I will be teaching are:
Sunday October 3 - A one-day tutorial on strategies for quantitative image analysis will be presented as part of the AOCS Conference on Food Structure and Quality in Cork, Ireland. Registration for the workshop is available at {http://www.aocs.org/meetings/fsq/courses.asp}
Tuesday, November 9 - Thursday, November 11 - A three-day hands-on course on Quantitative Image Analysis will be presented at the University of Missouri, Columbia, MO. Contact {rosslm-at-missouri.edu} Dr. Lou Ross, Electron Microscopy Core Facility, W136 Veterinary Medicine, University of Missouri, Columbia, MO 65211-5120, (573) 882-4777, fax 884-5414.
Wednesday, March 16 - Friday, March 18, 2005 - A three-day hands-on course on Photomicrography and Advanced Image Analysis will be presented at the McCrone Institute in Chicago. Contact {rweaver-at-mcri.org} Dr. Rob Weaver at the McCrone Institute, 2820 South Michigan Avenue, Chicago IL 60616, 312-842-7100. A brief description of the course contents is available at {http://www.mcri.org/Course_description.html#advdig} their website
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 2 08:17:25 2004
For a simple and effective method for making plant surface replicas for observation by LM or SEM, see my article in the Nov/Dec 2003 issue of Microscopy Today: Cellulose Acetate Replication of Plant Surfaces for SEM (plug plug!!). You'll see images of stomata there.
http://www.microscopy-today.com
However, if a plant surface is very thickly populated by a tangled mess of hairs, or trichomes, then replication may not be possible, as it will be full of holes left by the trichomes and may tear apart upon attempted removal from the surface. If the hair density is not too high, even though holes from trichomes may be present, you may still be able to see enough surface to get a good sample of the stomata.
-- Gib Ahlstrand, Scientist Electron Optical Facility, University of Minnesota, CBS Imaging Center, 35 Snyder Hall, St. Paul, MN. USA. 55108 (612)624-3454 (612)624-2785 FAX, ahlst007-at-tc.umn.edu http://www.cbs.umn.edu/ic/
"You can learn a lot by observation - just by lookin'!" - Yogi Berra
} Kusum } } One possible way to remove hairs that should work is to make a surface replica } of the surface using nail varnish or a mounting medium such as Shur Mount and } stripping off when dry. On most of the leaf tissues I've worked with it } removes hairs, fungi, surface debris but does not damage the surface itself. } } Ian } } } Ian Hallett } HortResearch } Mt Albert Research Centre, Private Bag 92 169 } Auckland, New Zealand } Fax +64 9 815 4201 } Telephone +64 9 815 4200 } EMail ihallett-at-hortresearch.co.nz } } } Below is the result of your feedback form (NJZFM-ultra-55). It was submitted } by (kn77-at-uwyo.edu) from } http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on } Tuesday, August 31, 2004 at 14:43:11 } --------------------------------------------------------------------------- } } Email: kn77-at-uwyo.edu } Name: Kusum Naithani } } Organization: University of Wyoming } } Education: Graduate College } } Location: Laramie, Wyoming, USA } } Question: Hi! } I'm working on plant leaf material (sagebrush). Leaf is covered by minute } white hairs and due to this reason I'm not able to find the distribution of } stomatal cells. Could you please suggest a way to remove these hairs so that I } can see stomatal cells. } 2nd question.. } I want to measure the size and depth of stomatal cells. Could you please tell } me the way to fix leaf in its living conditions. } I would greatly appreciate your help. } Thanks! } Kusum }
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 2 12:45:37 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jtd1-at-psu.edu) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, September 2, 2004 at 10:55:11 ---------------------------------------------------------------------------
Question: Our lab is considering various photo printers (} $1K) for production of electron micrographs. Currently the Epson 2200 is the stromg favorite. Are there any recomendations for other printers which we should consider? What are your reasons for the recomended printer?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (walter.bobrowski-at-pfizer.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, September 2, 2004 at 11:12:16 ---------------------------------------------------------------------------
Email: walter.bobrowski-at-pfizer.com Name: Walt Bobrowski
Organization: Pfizer Global R&D
Title-Subject: [Microscopy] [Filtered] DMP-30 vs. BDMA
Question: Can one substitute BDMA for DMP-30 in a Luft-based epoxy resin? If so, what would the proportion be? Currently, I add 2ml DMP-30 to 100 ml resin (PolyBed 812, NMA, DDSA). I believe I read you can substitute to produce a less viscous mixture, but can't find it. Any references appreciated!
} Email: walter.bobrowski-at-pfizer.com } Name: Walt Bobrowski } } Organization: Pfizer Global R&D } } Title-Subject: [Microscopy] [Filtered] DMP-30 vs. BDMA } } Question: Can one substitute BDMA for DMP-30 in a Luft-based epoxy } resin? If so, what would the proportion be? Currently, I add 2ml } DMP-30 to 100 ml resin (PolyBed 812, NMA, DDSA). I believe I read } you can substitute to produce a less viscous mixture, but can't find } it. Any references appreciated! } } Walt -
The only reason that DMP-30 is still around is tradition; by all means substitute the same amount of BDMA. Viscosities: BDMA, 0.85 cP, DMP-30, 20.5 cP. The quantity that you use is small, so there won't be a big change in the viscosity of the mix, but DMP-30 is so viscous that it can actually partition out of the mix during infiltration! AND DMP-30 is hygroscopic, which leads to more problems. The original reference is A. Glauert, Proc. RMS 22:264 (1987) and you'll find the data in chapter 6 of Glauert & Lewis, Biological Specimen Preparation for Transmission Electron Microscopy, Princeton,1998.
Why not use a less viscous epoxy, such as Spurr (with the new, safer ERL 4221) or Embed-It? They're both 65 cP, mixed.
-- Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/ Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 2 14:50:47 2004
So after years of hearing and reading about why BDMA is better than DMP-30, I switched to BDMA I used it at about 60-80% of the weight of DMP-30 and got equivalent results using freshly made resin mixes. But I routinely store my extra resin at -20 C and saw a difference. The BDMA mixtures got much more viscous (presumably partially polymerized) after 1-2 weeks at -20 compared to the DMP-30 mixtures. So I went back to DMP-30. Whichever one you use, I strongly advised you begin to dispense it and the other components by weight. I have a scale in my fume hood and make 50 ml batches of epoxy resins this way and they are much more consistent and the mess is significantly less. good luck. tom phillips
01:08 PM 09/02/04 -0500, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
I had a look at the archive and this had been discussed a few times in the last few years. Technology is changing fast especially in the digital world. We two would like to know the best printer (quality) out there if there is no money limitation and the best value for money meaning quality prints all editors of journals will be happy with. We got a comment like "the digital images are brilliant but the prints do not do them justice" from a editor. Please pass all communication to us as well. Thanks for all the help.
-----Original Message----- } From: by way of MicroscopyListserver [mailto:jtd1-at-psu.edu] Sent: Thursday, September 02, 2004 8:08 PM To: microscopy-at-microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jtd1-at-psu.edu) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, September 2, 2004 at 10:55:11 ---------------------------------------------------------------------------
Question: Our lab is considering various photo printers (} $1K) for production of electron micrographs. Currently the Epson 2200 is the stromg favorite. Are there any recomendations for other printers which we should consider? What are your reasons for the recomended printer?
In the same week there is news of significant change in two of the companies that were major players in silver image photography in the 20th century. Ilford has shed almost half its staff preparative to sale of the traditional photographic business while it is still a going concern, allowing the company to focus on its Swiss digital business. http://www.channel4.com/news/news_story.jsp?storyId=156722 Agfa has shed its traditional photographic film and consumer imaging business in a management buyout so that it can "focus on its core growth markets of Graphic Systems and HealthCare, which are rapidly going digital" http://news.agfa.com/corporate/news.nsf/news/F07C0210ECC86EA9C1256EF3004D27CE?opendocument
Events like these, and Kodak's announcement earlier this year that it would cease the production of its poneering APS cameras (though not of films) underline the fragility of the conventional photographic market in the face of the growth of digital imaging.
Which begs the question "can we rely on the continued availability of EM film", and if not, how long have we got to plan for the conversion to digital?
Dr. Chris Jeffree University of Edinburgh Schoolof Biological Sciences
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 05:34:31 2004
These units are designed to print only photo-quality and they do it well. As we have progressed towards keeping documents (including images) electronic and have thrown out the darkroom, and due to cost per print ($4.00/8x10), we only use the Pictrography to print images for manuscripts. If colleagues want review-quality, they either must review on their computer monitor or send image files to a B&W/Color LaserJet (yeah, lousy quality, but it's only for review).
Best regards,
Walter F. Bobrowski Investigative Pathology Safety Sciences Pfizer Global Research & Development Ann Arbor, MI 48105
-----Original Message----- } From: Coetzee, Mr S. H Physics Science [mailto:COETZEES-at-mopipi.ub.bw] Sent: Friday, September 03, 2004 1:51 AM To: by way of MicroscopyListserver Cc: microscopy-at-microscopy.com
Dear Tom
I had a look at the archive and this had been discussed a few times in the last few years. Technology is changing fast especially in the digital world. We two would like to know the best printer (quality) out there if there is no money limitation and the best value for money meaning quality prints all editors of journals will be happy with. We got a comment like "the digital images are brilliant but the prints do not do them justice" from a editor. Please pass all communication to us as well. Thanks for all the help.
-----Original Message----- } From: by way of MicroscopyListserver [mailto:jtd1-at-psu.edu] Sent: Thursday, September 02, 2004 8:08 PM To: microscopy-at-microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jtd1-at-psu.edu) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, September 2, 2004 at 10:55:11 ---------------------------------------------------------------------------
Question: Our lab is considering various photo printers (} $1K) for production of electron micrographs. Currently the Epson 2200 is the stromg favorite. Are there any recomendations for other printers which we should consider? What are your reasons for the recomended printer?
LEGAL NOTICE Unless expressly stated otherwise, this message is confidential and may be privileged. It is intended for the addressee(s) only. Access to this E-mail by anyone else is unauthorized. If you are not an addressee, any disclosure or copying of the contents of this E-mail or any action taken (or not taken) in reliance on it is unauthorized and may be unlawful. If you are not an addressee, please inform the sender immediately.
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 05:45:11 2004
On the Vibration testing, the kit is available from various companies but is expensive. We bought a complete unit from spicer consulting http://www.spicerconsulting.com/ Their SC11 kit costs around $10,000 but that does everything.
However there are other products available. ( http://www.predictech.com/CM/PT908.htm ) not sure though that they will be sensitive enough.
P.O. Box 2561 Honeydew 2040 Gauteng, South Africa -----Original Message----- } From: White, Woody N. [mailto:nwwhite-at-bwxt.com] Sent: 31 August 2004 11:02 To: 'edelmare-at-MUOhio.edu'; MicroscopyListServer
An inexpensive possibility...
The accelerometer may need a signal conditioner/buffer/amplifier - I have not checked it's specifications. If a PC sound card meets your frequency range and resolution requirements, the rest is simple and cheap.
A number of free/shareware programs that will turn your PC into an audio range spectrum analyzer are available. Adjust the accelerometer/amplifier output level to be compatible with the sound card "line in" requirement (typically 1 volt peak, max)and you are there...
Regards, Woody
Woody White BWXT Services: http://www.bwxt.com/bwxt.html My Site: http://woody.white.home.att.net
-----Original Message----- } From: Richard Edelmann [mailto:edelmare-at-MUOhio.edu] Sent: Tuesday, August 31, 2004 8:34 AM To: microscopy-at-MSA.Microscopy.com
With a number of on-going building modifications here as well as the
potential for relocating my EM Facility. we find ourselves in need of a vibration testing/monitoring system. What I am hoping to find is a simply system to plug into a Laptop. Having had various vibration testing done in the past, and where as I fully acknowledge the quality experience of testing service providers we just can't afford that kind of expense on a continuing basis. We are looking to do this in-house, monitor "baseline" building vibrations every couple of weeks or so.
I have picked up the recommendations for a Wilcoxon 731A/P31 Accelerometer but now I need a compatible PCMCIA spectrum analyzer interface board and software.
Any recommendations? And yes, vendors may respond directly back to me.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Director 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 07:36:56 2004
Group, In my view, simple inkjet printers do an extraordinary job. You can get one for 200 us dollars (or more or less depending on features). What is important is that the quality of the output is determined by the paper you put in (and some software toggles in the printer driver). So, there is a range of paper from high quality photo paper to zerox paper, with the cost per print scaling appropriately (and the time per print likewise). This makes it easy to get low, intermediate, high quality output on the same printer, with little fuss. And I have to say that for monochrome prints, the highest quality settings/paper produce prints equal to anything I have seen from the wetlab type of printers mentioned below. And even for color the differences are pretty small. Another issue is networking. The inexpensive inkjet that I have is not network-able (that's an intersting word) and I don't know if the more expensive ones give you that option.
My two pixels, Tobias
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I have already starting campaigning for a digital camera for our TEM!
I saw an advert in Microscopy and Analysis for a company which is (presumably) starting making TEM film as others leave the market.
Kodak did say recently on this listserver that they have no plans to drop EM film.
Dave
On Fri, 03 Sep 2004 09:05:45 +0100 Chris Jeffree {c.jeffree-at-ed.ac.uk} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } In the same week there is news of significant change in two of the } companies that were } major players in silver image photography in the 20th century. } Ilford has shed almost half its staff preparative to sale of the } traditional photographic } business while it is still a going concern, allowing the company to } focus on its Swiss digital business. } http://www.channel4.com/news/news_story.jsp?storyId=156722 } Agfa has shed its traditional photographic film and consumer imaging } business in a management buyout so that it can "focus on its core } growth markets of Graphic Systems and HealthCare, which are rapidly } going digital" } http://news.agfa.com/corporate/news.nsf/news/F07C0210ECC86EA9C1256EF3004D27CE?opendocument } } Events like these, and Kodak's announcement earlier this year that it } would cease the production of its poneering APS cameras (though not of } films) underline the fragility of the conventional photographic } market in the face of the growth of digital imaging. } } Which begs the question "can we rely on the continued availability of } EM film", and if not, how long have we got } to plan for the conversion to digital? } } } Dr. Chris Jeffree } University of Edinburgh } Schoolof Biological Sciences } } } } } This incoming email to UWE has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
This email has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 09:21:02 2004
Kodak has included a positive statement on its continued commitment to film products in its contribution to the "New and Interesting at M&M 2004" section of the September issue of Microscopy Today, at the printer now and in your mail boxes starting in a week or so.
Ron Anderson, MT Editor
-----Original Message----- } From: Chris Jeffree [mailto:c.jeffree-at-ed.ac.uk] Sent: Friday, September 03, 2004 4:06 AM To: microscopy-at-msa.microscopy.com
In the same week there is news of significant change in two of the companies that were major players in silver image photography in the 20th century. Ilford has shed almost half its staff preparative to sale of the traditional photographic business while it is still a going concern, allowing the company to focus on its Swiss digital business. http://www.channel4.com/news/news_story.jsp?storyId=156722 Agfa has shed its traditional photographic film and consumer imaging business in a management buyout so that it can "focus on its core growth markets of Graphic Systems and HealthCare, which are rapidly going digital" http://news.agfa.com/corporate/news.nsf/news/F07C0210ECC86EA9C1256EF3004D27C E?opendocument
Events like these, and Kodak's announcement earlier this year that it would cease the production of its poneering APS cameras (though not of films) underline the fragility of the conventional photographic market in the face of the growth of digital imaging.
Which begs the question "can we rely on the continued availability of EM film", and if not, how long have we got to plan for the conversion to digital?
Dr. Chris Jeffree University of Edinburgh Schoolof Biological Sciences
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 10:40:02 2004
I recently learned that Serco-Ilford is no more and I need service for my 2150RC. Does anyone know of anyone serving the New England area? Thanks in advance.
Mary
Mary McKee Program in Membrane Biology MGH-Charlestown (617)726-3696 --
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 11:18:33 2004
I freeze with DMP-30 in the tube. Generally I aliquot them so they are only thawed once. They are certainly good for a couple of weeks this way and longer in many cases. I make all my resin by weight (typically 20 g Embed 812, 10 g DDSA, 10 g NMA, and 0.6 g DMP-30) in a 50 ml plastic disposable tube and shake vigorously until well mixed. We used 0.8 g BDMA in place of DMP-30 in this formulation and saw no difference in cutting quality but did have the storage problem. The viscosity of the DDSA and NMA and Embed 812 is high and requires vigorous shaking regardless of the catylst so I don't see lower viscosity of BDMA as significant if you are measuring by weight. If you measure by volume, a viscous solution will be tougher to accurately measure and deliver and the percent error will be much higher for the small volume BDMA or DMP-30 component. I have never ever seen the DMP-30 come out of solution such as Caroline Schooley suggests in her e-mail; if this happens I would suspect improper mixing. My use of DMP-30 is based on careful consideration and comparison with BDMA and not on tradition. I used BDMA exclusively for over 1 year and then switched back so I think I gave it a fair shot.
At 10:22 AM 09/03/04 -0600, you wrote: } I have a question about freezing the resin mixtures - you freeze them with } the accelerator already added? How many freeze/thaw cycles can they stand? } Or, do you aliquot them to store and thaw only once?? I was taught to } store mine without DMP-30....if they'll last OK with it in, I'm all for it! } } Thanks, } } Tamara } } On Thu, 2 Sep 2004, Tom Phillips wrote: } } } } } } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
For a less viscous mixture, you could try one percent DMP-30 instead of two percent. The final cure might not be quite so hard, though, so depending on your tissue this may or may not help.
Lesley Weston.
----- Original Message ----- } From: walter.bobrowski-at-pfizer.com (by way of MicroscopyListserver) Sent: 9/2/2004 11:08:46 AM To: microscopy-at-microscopy.com
} - } } I freeze with DMP-30 in the tube. Generally I aliquot them so they } are only thawed once. They are certainly good for a couple of weeks } this way and longer in many cases. I make all my resin by weight } (typically 20 g Embed 812, 10 g DDSA, 10 g NMA, and 0.6 g DMP-30) in } a 50 ml plastic disposable tube and shake vigorously until well } mixed. We used 0.8 g BDMA in place of DMP-30 in this formulation } and saw no difference in cutting quality but did have the storage } problem. The viscosity of the DDSA and NMA and Embed 812 is high } and requires vigorous shaking regardless of the catylst so I don't } see lower viscosity of BDMA as significant if you are measuring by } weight. If you measure by volume, a viscous solution will be tougher } to accurately measure and deliver and the percent error will be much } higher for the small volume BDMA or DMP-30 component. I have never } ever seen the DMP-30 come out of solution such as Caroline Schooley } suggests in her e-mail; if this happens I would suspect improper } mixing. My use of DMP-30 is based on careful consideration and } comparison with BDMA and not on tradition. I used BDMA exclusively } for over 1 year and then switched back so I think I gave it a fair } shot.
You missed my point, Tom; when I said that the DMP-30 can partition during infiltration, I meant that it doesn't enter the tissue as rapidly as the other resin components. The symptom is soft tissue in a normal, hard block. I agree with you that the high DMP-30 viscosity is going to have little effect on the viscosity of the mixed epoxy.
Caroline -- Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/ Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 15:21:20 2004
Do you have evidence that the DMP-30 doesn't get in or is that your hypothesis to explain the problem? How do you know it is a sign that the DMP-30 didn't get in and not the complete mixture or one of the equally viscous components? I find it hard to believe the DMP-30 differentially inflitrates. In addition, the DMP-30 (and BDMA) has probably already begun to react with the other components and would be "carried" in covalently bound to the one of them. Poor infiltration of the entire mix would (and does) result in the symptom you describe. Take a resin mix made with BDMA and do a short infiltration on a tough to infiltrate tissue like maize endosperm and the center will be softer than the perimeter and free resin regions. BDMA is frequently touted as superior due to its lower viscosity and less hydrophilic. It is lower viscosity but I don't see that as a problem or benefit. I have no data on the relative hydroscopic properties but I have never had a bottle of DMP-30 go bad on me in the 25 years I have been doing TEM. Maybe I use my bottles up before they goes bad. We do close it up promptly after using but I assume most labs do. I have nothing against BDMA if one is making the resin up fresh each time (which is the best practice regardless of whether you use BDMA or DMP-30). But for our routine samples, we commonly make 40-50 gm batches and store the resin at -20 C for 1-3 weeks. This works when we use DMP-30 but not with BDMA. I agree that lots of what microscopists do is because of "tradition" but the selection of DMP-30 can be the result of a careful, reasoned and experimentally tested decision process. Tom Phillips
At 01:21 PM 09/03/04 -0700, you wrote: } } - } } } } I freeze with DMP-30 in the tube. Generally I aliquot them so they are } } only thawed once. They are certainly good for a couple of weeks this way } } and longer in many cases. I make all my resin by weight (typically 20 g } } Embed 812, 10 g DDSA, 10 g NMA, and 0.6 g DMP-30) in a 50 ml plastic } } disposable tube and shake vigorously until well mixed. We used 0.8 g } } BDMA in place of DMP-30 in this formulation and saw no difference in } } cutting quality but did have the storage problem. The viscosity of the } } DDSA and NMA and Embed 812 is high and requires vigorous shaking } } regardless of the catylst so I don't see lower viscosity of BDMA as } } significant if you are measuring by weight. If you measure by volume, a } } viscous solution will be tougher to accurately measure and deliver and } } the percent error will be much higher for the small volume BDMA or DMP-30 } } component. I have never ever seen the DMP-30 come out of solution such } } as Caroline Schooley suggests in her e-mail; if this happens I would } } suspect improper mixing. My use of DMP-30 is based on careful } } consideration and comparison with BDMA and not on tradition. I used BDMA } } exclusively for over 1 year and then switched back so I think I gave it a } } fair shot. } } You missed my point, Tom; when I said that the DMP-30 can partition during } infiltration, I meant that it doesn't enter the tissue as rapidly as the } other resin components. The symptom is soft tissue in a normal, hard } block. I agree with you that the high DMP-30 viscosity is going to have } little effect on the viscosity of the mixed epoxy. } } Caroline } -- } Caroline Schooley } Project MICRO Coordinator } Microscopy Society of America } Box 117, 45301 Caspar Point Road } Caspar, CA 95420 } Phone/FAX (707)964-9460 } Project MICRO: http://www.msa.microscopy.com/ProjectMicro/ } Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
I'm looking for sources of acoustic/anachoic blocks of foam to reduce SEM room interference.
These are like standard anechoic chamber panels. I'm having trouble getting above 400KX without mechanical noise from the scroll pump that is located in a separate room. The acoustical isolation from one room to the other is not all that great, I suppose.
Does anyone have experience with suppliers of these panels? I'd like to glue them to drywall.
Supplier responses are welcomed as off-line messages.
gary g.
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 3 21:05:34 2004
You can try http://www.illbruck-sonex.com/ for what you need. I'm familiar with this product when we used it at the High Voltage EM lab at UW Madison. Worked well.
Damian Neuberger
-----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] Sent: Friday, September 03, 2004 8:31 PM To: MSA listserver
I'm looking for sources of acoustic/anachoic blocks of foam to reduce SEM room interference.
These are like standard anechoic chamber panels. I'm having trouble getting above 400KX without mechanical noise from the scroll pump that is located in a separate room. The acoustical isolation from one room to the other is not all that great, I suppose.
Does anyone have experience with suppliers of these panels? I'd like to glue them to drywall.
Supplier responses are welcomed as off-line messages.
gary g.
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 4 07:05:23 2004
We use a Polaroid SprintScan 45 multi-format film scanner to scan our EM negatives. It has been a work horse for us for 7 years, and the hardware is still in a good shape. However, We lost the original disk with driver (for Mac) on it and we could not get support from the company anymore because Polaroid has discontinued scanner business. Does anyone out there have the same scanner and would be willing to lend us the driver software? Our machine is currently down because the computer could not locate it even though the scanner was on and all cables were connected. We need to re-install the driver as the first trouble shooting step.
If we can not find a driver, we might have to purchase a new film scanner. Does anyone has a recommendation on makes and models?
Thank you very much in advance.
Hong Emory EM hyi-at-emory.edu
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 4 11:39:26 2004
Bill Robertson in his reply posting pointed out a very good point about wall mass. My initial thoughts were to put the acoustic tiles in the SEM room. But the noise is more likely coming from the adjacent room where the scroll pump and specimen interchange pump are located. The specimen interchange pump is not at issue. The scroll pump is an Edwards XDS 10. Based on limited experience, it does not sound like it is running as it should. Vacuum is fine but the sound is odd. It makes a mechanical knocking noise. I'm told that either these pumps work almost forever when new or die quickly. Sounds like I have the latter.
In either case, wall mass has a high probability of attention. The intervening wall is standard drywall with 2x4 studs. There is no insulation in the wall. I'm thinking of having liquid foam insulation put in the wall cells and seeing if that helps. In the mean time, the plinth anti-vibration system is not exactly right and will get adjusted shortly. then, based on how that turns out, acoustic tiles in the pump room may be a good solution. If that does not help much, then foam insulation.
The chiller is in the same room as the SEM but turning it off makes no change in image noise. So the noise is external.
Thanks for the replies. Will work on this in the next couple of weeks.
gary g.
At 08:38 AM 9/4/2004, you wrote: } Gary, } } They can be pricey. One reason is that most jurisdictions require that } insulation } applied to an open wall surface be flame proof. Not only that, the } adhesive used } to apply it must also be flame proof, per the fire department and building } codes. } (Recall the disasterous fire in the Rhode Island night club a couple years } back). } Some years ago I found that the material required behind an XRD system in } a small } room ran several hundred dollars for a half dozen panels. However, I } found that I } could get a big improvement by placing a small number of them so as to } block the } noise at the source, rather than covering the more distant wall } surfaces. Buy a } small number to try out, or experiment with packing foam. Have an } assistant hold } them in different locations so as to block acoustic reflections. You may } find that } suspending one from the ceiling in the plane of the instrument helps alot with } noise at the position where the operator sits. } } John Twilley } } Gary Gaugler wrote: } } } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } ------------------------------------------------------------------------------- } } } } I'm looking for sources of acoustic/anachoic } } blocks of foam to reduce SEM room interference. } } } } These are like standard anechoic chamber panels. } } I'm having trouble getting above 400KX without } } mechanical noise from the scroll pump that is } } located in a separate room. The acoustical } } isolation from one room to the other is not } } all that great, I suppose. } } } } Does anyone have experience with suppliers of } } these panels? I'd like to glue them to drywall. } } } } Supplier responses are welcomed as off-line } } messages. } } } } gary g.
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 4 11:44:55 2004
I was a fan of Epson photo printers for quite some time. Notable was the 890 & 980. It is a small format printer compared to the 2200. I recently (a year or so ago) bought a Epson Stylus Photo 2000. It lasted about six months and then jammed constantly. In-warranty customer service and any idea of repair was on a wish list. It never happened. The printer was scrapped.
The 2200 may have solved teething problems with large format printers. However, the 2000 was VERY slooooow using photo paper. When it worked, the results were stunning. Many times (too many) it would stop printing 1/4 or 1/2 way through the print and just die. The job hung (Win2K Pro) and had to be restarted with a new sheet of paper.
The Epson and Canon small format printers seem to do a better, more reliable job. As a result of being burned by Epson, I now take print jobs to a local service bureau. they do a very nice job for not much cost. These are mostly for 24" x 48" glossy mounted prints. Small ones are done on my HP 4550 color laser printer. If the color gamut is matched well between the monitor and Photoshop, the HP does a nice job for reports. For transparencies (not much used any longer), the Kodak dye sub is excellent.
Let us know what you find. There are a lot of options. Also, check out the Ethernet print servers that will connect a non-network printer to a LAN and allow all to use it. HP and others make these. they usually cost about $100 or so.
gary g.
At 11:08 AM 9/2/2004, you wrote:
} Email: jtd1-at-psu.edu } Name: Tom Doman } } Organization: Penn State University } } Title-Subject: [Microscopy] [Filtered] MListserver: } } Question: Our lab is considering various photo printers (} $1K) for } production of electron micrographs. Currently the Epson 2200 is the stromg } favorite. Are there any recomendations for other printers which we should } consider? What are your reasons for the recomended printer? } } Thanks in advance! } } Tom } } ---------------------------------------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 4 13:33:46 2004
From your description of the sound coming from your pump, it sounds like your issue might be low frequency. For low frequencies, acoustic tiles and the like are not very effective. The strategy for reducing high frequencies is to absorb the energy, and usually involves foam or fibruous materials that will vibrate and dissipate the energy. For low frequencies, the strategy is to block/reflect the energy, and this requires rigidity and mass. The best is a very solid wall, but there are also a number of lead-backed sheet materials, either separately or in combination with absorbers. If you have access to a McMaster Carr catalog, I suggest you check out "Sound Absorbers" (or visit their web site at www.mcmaster.com and do a keyboard search for "sound". There is also a helpful summary on page 3266 of their online catalog which explains the various types and ratings systems. (No financial interest in McMaster Carr, but use them all the time.) Fred Schamber ASPEX, LLC
Gary Gaugler wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } } Bill Robertson in his reply posting pointed out } a very good point about wall mass. My initial } thoughts were to put the acoustic tiles in the SEM room. } But the noise is more likely coming from the adjacent } room where the scroll pump and specimen interchange } pump are located. The specimen interchange pump } is not at issue. The scroll pump is an Edwards XDS 10. } Based on limited experience, it does not sound like } it is running as it should. Vacuum is fine but the } sound is odd. It makes a mechanical knocking noise. } I'm told that either these pumps work almost forever } when new or die quickly. Sounds like I have the latter. } } In either case, wall mass has a high probability } of attention. The intervening wall is standard } drywall with 2x4 studs. There is no insulation } in the wall. I'm thinking of having liquid foam } insulation put in the wall cells and seeing if that } helps. In the mean time, the plinth anti-vibration } system is not exactly right and will get adjusted } shortly. then, based on how that turns out, acoustic } tiles in the pump room may be a good solution. If that } does not help much, then foam insulation. } } The chiller is in the same room as the SEM but turning } it off makes no change in image noise. So the noise } is external. } } Thanks for the replies. Will work on this in the } next couple of weeks. } } gary g. } } } At 08:38 AM 9/4/2004, you wrote: } } } Gary, } } } } They can be pricey. One reason is that most jurisdictions require } } that insulation } } applied to an open wall surface be flame proof. Not only that, the } } adhesive used } } to apply it must also be flame proof, per the fire department and } } building codes. } } (Recall the disasterous fire in the Rhode Island night club a couple } } years back). } } Some years ago I found that the material required behind an XRD } } system in a small } } room ran several hundred dollars for a half dozen panels. However, I } } found that I } } could get a big improvement by placing a small number of them so as } } to block the } } noise at the source, rather than covering the more distant wall } } surfaces. Buy a } } small number to try out, or experiment with packing foam. Have an } } assistant hold } } them in different locations so as to block acoustic reflections. You } } may find that } } suspending one from the ceiling in the plane of the instrument helps } } alot with } } noise at the position where the operator sits. } } } } John Twilley } } } } Gary Gaugler wrote: } } } } } } } ------------------------------------------------------------------------------ } } } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } } America } } } To Subscribe/Unsubscribe -- } } http://www.msa.microscopy.com/MicroscopyListserver } } } On-Line Help } } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } } } ------------------------------------------------------------------------------- } } } } } } } } I'm looking for sources of acoustic/anachoic } } } blocks of foam to reduce SEM room interference. } } } } } } These are like standard anechoic chamber panels. } } } I'm having trouble getting above 400KX without } } } mechanical noise from the scroll pump that is } } } located in a separate room. The acoustical } } } isolation from one room to the other is not } } } all that great, I suppose. } } } } } } Does anyone have experience with suppliers of } } } these panels? I'd like to glue them to drywall. } } } } } } Supplier responses are welcomed as off-line } } } messages. } } } } } } gary g. } } } }
From MicroscopyL-request-at-ns.microscopy.com Sun Sep 5 11:32:17 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jbarclay-at-southpointresources.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, September 5, 2004 at 08:40:08 ---------------------------------------------------------------------------
Email: jbarclay-at-southpointresources.com Name: Jim Barclay
Organization: Calgary, Alberta
Title-Subject: [Microscopy] [Filtered] Wild Heerbrug M5A microscope & camera attachment
Question: I am trying to see if I can find some type of adaptor that will fit my older Wild Heerbrug M5A binocular microscope (15-20 years old?) and allow me to fit a digital camera to the scope. I would need some type of beam splitter that would continue to allow simultaneosu viewing of samples while also allowing occasional photo taking.
I have already contacted and am waiting for reply from a Leica Microsystems representative which owns the Wild Leitz brand.
Thank you for listening to a newbie to this board. Any suggestions welcome.
} I'm looking for sources of acoustic/anachoic } blocks of foam to reduce SEM room interference. } } These are like standard anechoic chamber panels. } I'm having trouble getting above 400KX without } mechanical noise from the scroll pump that is } located in a separate room. The acoustical } isolation from one room to the other is not } all that great, I suppose. } } Does anyone have experience with suppliers of } these panels? I'd like to glue them to drywall. } } Supplier responses are welcomed as off-line } messages. } Dear Gary, Fred has it right about the difference between low and high frequencies. The depth of the invaginations in anechoic panels has to be 1/4 of the wavelength of the sound in order to be effective, so for low frequencies, the panels could take up the entire room, leaving no space for the equipment. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Sun Sep 5 18:43:56 2004
I don't have any direct way of knowing whether the noise is high or low frequency. It only shows up above 60KX as sinusoids on the edges of specimen details.
I can probably use FFT to compute the frequency. It is consistent.
gary g.
At 03:34 PM 9/5/2004, you wrote:
} On Sep 3, 2004, at 6:30 PM, Gary Gaugler wrote: } } } I'm looking for sources of acoustic/anachoic } } blocks of foam to reduce SEM room interference. } } } } These are like standard anechoic chamber panels. } } I'm having trouble getting above 400KX without } } mechanical noise from the scroll pump that is } } located in a separate room. The acoustical } } isolation from one room to the other is not } } all that great, I suppose. } } } } Does anyone have experience with suppliers of } } these panels? I'd like to glue them to drywall. } } } } Supplier responses are welcomed as off-line } } messages. } Dear Gary, } Fred has it right about the difference between low and high } frequencies. The depth of the invaginations in anechoic panels has to be } 1/4 of the wavelength of the sound in order to be effective, so for low } frequencies, the panels could take up the entire room, leaving no space } for the equipment. } Yours, } Bill Tivol, PhD } EM Scientist and Manager } Cryo-Electron Microscopy Facility } Broad Center, Mail Code 114-96 } California Institute of Technology } Pasadena CA 91125 } (626) 395-8833 } tivol-at-caltech.edu } }
From MicroscopyL-request-at-ns.microscopy.com Sun Sep 5 23:13:35 2004
If you are building a wall that may transmuted vibration making the wall a 6 inch wall with double 4 inch studs that are every 8 inches with every other stud supporting opposite faces of the wall. If you use a sound deadening blanket widen the wall to leave room to weave the padding between the studs with out compressing it.
Two layers of 5/8 inch gypsum wall board are generaly considered fire proof enough for university buildings at Oklahoma State. The last I knew there were no paints that could be used on anything but aluminum to increase their fire rating. But that has been a while. In actual practice there are paints that improve the fire resistance of anything but last I knew it changed so much with age that for anything but metal it was too unpredictable to approve.
Gordon Gordon Couger gcc-at-couger.com
I collect links on information related to light microscopes. http://www.couger.com/microscope/links/gclinks.html Please forward any links or information you think might be useful to others. Microscope Manual at www.science-info.org
} From: "Gary Gaugler" {gary-at-gaugler.com} : : Bill Robertson in his reply posting pointed out : a very good point about wall mass. My initial : thoughts were to put the acoustic tiles in the SEM room. : But the noise is more likely coming from the adjacent : room where the scroll pump and specimen interchange : pump are located. The specimen interchange pump : is not at issue. The scroll pump is an Edwards XDS 10. : Based on limited experience, it does not sound like : it is running as it should. Vacuum is fine but the : sound is odd. It makes a mechanical knocking noise. : I'm told that either these pumps work almost forever : when new or die quickly. Sounds like I have the latter. : : In either case, wall mass has a high probability : of attention. The intervening wall is standard : drywall with 2x4 studs. There is no insulation : in the wall. I'm thinking of having liquid foam : insulation put in the wall cells and seeing if that : helps. In the mean time, the plinth anti-vibration : system is not exactly right and will get adjusted : shortly. then, based on how that turns out, acoustic : tiles in the pump room may be a good solution. If that : does not help much, then foam insulation. : : The chiller is in the same room as the SEM but turning : it off makes no change in image noise. So the noise : is external. : : Thanks for the replies. Will work on this in the : next couple of weeks. : : gary g. : : : At 08:38 AM 9/4/2004, you wrote: : } Gary, : } : } They can be pricey. One reason is that most jurisdictions require that : } insulation : } applied to an open wall surface be flame proof. Not only that, the : } adhesive used : } to apply it must also be flame proof, per the fire department and building : } codes. : } (Recall the disasterous fire in the Rhode Island night club a couple years : } back). : } Some years ago I found that the material required behind an XRD system in : } a small : } room ran several hundred dollars for a half dozen panels. However, I : } found that I : } could get a big improvement by placing a small number of them so as to : } block the : } noise at the source, rather than covering the more distant wall : } surfaces. Buy a : } small number to try out, or experiment with packing foam. Have an : } assistant hold : } them in different locations so as to block acoustic reflections. You may : } find that : } suspending one from the ceiling in the plane of the instrument helps alot with : } noise at the position where the operator sits. : } : } John Twilley : } : } Gary Gaugler wrote: : } : } } : } ------------------------------------------------------------------ ------------ : } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America : } } To Subscribe/Unsubscribe -- : } http://www.msa.microscopy.com/MicroscopyListserver : } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html : } } : } ------------------------------------------------------------------ ------------- : } } : } } I'm looking for sources of acoustic/anachoic : } } blocks of foam to reduce SEM room interference. : } } : } } These are like standard anechoic chamber panels. : } } I'm having trouble getting above 400KX without : } } mechanical noise from the scroll pump that is : } } located in a separate room. The acoustical : } } isolation from one room to the other is not : } } all that great, I suppose. : } } : } } Does anyone have experience with suppliers of : } } these panels? I'd like to glue them to drywall. : } } : } } Supplier responses are welcomed as off-line : } } messages. : } } : } } gary g. : : :
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 01:14:19 2004
not wanting to sound paranoid, i think we have to consider chris's words carefully.
while digital systems are good, after 35 years i cannot get the same results from a digital image than i can from film, no matter how hard i try. double exposures with polychromatic paper using different filters, dodging, burning, combined, can contribute to excellent prints. perhaps others can get the same results from digitized images, after 5 years, i cannot.
we currently maintain two older Philips 201 and one cm10 microscopes. they have 35mm cameras. Kodak has discontinued manufacture of Direct Positive 5302, which we use for these instruments. the only sources of which i now know for this film is the different suppliers. but how long will their stocks last? what other 35mm format films are there that have the same high resolution that is found with 5302?
all in all, the promise from Kodak is fine and dandy, but will that hold when Kodak decides that the digital market has made it no longer viable to support wet chemistry with specialized, high resolution films such as we require.
of course, as far as i am concerned, i have 25 roles of 5302 in the freezer, so i'm set until our microscopes die. but it is an ongoing concern for the rest of you who were not ordering at the time that Kodak made their decision and were not able to stock up.
paul
Paul R. Hazelton, PhD Electron Microscope Unit University of Manitoba Department of Medical Microbiology 531 Basic Medical Sciences Building 730 William Avenue Winnipeg, Manitoba, Canada, R3E 0W3 e-mail: paul_hazelton-at-umanitoba.ca Phone:204-789-3313 Pager:204-931-954 Cell:204-781-1502 Fax:204-789-3926
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 01:57:23 2004
we produce c-mount adapters which can be connected to the eyepiece of a microscope. The eyepiece remains in place, you put our eyepiece adapter on top of it. Our eyepiece adapter has a lens built-in. This has the advantage of capturing most of what you see through the eyepiece. You can easily remove the adapter and look through the eyepiece again. We offer different sizes of eyepiece adapters. We also manufacture on demand.
If you would like to get more information please contact me.
bwoM} ------------------------------------------------------------------------------ bwoM} The Microscopy ListServer -- Sponsor: The Microscopy Society of America bwoM} To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver bwoM} On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html bwoM} -------------------------------------------------------------------------------
bwoM} Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jbarclay-at-southpointresources.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on bwoM} Sunday, September 5, 2004 at 08:40:08 bwoM} ---------------------------------------------------------------------------
bwoM} Email: jbarclay-at-southpointresources.com bwoM} Name: Jim Barclay
bwoM} Question: I am trying to see if I can find some type of adaptor that will fit my older Wild Heerbrug M5A binocular microscope (15-20 years old?) and allow me to fit a digital camera to the bwoM} scope. I would need some type of beam splitter that would continue to allow simultaneosu viewing of samples while also allowing occasional photo taking.
bwoM} I have already contacted and am waiting for reply from a Leica Microsystems representative which owns the Wild Leitz brand.
bwoM} Thank you for listening to a newbie to this board. Any suggestions welcome.
Chris Jeffree wrote the following: ====================================================================== In the same week there is news of significant change in two of the companies that were major players in silver image photography in the 20th century. Ilford has shed almost half its staff preparative to sale of the traditional photographic business while it is still a going concern, allowing the company to focus on its Swiss digital business. http://www.channel4.com/news/news_story.jsp?storyId=156722 Agfa has shed its traditional photographic film and consumer imaging business in a management buyout so that it can "focus on its core growth markets of Graphic Systems and HealthCare, which are rapidly going digital" http://news.agfa.com/corporate/news. nsf/news/F07C0210ECC86EA9C1256EF3004D27CE?opendocument
Events like these, and Kodak's announcement earlier this year that it would cease the production of its poneering APS cameras (though not of films) underline the fragility of the conventional photographic market in the face of the growth of digital imaging.
Which begs the question "can we rely on the continued availability of EM film", and if not, how long have we got to plan for the conversion to digital? ============================================================================ Chris is correct in that there has been a real decline worldwide of emulsion based photographic products.
But for the TEM film, the main people who are spreading the "fear" of a "filmless day" soon to arrive are the ones who would benefit the most if one did convert to all digital recording. There could be legitimate reasons to do that, of course, but the inability to purchase high quality TEM film is not going to be one of them at least not in the near and intermediate term future. And besides, someone with a ten or more year old TEM probably is not going to be too keen on making the large capital investment needed to convert to digital anyhow, since the digital add-on would be worth far more than the TEM onto which it is going.
When a large manufacturer decides to get out of a particular business, it is not at all uncommon or unusual for them to find a smaller firm to continue the manufacturing, marketing and distribution of the soon-to-be discontinued product. This makes sense ethically as well since that way they don't leave their existing customers "cut off at their knees". And what would seem like "peanuts" to a large global manufacturer if not also a nuisance could be seen as gigantic volume to a much smaller firm. This kind of licensing of "mature" products being discontinued goes on all the time, including even the marketplace for TEM film. For example, when Agfa " discontinued" the Agfa Scientia brand of TEM film, they licensed a highly reputable German photographic film manufacturing firm, MACO, to continue to manufacturer the TEM films that users around the world had used for their work. And the MACO TEM film is available from PLANO in Germany and from SPI Supplies everywhere else. Everyone can rest well assured that MACO will continue to manufacture TEM film well into the future, farther in fact than most would even want to look.
More information and prices about the MACO film could be found at URL http://www.2spi.com/catalog/photo/maco-TEM-film.html
Disclaimer: SPI Supplies is the worldwide distributor for the MACO TEM film so we have an obviously vested interest in publicizing that fact.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 05:32:46 2004
Chuck Thanks for this clear statement of the position, which will surely reassure many TEM users like myself with a middle-aged instrument and little prospect of going digital at the moment. I will try to relax, but our audio-visual services have just refreshed my paranoia by announcing that they are no longer able to source 35mm slide projectors and lenses suitable for use in lecture theatres. (Is that really true??) We are therefore being encouraged to reduce our dependence on slides.
Best wishes Chris
Dr. Chris Jeffree
----- Original Message ----- } From: "Garber, Charles A." {cgarber-at-2spi.com} To: "MICROSCOPY BB" {Microscopy-at-MSA.Microscopy.com} Sent: Monday, September 06, 2004 9:45 AM
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (stokes-at-saturn.med.nyu.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, September 5, 2004 at 19:04:28 ---------------------------------------------------------------------------
Email: stokes-at-saturn.med.nyu.edu Name: David Stokes
Organization: NYU Skirball Inst
Title-Subject: [Microscopy] [Filtered] EM Core Director
Question: The Skirball Institute of New York University School of Medicine seeks an individual to set up and direct an imaging core facility with an initial emphasis on electron microscopy. The Skirball Insitute, located in midtown Manhattan, consists of 35 laboratories with a diverse array of biomedical research projects, which are described in detail at http://saturn.med.nyu.edu. Individual laboratories currently operate two electron microscopes and several confocal microscopes together with various ancillary equipment for specimen preparation and image analysis. To organize these into a shared facility, we seek an individual with extensive experience in conventional thin sectioning and immunolabeling of biological organisms. The ideal individual will also have management skills and an ambition to develop a comprehensive facility with additional staff offering a wide range of imaging services. Applicants should send their curriculum vitae along with the names and addresses of three references to: Dr. David Stokes at stokes-at-saturn.med.nyu.edu.
We here at the end of the world have already suffered from this "problem" for the last 4 or 5 years where Kodak could not supply us with "EM" grade film. We still use film for all the reasons already quoted; particularly that it still produces the best results compared to digital.
Four years ago we switched suppliers to Agfa and use a product called Copex Positive Pet 10. Code number 2OYAT CNP3 NP EI This is in the 35mm format
Good Luck Raymond Bennett
Keith Williamson EM Unit HortResearch Private Bag 11030 Palmerston North NEW ZEALAND
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chris, ron, and everyone else...
not wanting to sound paranoid, i think we have to consider chris's words carefully.
while digital systems are good, after 35 years i cannot get the same results from a digital image than i can from film, no matter how hard i try. double exposures with polychromatic paper using different filters, dodging, burning, combined, can contribute to excellent prints. perhaps others can get the same results from digitized images, after 5 years, i cannot.
we currently maintain two older Philips 201 and one cm10 microscopes. they have 35mm cameras. Kodak has discontinued manufacture of Direct Positive 5302, which we use for these instruments. the only sources of which i now know for this film is the different suppliers. but how long will their stocks last? what other 35mm format films are there that have the same high resolution that is found with 5302?
all in all, the promise from Kodak is fine and dandy, but will that hold when Kodak decides that the digital market has made it no longer viable to support wet chemistry with specialized, high resolution films such as we require.
of course, as far as i am concerned, i have 25 roles of 5302 in the freezer, so i'm set until our microscopes die. but it is an ongoing concern for the rest of you who were not ordering at the time that Kodak made their decision and were not able to stock up.
paul
Paul R. Hazelton, PhD Electron Microscope Unit University of Manitoba Department of Medical Microbiology 531 Basic Medical Sciences Building 730 William Avenue Winnipeg, Manitoba, Canada, R3E 0W3 e-mail: paul_hazelton-at-umanitoba.ca Phone:204-789-3313 Pager:204-931-954 Cell:204-781-1502 Fax:204-789-3926
The contents of this e-mail are privileged and/or confidential to the named recipient and are not to be used by any other person and/or organisation. If you have received this e-mail in error, please notify the sender and delete all material pertaining to this e-mail. ______________________________________________________
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 15:59:54 2004
Industrial Research Limited is New Zealand's leading industrial scientific research organisation. Our role is to develop leading-edge technologies into practical business opportunities (http://www.irl.cri.nz).
IRL is strengthening its research efforts in the area of superconducting materials and is now seeking expressions of interest from suitably qualified scientists for a two year Post-doctoral position, at our Gracefield site (near Lower Hutt, Wellington). This is an opportunity to join an internationally renowned team working on both Government and industry funded projects in the area of superconducting technologies.
Key responsibilities, knowledge and qualifications: * To undertake applied research and development projects in superconducting materials, including the preparation and analysis of samples of so-called "2nd generation" Yttrium Barium Copper OxideYBCO tapes * PhD or equivalent in materials science or a related field with a focus on TEM work. * Experience in preparing TEM samples of composite materials is highly desirable * To develop and contribute to the development of new research areas * To take an active role in the technology transfer process * Experience with superconducting materials is desirable but not essential * * * The successful applicant will possess: * Strong aptitude for experimental research and development * Demonstrated TEM experience * Good planning, organisational and problem solving skills * An ability to work independently and as part of a team * Superior oral and written communication skills
An attractive remuneration package commensurate with qualifications and experience will be offered to the successful candidate as well as a wonderful opportunity to live in a vibrant capital city in a country renowned for its quality of life and outdoor activities.
'Expressions or Interest' are invited and should be forwarded by 17 September 2004 to: Jennie Scott, Industrial Research Limited, P O Box 31-310, Lower Hutt, Wellington, Phone: (04) 931 3094, Fax: (04) 569 0019, E-mail: j.scott-at-irl.cri.nz {mailto:j.scott-at-irl.cri.nz} .
Industrial Research Limited is an Equal Opportunities Employer
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 16:26:07 2004
i had been thinking of looking up agfa, then they went out of the field. does MACO produce a 35mm film, and is it of similar grain?
paul
Paul R. Hazelton, PhD Electron Microscope Unit University of Manitoba Department of Medical Microbiology 531 Basic Medical Sciences Building 730 William Avenue Winnipeg, Manitoba, Canada, R3E 0W3 e-mail: paul_hazelton-at-umanitoba.ca Phone:204-789-3313 Pager:204-931-954 Cell:204-781-1502 Fax:204-789-3926
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 20:12:30 2004
-----Original Message----- } From: Paul Hazelton [mailto:paul_hazelton-at-umanitoba.ca] Sent: Sunday, September 05, 2004 11:07 AM
double exposures with polychromatic paper using different filters, dodging, burning, combined, can contribute to excellent prints.
---------------------------------------
I don't mean to sound sarcastic, and I know it's going to, so please understand that's not how I mean it.
How is the above different from manipulating a digital image?
I guess there are two parts to the question. First, is this any less of a manipulation (and hence potential for inaccurate or inappropriate artifacts) than using digital techniques to improve the appearance of an image? I mention this in the context of other discussions on digital image integrity.
Second, is the specific problem you're describing a limit of digital technology, or a limit of the skills and resources available? It is certainly a different set of skills to work in a "wet" darkroom than that used in the "digital" darkroom. (Having worked some in both, I too found the digital harder. None the less, I can see where given the right set of tools and skills, the "useable quality" of digital and film could be equal.)
John R.
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 21:41:28 2004
Raymond Bennett wrote: =============================================== We here at the end of the world have already suffered from this "problem" for the last 4 or 5 years where Kodak could not supply us with "EM" grade film. We still use film for all the reasons already quoted; particularly that it still produces the best results compared to digital. =============================================== I am sympathetic to your problem but it is a different kind of problem, not to be confused with the issue already being discussed on the listserver. Your problem is a distribution problem and is the result of the way Kodak (and other manufacturers) distribute(s) their films and other photographic products. Now I am not the last word on this but I am sure someone from Kodak would correct me quite quickly should I be wrong.
My perception is that Kodak establishes an exclusive distributorship, sometimes their own subsidiary, to distribute film in a specific market, such as NZ. But it is up to that particular distributor to decide what they will either a) keep in stock or b) "handle" even on special order. Unfortunately, too many such distributors decide that because the volume is so low, and perhaps they don't want to end up with stale-dated film, they just don't want to be bothered with the handling of such a specialty item like the EM films so they tell their customers it is "not available" but of course, they are really saying it is not available from them, even thought it certainly could be avalable generally, such as in the USA or other countries.
That is why so many end users in New Zealand purchase their Kodak EM film from those firms already providing EM consumables, such as SPI Supplies, Ted Pella, or Ladd (to name a few) in the USA. By ordering this way, you can combine all your other needs for TEM supplies and consumables and the end result is that the incremental shipping costs associated with the film can become almost negligible if not zero. This problem is far from being limited to NZ and in fact gets repeated in many other countries and markets around the world.
With regard to getting superior results with film vs.digital, I hear this all the time, but some would say it would depend on the quality of the digital system you are using to make the comparison. Could you comment on what digital system you used to make your comparison? Those of us who have not converted yet could find your answer very interesting.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 22:09:56 2004
Paul R. Hazelton wrote: ============================================================================ == i had been thinking of looking up agfa, then they went out of the field. does MACO produce a 35mm film, and is it of similar grain? ============================================================================ === Was this a 35 mm film that actually went into the vacuum of the TEM or are you talking about using it with a camera that photographed the screen through a viewing glass from outside the vacuum? Do you have a former Agfa product number?
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 6 23:01:14 2004
certainly your comments are taken well. probably because i agree, and that the whole intent of what i said is that i can manipulate the images much better with wet chemistry after 35 years than i can after 4-5 years digital. i know someone in the advertising business who tells me digital is great, he just hires someone to do the digital work. i cannot do that.
paul
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 02:28:03 2004
Kodak is not the only maker of film on earth and even if 5302 is the best film for the job it is not the only film that will work. Any film can be a direct positive film with the right chemistry. Unless there is some reason it needs to insensitive to red light there are any number of films that with the right developers will work. If you limit it to ortho film the field is a bit limited but there are still a few others.
: : chris, ron, and everyone else... : : not wanting to sound paranoid, i think we have to consider chris's words : carefully. : : while digital systems are good, after 35 years i cannot get the same : results from a digital image than i can from film, no matter how hard i : try. double exposures with polychromatic paper using different filters, : dodging, burning, combined, can contribute to excellent prints. perhaps : others can get the same results from digitized images, after 5 years, i : cannot. : : we currently maintain two older Philips 201 and one cm10 microscopes. : they have 35mm cameras. Kodak has discontinued manufacture of Direct : Positive 5302, which we use for these instruments. the only sources of : which i now know for this film is the different suppliers. but how long : will their stocks last? what other 35mm format films are there that : have the same high resolution that is found with 5302? : : all in all, the promise from Kodak is fine and dandy, but will that hold : when Kodak decides that the digital market has made it no longer viable : to support wet chemistry with specialized, high resolution films such as : we require. : : of course, as far as i am concerned, i have 25 roles of 5302 in the : freezer, so i'm set until our microscopes die. but it is an ongoing : concern for the rest of you who were not ordering at the time that Kodak : made their decision and were not able to stock up. : : paul : : Paul R. Hazelton, PhD : Electron Microscope Unit : University of Manitoba : Department of Medical Microbiology : 531 Basic Medical Sciences Building : 730 William Avenue : Winnipeg, Manitoba, Canada, R3E 0W3 : e-mail: paul_hazelton-at-umanitoba.ca : Phone:204-789-3313 : Pager:204-931-954 : Cell:204-781-1502 : Fax:204-789-3926 : : : : : :
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 07:16:40 2004
Unless the eye alone is used to view a scene (and almost every human eye is different) some form of manipulation has occurred. The minute that an image detector other than the eye is used, its response characteristics define what can be seen or not seen. By definition, every mode of imaging other than that of visible light has been through the selective filtration of some device. As soon as the interpretive and adaptive mechanisms of the brain are divorced from the immediate act of seeing a scene in context, in visible light, a greatly diminished amount of information is available.
For conventional film photography every image has been manipulated from the moment that the decision was made to record it. The choice of one film over another is a choice to use a certain set of contrast and density limits over some other. This is what professionals do - they make informed decisions about work that they are uniquely trained to do. Anyone who is not prepared to take that responsibility for the product of their work probably should stick to an arbitrary formula since they apparently are no more qualified to make those judgments than those to whom any exercise of professional judgment somehow looks suspicious.
John Twilley
Chiphead wrote:
} How is the above different from manipulating a digital image? } is this any less of } a manipulation (and hence potential for inaccurate or inappropriate } artifacts) than using digital techniques to improve the appearance of an } image?
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 08:05:29 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tnicklee-at-uhnresearch.ca) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 6, 2004 at 13:33:11 ---------------------------------------------------------------------------
Question: Has any one performed a dual immunohistochemistry staining, then removed, destained or stripped the antibodies and performed either a second series of immunofluoresecnce or immunohistochemistry? I have heard this discussed at conferences, but can not find a recent reference. We initially are staining for HIF, CA9 and EF5 in fluorescence, which are working. We are then interested in restaining for CD34 (Nova Red in immunohistochemistry) which is a very robust antibody. CD34 should be in regions where HIF CA9 and EF5 are not, but for some reason it is not staining. Thanks for any suggestions.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (sarbu-at-mf.mpg.de) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 6, 2004 at 11:52:53 ---------------------------------------------------------------------------
Email: sarbu-at-mf.mpg.de Name: Corneliu Sarbu
Organization: National Institute for Materials Research, Bucharest
Question: Dear colleagues, does anybody know of a software able to assist in the geometrical drawing of the relationship between the Buergers vector, dislocation line direction, diffraction vector and perhaps electron beam direction (all expressed in a crystallographical way) in in a given material (of cubic symmetry) whose crystallographical constants are known ? The task can be fulfilled by hand drawing, in an approximate way, but I would be pleased to do it in authomatically, as I have to cope with a large variety of such configurations. A software able to display dinamically the variation of all the above mentioned parameters would be great.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Stacey.Andringa-at-uc.edu) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, September 7, 2004 at 06:04:51 ---------------------------------------------------------------------------
Question: I processed quite a few blocks of tissue in Spurr last week. Some of the tissue was actually cell pellets embedded in conical capsules. These blocks and some others in regular capsules did not polymerize in the 2 days as I expected. I left them in the oven over the Holiday, but they are still soft. I may have made the Spurr wrong. Does anyone know if these will eventually cure or can I dig out the tissue, put it into a Spurr/Propylene oxide mixture and try to re-embed it? Of course these were the most important blocks in all that I embedded! Thanks. Stacey Andringa
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (walter.bobrowski-at-pfizer.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, September 7, 2004 at 07:16:18 ---------------------------------------------------------------------------
Email: walter.bobrowski-at-pfizer.com Name: Walt Bobrowski
Organization: Pfizer Global R&D
Title-Subject: [Microscopy] [Filtered] RE: DMP-30 vs. BDMA
Question: Thanks all for the wonderful advice. Yes, old habits die hard! Caroline, I've "always" used the Mollenhauer recipe as I liked it's cutting properties (it felt 'just right'). However, I'm attempting to optimize our 13-year old Lynx el automated processor and moving toward a less-viscous epoxy formulation (Luft with BDMA) for our routine embedding. Yes, I know that Spurr's is even less viscous, but didn't know they had re-formulated the components to be less toxic. I'll have to look into that.
Best regards,
Walter F. Bobrowski Investigative Pathology Safety Sciences Pfizer Global Research & Development Ann Arbor, MI 48105
There are sound-deadening metal channels that are attached to the wall studs, then drywall attached to them. A drywall supply firm could supply them. These, and a second layer of 'rock' over the old one could obviate tearing out the wall. With the enduring popularity of 'bass you can feel', in popular 'music', this should be an ongoing concern. :0)
Paul Grover
-----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] Sent: Saturday, September 04, 2004 12:01 PM To: John Twilley Cc: MSA listserver
Bill Robertson in his reply posting pointed out a very good point about wall mass. My initial thoughts were to put the acoustic tiles in the SEM room. But the noise is more likely coming from the adjacent room where the scroll pump and specimen interchange pump are located. The specimen interchange pump is not at issue. The scroll pump is an Edwards XDS 10. Based on limited experience, it does not sound like it is running as it should. Vacuum is fine but the sound is odd. It makes a mechanical knocking noise. I'm told that either these pumps work almost forever when new or die quickly. Sounds like I have the latter.
In either case, wall mass has a high probability of attention. The intervening wall is standard drywall with 2x4 studs. There is no insulation in the wall. I'm thinking of having liquid foam insulation put in the wall cells and seeing if that helps. In the mean time, the plinth anti-vibration system is not exactly right and will get adjusted shortly. then, based on how that turns out, acoustic tiles in the pump room may be a good solution. If that does not help much, then foam insulation.
The chiller is in the same room as the SEM but turning it off makes no change in image noise. So the noise is external.
Thanks for the replies. Will work on this in the next couple of weeks.
gary g.
At 08:38 AM 9/4/2004, you wrote: } Gary, } } They can be pricey. One reason is that most jurisdictions require that } insulation } applied to an open wall surface be flame proof. Not only that, the } adhesive used } to apply it must also be flame proof, per the fire department and building } codes. } (Recall the disasterous fire in the Rhode Island night club a couple years } back). } Some years ago I found that the material required behind an XRD system in } a small } room ran several hundred dollars for a half dozen panels. However, I } found that I } could get a big improvement by placing a small number of them so as to } block the } noise at the source, rather than covering the more distant wall } surfaces. Buy a } small number to try out, or experiment with packing foam. Have an } assistant hold } them in different locations so as to block acoustic reflections. You may } find that } suspending one from the ceiling in the plane of the instrument helps alot with } noise at the position where the operator sits. } } John Twilley } } Gary Gaugler wrote: } } } } ---------------------------------------------------------------------------- -- } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } ---------------------------------------------------------------------------- --- } } } } I'm looking for sources of acoustic/anachoic } } blocks of foam to reduce SEM room interference. } } } } These are like standard anechoic chamber panels. } } I'm having trouble getting above 400KX without } } mechanical noise from the scroll pump that is } } located in a separate room. The acoustical } } isolation from one room to the other is not } } all that great, I suppose. } } } } Does anyone have experience with suppliers of } } these panels? I'd like to glue them to drywall. } } } } Supplier responses are welcomed as off-line } } messages. } } } } gary g.
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 09:23:18 2004
If you can see the noise as discrete horizontal sinusoids, it is probably low frequency. If you increase your scan speed and the spacing widens proportionately, then you are looking at the true frequency (not an 'aliasing' effect). Also, when aliasing is occurring, the disturbance will often appear to be propagating at an angle, rather than purely horizontal.
A simple way to estimate low frequencies is to employ something that generates a strong 60Hz magnetic field as a reference (I have often used a hand-held magnetic tape eraser). Set up with a relatively high scan speed so that your noise appears as nice sinusoidal disturbances. Count how many cusps you get in a convenient vertical distance. Then turn on the magnetic field generator and do the same for its disturbance. Calculate the ratio vs 60 Hz and you have a crude frequency estimate.
However, if you are seeing discrete sinusoidal disturbances and especially since this is coming from a piece of equipment in a neighboring room that is described as making a "knocking" sound, relatively low frequencies sound like a good bet.
Fred Schamber ASPEX, LLC
Gary Gaugler wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } I don't have any direct way of knowing whether the } noise is high or low frequency. It only shows up } above 60KX as sinusoids on the edges of specimen } details. } } I can probably use FFT to compute the frequency. } It is consistent. } } gary g. } } } At 03:34 PM 9/5/2004, you wrote: } } } } } On Sep 3, 2004, at 6:30 PM, Gary Gaugler wrote: } } } } } I'm looking for sources of acoustic/anachoic } } } blocks of foam to reduce SEM room interference. } } } } } } These are like standard anechoic chamber panels. } } } I'm having trouble getting above 400KX without } } } mechanical noise from the scroll pump that is } } } located in a separate room. The acoustical } } } isolation from one room to the other is not } } } all that great, I suppose. } } } } } } Does anyone have experience with suppliers of } } } these panels? I'd like to glue them to drywall. } } } } } } Supplier responses are welcomed as off-line } } } messages. } } } } Dear Gary, } } Fred has it right about the difference between low and high } } frequencies. The depth of the invaginations in anechoic panels has } } to be 1/4 of the wavelength of the sound in order to be effective, so } } for low frequencies, the panels could take up the entire room, } } leaving no space for the equipment. } } Yours, } } Bill Tivol, PhD } } EM Scientist and Manager } } Cryo-Electron Microscopy Facility } } Broad Center, Mail Code 114-96 } } California Institute of Technology } } Pasadena CA 91125 } } (626) 395-8833 } } tivol-at-caltech.edu } } } } } } }
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 09:38:34 2004
Any professional photo lab could do this for you. Since you are in New York City finding a pro lab should be easy. However, high quality film scanners (from Nikon, Minolta, etc). that do 35 mm slides are relatively cheap these days, you might look into buying one and scanning the slides yourself.
Geoff
Sandra Masur wrote:
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-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 10:44:17 2004
} Can anyone recommend a good commercial photolab that is } also reasonably priced, for scanning hundreds of 35 mm } slides and saving on CDs? } Dear Sandra, I have used Dale Labs in Florida for many years for my personal photography, and one of the services they offer is to scan 35 mm slides onto CDs. Their phone number is (800) 327-1776. I have no affiliation with them except as a long-time satisfied customer. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 10:50:14 2004
Suggest that you contact the Royal Microscopical Society in Oxford, UK. They have a wide range of courses.
Here in the US, John Russ' course at North Carolina State is one of the best respected courses.
Hope this was helpful, Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^& Need a good general text on light microscopy? MME still has copies of Optimizing Light Microscopy available, with discounts for class-sized orders (10 or more). Visit www.MicroscopyEducation.com for details.
At 01:53 AM 9/2/2004, Gareth Morgan wrote:
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From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 11:05:12 2004
Dear Gary, I have had some success here isolating the noise from the pump by putting the pump up on a sound-deadening material such as two-inch-thick packing foam, with a piece of plywood on top. I now have all my rotary pumps up on pads. There can be considerable noise transmission through the floor to your SEM column. Regards and good luck, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: "Gary Gaugler" {gary-at-gaugler.com} To: "John Twilley" {jtwilley-at-sprynet.com} Cc: "MSA listserver" {Microscopy-at-MSA.Microscopy.Com} Sent: Saturday, September 04, 2004 10:00 AM
MACO have contacted me to draw my attention to their company, which produces EM film, X-ray film and other specialist B&W films, printing paper and chemicals. The company partners Oriental in Japan and Lucky in China, both large producers of silver photographic emulsions and coated papers for digital printers.
I had not been aware of the existence of this company until I started this thread on continued availability of EM film, and I suspect that is true of many other EM users on this list.
MACO's web site is at http://www.mahn.net
Prompted by Nestor, I have suggested to MACO that they should list themselves as a company on the MSA Commercial Organisations site http://www.amc.anl.gov/docs/nonanl/wwwform.html
This is not an attempt to promote MACO in particular, and I have no financial interest whatever in this particular company, although I will be trying out their EM film if I can obtain a sample of it here. I merely wish to draw attention to the fact that, as Chuck has pointed out, this is a new company already poised to take over film production if the big companies pull out. It would be a service to us all if other emerging companies producing EM films and related photographic materials and equipment could also be encouraged to register with the Commercial Organisations site.
Best wishes Chris
Dr. Chris Jeffree University of Edinburgh
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 12:36:21 2004
I have inherited some aged vials of crystalline osmium. They are a tad older than normal which is to say 1940! But they look quite normal and are sealed in glass ampules just like a more modern vintage. I have offered 30 x 1 gm vials so it was hard to not accept the gift; this would be a lifetime supply. I intend to try them out in an experiment tomorrow. If anyone knows why this is doomed to failure, please let me know asap. Otherwise I will let the list know how it works out. Tom
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
I need to do some embedding in Lowicryl HM20 and will be ordering new resin kit. I noticed EMS is now offering "MonoStep" - a pre-mixed variation of the normal kit that supposedly gets rid of all the manipulations. Does anybody have some experience with this resin? I am a bit cautious about possible partial polymerization of the pre-made mixture.
Thanks,
Michael
-- Michael Jarnik, Ph.D. Electron Microscope Facility Fox Chase Cancer Center 7701 Burholme Ave. Philadelphia, PA 19111 Tel. 215-728-5675 Fax 215-728-2412
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 15:54:56 2004
} I have inherited some aged vials of crystalline osmium. They are a tad } older than normal which is to say 1940! But they look quite normal and are } sealed in glass ampules just like a more modern vintage. I have offered 30 } x 1 gm vials so it was hard to not accept the gift; this would be a } lifetime supply. I intend to try them out in an experiment tomorrow. If } anyone knows why this is doomed to failure, please let me know } asap. Otherwise I will let the list know how it works out. Tom } } } } Thomas E. Phillips, PhD } Professor of Biological Sciences } Director, Molecular Cytology Core } 3 Tucker Hall } University of Missouri } Columbia, MO 65211-7400 } } 573-882-4712 (office) } 573-882-0123 (fax) } PhillipsT-at-missouri.edu } } } }
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 16:38:43 2004
I know nothing about "dual" immunostaining" but would think that any "destaining", stripping etc will equally affect antibodies and antigens, so I don't expect anything good from it. From another hand, formally speaking you do not limited to only "double" staining in immunofluorescence. With luck, you may use for instance mouse, rat, rabbit, chicken primary antibodies and correspondent secondary with different fluorochromes. Modern confocal microscopes permits to analyze the whole spectrum of emission, so you could separate signals quite easily. I don't know is my approach realistic or not. It would be nice to hear expert's opinion. From biochemical point of view, you could "strip" antibodies by low pH (=2) or a few moles of urea, but it would affect your antigen as well (for bad or good). Sergey
At 08:30 AM 9/7/2004 -0500, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Tue Sep 7 16:50:20 2004
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jcai-at-nanostellar.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, September 7, 2004 at 12:57:14 ---------------------------------------------------------------------------
Email: jcai-at-nanostellar.com Name: Juan
Organization: Nanostellar
Title-Subject: [Microscopy] [Filtered] selective etching method for alumina
Question: Hi all,
I am looking for a way to chemically etch gama-alumina powders, without etching Ag and Cu nano-pariticles mixed in them.
The weight percentage of Ag or Cu in Alumina is around 0.5-1%. We have tried HF, but it turned out Ag and Cu are also etched out.
Hello, Please, could anybody of Philips CM100 users send me a list of coil currents (C1, C2, OBJ etc.)in LM and M Mode at 80kV for magnifications of 1.4kx, 10.5kx and 34kx, respectively. We have some troubles with our EM and unfortunately we do not have a record of these values. Thanking you in advance. Best regards from Prague Oldrich
+-----------------------------------+ Oldrich Benada Acad. Sci. CR Institute of Microbiology Laboratory of electron microscopy Videnska 1083 CZ - 142 20 Prague 4 - Krc Czech Republic +------------------------------------+
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 8 03:17:40 2004
it's unlikely that the samples will polymerise much more after a couple of days. You don't say whether you use Spurr's regularly and this is a first batch to go wrong.
You can certainly try re-embedding but be prepared for a slightly poorer block but be aware of the points below in case it happens again.
Spurr's can poorly embed for a variety of reasons: 1. incomplete dehydration, incomplete removal of alcohol - although if you're using propylene oxide as well I wouldn't have thought so. 2. incomplete impregnation with resin because too little time, not enough stages (eg 50%, 75%, 2x100%), not enough agitation/rotation. 3. bad mix either due to wrong amounts or incomplete mix. 4. One of the components has deteriorated (S1 curing agent may have a shelf life of 6-12 months; I believe that the NSA anhydride hardener can go off especially if exposed to moisture over time)
Spurr's can be affected by moisture so if you chill or freeze it, allow plenty of time for it to reach room temperature and don't leave the lids of the components or mixture for too long.
Malcolm
Malcolm Haswell e.m. unit School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear SR1 3SD UK e-mail: malcolm.haswell-at-sunderland.ac.uk
----- Original Message ----- } From: Stacey.Andringa-at-uc.edu (by way of MicroscopyListserver)
Announcement and Call for Papers Michigan Microscopy &Microanalysis Society Fall Meeting, 2004
Kellogg Center East Lansing, MI November 5, 2004
Abstract Deadline: October 1, 2004
The Fall Meeting of the Michigan Microscopy and Microanalysis Society will be held on November 5th, 2004 at the Kellogg Center in East Lansing, MI. In this one-day conference, there will be two sessions. One is a platform session. This session will have approximately 8-10 speakers representing industry, academia, and research laboratories. The other is a poster session. Please encourage your colleagues who prefer to avoid a platform presentation to submit abstracts for the poster presentations. Some selected abstracts for the oral presentation can be transferred to the poster session if too many abstracts come for the platform session. In addition to the speakers, vendors will exhibit a wide range of products and services of interest to the microscopy community. Presentations are being solicited from researchers in the Physical and Biological Sciences, including one vendor presentation and an invited speaker. Student participation is particularly encouraged. Also, vendors are encouraged to contact the below address to reserve space for product display.
Abstract Submission Please submit a 300 to 350 word abstract by October 1st indicating which session you prefer (poster or presentation)
Geoff Williams MMM President 217 Brooks Hall Biology Department Central Michigan University Mt. Pleasant, MI 48859 Ph 989 774 3576 Fax 989 774 3462 Email: ge.willi-at-cmich.edu
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 8 14:13:31 2004
I need a non standard band pass filter and grating for a B &L monochromatic grating. for 500 nm to 1,000 nm. I know these are not standard. The band pass filter can be wider and a water cell would be actable. I would also like to find a hot or cold mirror that had a 1,100 to 1,400 cut off. All the filters need to have pretty high transmittance in the pass band.
Thanks Gordon Gordon Couger gcc-at-couger.com
I collect links on information related to light microscopes. http://www.couger.com/microscope/links/gclinks.html Please forward any links or information you think might be useful to others. Microscope Manual at www.science-info.org -----
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 8 16:13:41 2004
That's interesting - what was it used for in 1940?
Lesley Weston.
on 07/09/2004 11:03 AM, Tom Phillips at phillipst-at-missouri.edu wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- --} - } } I have inherited some aged vials of crystalline osmium. They are a tad } older than normal which is to say 1940! But they look quite normal and are } sealed in glass ampules just like a more modern vintage. I have offered 30 } x 1 gm vials so it was hard to not accept the gift; this would be a } lifetime supply. I intend to try them out in an experiment tomorrow. If } anyone knows why this is doomed to failure, please let me know } asap. Otherwise I will let the list know how it works out. Tom } } } } Thomas E. Phillips, PhD } Professor of Biological Sciences } Director, Molecular Cytology Core } 3 Tucker Hall } University of Missouri } Columbia, MO 65211-7400 } } 573-882-4712 (office) } 573-882-0123 (fax) } PhillipsT-at-missouri.edu } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 8 17:16:25 2004
Oh, what a good question! I do think osmium was used pre-EM days for some speciality stains (neuro?) but I was given at least 30 ampules so that seems like a lot for someone doing a speciality stain. I don't know which department this chemical stock came from. This came to me via the Environmental Health and Safety people on campus who try to recycle "unwanted" but unused chemicals (a great program you should all get your schools to copy since it says lots of disposal costs and reduces the researcher expenses). It may have been used in some chemical lab but once again seems like a lot. These vials were packaged by Merck (stamped made in Germany), sold by Arthur Thomas and each encased in a nice little wood case. They have 8/13/40 or 1940 written on them. I will let the list know how the expt comes out. Tom
At 02:39 PM 09/08/04 -0700, you wrote: } That's interesting - what was it used for in 1940? } } Lesley Weston. } } } on 07/09/2004 11:03 AM, Tom Phillips at phillipst-at-missouri.edu wrote: } } } } } } } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } ---------------------------------------------------------------------------- } --} - } } } } I have inherited some aged vials of crystalline osmium. They are a tad } } older than normal which is to say 1940! But they look quite normal and are } } sealed in glass ampules just like a more modern vintage. I have offered 30 } } x 1 gm vials so it was hard to not accept the gift; this would be a } } lifetime supply. I intend to try them out in an experiment tomorrow. If } } anyone knows why this is doomed to failure, please let me know } } asap. Otherwise I will let the list know how it works out. Tom } } } } } } } } Thomas E. Phillips, PhD } } Professor of Biological Sciences } } Director, Molecular Cytology Core } } 3 Tucker Hall } } University of Missouri } } Columbia, MO 65211-7400 } } } } 573-882-4712 (office) } } 573-882-0123 (fax) } } PhillipsT-at-missouri.edu } } } } } }
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
OsO4 is used as a histological stain for light microscopy, this use may have preceded its use in Electron Microscopy.
Aaron Hicks Electron Microscopy Preparation Technician
Comparative Physiology and Anatomy Institute of Veterinary, Animal, and Biomedical Sciences Massey University
PN-412 Private Bag 11 222 Palmerston North New Zealand
Phone +64 06 350 4470
-----Original Message----- } From: Lesley Weston [mailto:lesley-at-vancouverbc.net] Sent: Thursday, 9 September 2004 9:39 a.m. To: Tom Phillips; Microscopy-at-msa.microscopy.com
That's interesting - what was it used for in 1940?
Lesley Weston.
on 07/09/2004 11:03 AM, Tom Phillips at phillipst-at-missouri.edu wrote:
} } } ---------------------------------------------------------------------- } -------- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------ ---- --} - } } I have inherited some aged vials of crystalline osmium. They are a } tad older than normal which is to say 1940! But they look quite } normal and are sealed in glass ampules just like a more modern } vintage. I have offered 30 x 1 gm vials so it was hard to not accept } the gift; this would be a lifetime supply. I intend to try them out } in an experiment tomorrow. If anyone knows why this is doomed to } failure, please let me know asap. Otherwise I will let the list know } how it works out. Tom } } } } Thomas E. Phillips, PhD } Professor of Biological Sciences } Director, Molecular Cytology Core } 3 Tucker Hall } University of Missouri } Columbia, MO 65211-7400 } } 573-882-4712 (office) } 573-882-0123 (fax) } PhillipsT-at-missouri.edu } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 8 18:50:27 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (wwiggins-at-carolinas.org) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 8, 2004 at 12:41:52 ---------------------------------------------------------------------------
Question: Does anyone have any details about Ilford going under? There was a blurb in the Guardian about it last month but I've heard nothing here in the States. Worried because I just got an Ilford 2150XL processor 2 years ago and hoped I could stave off digital equipment for a longer while.
Winston I posted a comment about this to the list a few days ago. See below Some further links: http://www.guardian.co.uk/arts/news/story/0,11711,1289459,00.html http://www.manchesteronline.co.uk/business/general/s/128/128120_fears_for_700_jobs_as_ilford_faces_closure.html By contrast, Ilford's web site suggests situation normal! Hope this helps Chris
Dr. Chris Jeffree Inveresk Cottage 26, Carberry Road Inveresk Musselburgh Midlothian EH21 8PR Tel: +44 131 665 6062 FAX +44 131 653 6248 Mobile 07710 585 401 ----- Original Message ----- } From: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} To: {microscopy-at-msa.microscopy.com} Sent: Friday, September 03, 2004 9:05 AM
Dear listserver readers, I am in the lucky position of getting a second hand Micrion 2500 focused ion beam microscope. All the bits are in the room, installation (via FEI) should start soon. However I have no manual, and no information on how it was configured in its previous life. Are there any users of this machine out there who can give me some advice on the 'standard' configuration (particularly things like the gas supplies) - and what it is like to use/maintain? It looks quite frightening to an old TEM/SEM user like me, UHV stainless steel and a million wires and tubes, more like some experimental surface science kit than a routine spec prep tool..
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From MicroscopyL-request-at-ns.microscopy.com Thu Sep 9 07:42:15 2004
OsO4 is a classic stain for the basal bodies of ciliates. It's important in ciliate systematics and morphology, as it reveals the ciliary patterns at the light microscope level. But. It was usually applied to a dish of critters while watching through a microscope. Think Van Gogh's "Starry Night" ...
Phil
} That's interesting - what was it used for in 1940? } } Lesley Weston. } } } on 07/09/2004 11:03 AM, Tom Phillips at phillipst-at-missouri.edu wrote: } } } } } } } } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } ---------------------------------------------------------------------------- } --} - } } } } I have inherited some aged vials of crystalline osmium. They are a tad } } older than normal which is to say 1940! But they look quite normal and are } } sealed in glass ampules just like a more modern vintage. I have offered 30 } } x 1 gm vials so it was hard to not accept the gift; this would be a } } lifetime supply. I intend to try them out in an experiment tomorrow. If } } anyone knows why this is doomed to failure, please let me know } } asap. Otherwise I will let the list know how it works out. Tom } } } } } } } } Thomas E. Phillips, PhD } } Professor of Biological Sciences } } Director, Molecular Cytology Core } } 3 Tucker Hall } } University of Missouri } } Columbia, MO 65211-7400 } } } } 573-882-4712 (office) } } 573-882-0123 (fax) } } PhillipsT-at-missouri.edu } } } } } }
-- Philip Oshel Supervisor, BBPIC microscopy facility Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 - 1284 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 9 08:00:08 2004
Perhaps for Mann's osmium-sublimate fixative, Flemming's fixative or for lipid staining.
Lesley Weston wrote:
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-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 9 09:55:13 2004
Congratulations on your new 2500! It is a great tool, but you are right, it is not really designed to be easy service-able.
The "standard" configuration of gas supply boxes would depend on what are you planning to do with the tool. If you could please describe your intended applications in some detail (materials you plan to deal with and a type of work you intend to do), I would be more then glad to help you with selection of he gas configuration. There are also a lot of routine service and periodic maintenance that you can do on your own with some skill and perhaps a bit of training.
If it is a standard, single-column 2500, then there is no UHV involved, so you would not have to deal with bake-outs.
Please contact me with your application details and if (or rather when) you will have more questions regarding the tool.
Best Regards, Valery Ray Particle Beam Systems & Technology www.partbeamsystech.com
--- Richard Beanland {richard.beanland-at-bookham.com} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The } Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Dear listserver readers, } I am in the lucky position of getting a second hand } Micrion 2500 } focused ion beam microscope. All the bits are in } the room, installation } (via FEI) should start soon. However I have no } manual, and no information } on how it was configured in its previous life. Are } there any users of this } machine out there who can give me some advice on the } 'standard' } configuration (particularly things like the gas } supplies) - and what it is } like to use/maintain? It looks quite frightening to } an old TEM/SEM user } like me, UHV stainless steel and a million wires and } tubes, more like some } experimental surface science kit than a routine spec } prep tool.. } } Many thanks indeed } } Richard } } _______________________________ } Richard Beanland } Analytical Services } Bookham Technology plc } Caswell, } Towcester, } Northamptonshire NN12 8EQ } UK } Tel: +44 (0) 1327 356362 } Fax: +44 (0) 1327 356775 } http://www.bookham.com } } } ======================================================================= } This e-mail is intended for the person it is } addressed to only. The } information contained in it may be confidential } and/or protected by } law. If you are not the intended recipient of this } message, you must } not make any use of this information, or copy or } show it to any } person. Please contact us immediately to tell us } that you have } received this e-mail, and return the original to us. } Any use, } forwarding, printing or copying of this message is } strictly prohibited. } No part of this message can be considered a request } for goods or } services. } ======================================================================= } }
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 9 11:10:22 2004
Dear Listers: We have an SIS Megaview II mounted on the 35mm port of our Hitachi H-7000. This interferes with the proper operation of the left specimen traverse control. If there is anyone out there who has managed to customize the system so that it operates properly, I would appreciate hearing from them. Thanks much Greg
Gregory W. Erdos Ph.D. Assistant Director, Biotechnology Program Scientific Director, Electron Microscopy P.O. Box 118525 217 Carr Hall University of Florida Gainesville, FL 32611 gwe-at-ufl.edu 352-392-1295 fax- 352-846-0251
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 9 16:30:00 2004
We have at least one other installation on an H-7000, where a pulley system was used to "redirect" the control rod. I will send you a document directly that shows the modifications. If you'd like I can get more information for you (parts, prices, etc.)
mike
Michael Bode, Ph.D. Soft Imaging System Corp. 12596 West Bayaud Avenue Suite 300 Lakewood, CO 80228 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: Greg Erdos [mailto:gwe-at-biotech.ufl.edu] Sent: Thursday, September 09, 2004 10:35 To: Microscopy-at-sparc5.microscopy.com
Dear Listers: We have an SIS Megaview II mounted on the 35mm port of our Hitachi H-7000. This interferes with the proper operation of the left specimen traverse control. If there is anyone out there who has managed to customize the system so that it operates properly, I would appreciate hearing from them. Thanks much Greg
Gregory W. Erdos Ph.D. Assistant Director, Biotechnology Program Scientific Director, Electron Microscopy P.O. Box 118525 217 Carr Hall University of Florida Gainesville, FL 32611 gwe-at-ufl.edu 352-392-1295 fax- 352-846-0251
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 9 21:54:27 2004
The easy way (perhaps the easiest way) is to use flexible shaft. We use flexible shafts from Small Parts Inc. (Miami Lakes, FL), www.smallparts.com or 800-220-4242 . Choose panel type, with 1/4" diameter bore at one end and 1/4" diameter shaft at the other end. Then you can simply remove existing left specimen translation knob, attach bore end of the flexible shaft to the left control of the specimen translation control axis, and attach removed knob to the rod end of the flexible shaft. You need also a small (rectangular) piece of aluminum or whatever suitable material to hold the outer end of the shaft at the new location. External end of the flexible shaft attaches to a panel (aluminum piece) in the same way as regular potentiometer or a toggle switch. Flexible shafts 6" and 12" long are standard. Several can be joined for greater length. Part numbers Y-FDCP-4/6 and Y-FDCP-4/12 respectively, cost $19 and $23.
Please see one such camera installation on Hitachi HF-2000 TEM on our web site at http://www.sia-cam.com/2/photo2.html ; http://www.sia-cam.com/2/photo3.html ; http://www.sia-cam.com/2/photo4.html . Both knobs were moved there. It took us 1 hour to complete this modification from scratch.
Do not use flex. shaft much longer than needed, in order to avoid excessive free-play.
Rigid U-joint shafts can be used too, but modification becomes much more complex, since bearing support will be required on each side of a U-joint.
You are welcome to contact me for further details.
Vitaly Feingold Scientific Instruments and Applications 2773 Heath Lane, Duluth GA 30096 (770)232-7785 ph. (770)232-1791 fax (678)467-0012 mobile www.sia-cam.com
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----- Original Message ----- } From: "Greg Erdos" {gwe-at-biotech.ufl.edu} To: "Microscopy-at-sparc5.microscopy.com" {Microscopy-at-MSA.Microscopy.Com} Sent: Thursday, September 09, 2004 12:35 PM
Anyone know of a company that specializes in blue filters for microscopy. We are looking for various sizes and need quite a few for student scopes, some that are not standard sizes. Obviously the dealers have some sizes, but I am really looking for a company that carries a large range of sizes and types.
Thanks for the help! David Burton Optical Specialist University of Washington
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 10 05:10:01 2004
Dear all, Would anyone be kind enough to comment on the relative merits of lens-coupled and fibre-optic-coupled CCD cameras for use at the wide angle (35 mm) port? I am currently investigating the best choice for the recording of diffraction patterns on a FEI Tecnai.
Thanks
-- Ian MacLaren Department of Physics and Astronomy University of Glasgow Glasgow G12 8QQ Scotland http://www.ssp.gla.ac.uk/
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 10 09:05:26 2004
We have a Hitachi H-7500 TEM, and want to have objective aperture strips with four 20 micron apertures made for our scope. Can anyone recommend a company? Thank you.
Hong Emory EM
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 10 12:23:58 2004
Yes, Ladd Research can provide a customized aperture for your scope. I believe we are the only supplier of customized apertures in the U.S., but we can make them available through other EM suppliers if you choose.
Deb Sicard
*Disclaimer: Ladd Research is a dealer of microscopy supplies and accessories
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: ladres-at-att.net
----- Original Message ----- } From: "Hong Yi" {hyi-at-emory.edu} To: {microscopy-at-ns.microscopy.com} Sent: Friday, September 10, 2004 10:30 AM
Dear All:
We have a Hitachi H-7500 TEM, and want to have objective aperture strips with four 20 micron apertures made for our scope. Can anyone recommend a company? Thank you.
Hong Emory EM
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 10 13:52:14 2004
In a message dated 9/9/2004 10:07:31 PM US Mountain Standard Time, dburton-at-nwlink.com writes: Anyone know of a company that specializes in blue filters for microscopy. We are looking for various sizes and need quite a few for student scopes, some that are not standard sizes. Obviously the dealers have some sizes, but I am really looking for a company that carries a large range of sizes and types. David,
The "default" size for such filters seems to be 24 mm or 1 inch, which is probably what you'll find from most of the suppliers unless you want to get plastic filter material in sheet form and make your own.
Try the following:
Edmund Optical at [www.edmundoptical.com]. Click on their Online Catalog, then =} Optics =} Filters and Diffusers =} Color. You'll get several options, such as plastic, glass, wratten filters, etc.
Chroma Technology at [www.chroma.com], Toll Free: 1-800-824-7662. They're a specialty filter manufacturer, but I've used them before for simple blue scope filters. They can cut and mount the filters in any size you want. Paul Millman is the person to talk to.
Good luck, hope this helps.
Cheers,
Bob Chiovetti RMC Products / Boeckeler Instruments, Inc.
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 10 14:58:15 2004
I am immunolabeling cryothin sections of tissue that has been lightly fixed in 2 or 4% paraformaldehyde with 0.1% glut, then cryoprotected, frozen and thin sectioned. Does anyone have antigen unmasking techniques that you are very pleased with for cryothins? If so I would be so happy to hear about them. I have tried 0.005% trypsin, hot citrate buffer, hot tris 9.0 and 0.3% SDS and have had no effect on the antigens that I am trying to unmask. Morphology, however, has suffered! Thank you for any info advance.
Robert Underwood Research Scientist University of Washington
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 10 18:20:54 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (konishi-at-geofourpeaks.com) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Friday, September 10, 2004 at 14:57:52 ---------------------------------------------------------------------------
Question: } I am asking about how to estimate the number of atoms at the surface of nano particles. } } You can roughly estimate the number using volumes of sphere and unit cell. If you assume that ìsurfaceî is the region between two spheres that have a slightly different diameter, you can calculate the volume of ìsurfaceî and estimate the number of atoms in the ìsurfaceî. However, it is a very rough estimation, and it is not clear how to chose the depth to define the surface. } } I would like to know general method in calculation of the number of atoms on surface. Also, if there is a program for such calculation, please advise. } } Thank you, } Hiromi Konishi } Indiana University
PS. I am a member of the list, but currently I cannot post my message for unknow reason, so I used this form.
I would like to transfer the license of MacTempas (HRTEM Image Simulation Software package for Apple) with USB hardware key (fair price!). Interested please contact : benyam-at-recite.ca
Regards, Benyam
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 11 16:10:56 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tauria-at-hotmail.com) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Saturday, September 11, 2004 at 12:06:51 ---------------------------------------------------------------------------
Email: tauria-at-hotmail.com Name: FRANCIS J. PRONESTI
Organization: WORLD ENERGY SERVICES
Education: Graduate College
Location: CASTELLANA GROTTE (BARI), ITALY
Question: I HAVE AN OLD UNITRON SERIES "N" METALURGICAL MICROSCOPE, PURCHASED RECENTLY ON EBAY. I DO OWE MANY OTHER MICROSCOPES, BUT I HAVE FALLEN IN LOVE WITH THIS BEAUTIFUL, OPTICALLY COMPLEX INSTRUMENT. THERE IS NOTHING INTUITIVE ABOUT ITS SET-UP AND USE, SO I WONDER IF ANYONE WOULD KNOW WHERE I CAN BUY A COPY OF THE OPERATING MANUAL FOR THIS SCOPE. FRANCIS J. PRONESTI MSME, MSEE, MEMBER IEEE. PRESIDENT AND OWNER, WORLD ENERGY SERVICES LTD.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dmk8533-at-louisiana.edu) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, September 10, 2004 at 14:05:22 ---------------------------------------------------------------------------
Email: dmk8533-at-louisiana.edu Name: David Krayesky
Question: I have a Olympus Vanox fluoresence microscope in my lab. I need a manual for the microscope to explain what all the different markings mean on the various filters, so I can figure out what the filters do. Does anyone know where I can get this information? I haven't had a lot of luck at Olympus's web site or the FSU microscopy Primer web site.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (burnin1970-at-hotmail.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, September 10, 2004 at 23:17:54 ---------------------------------------------------------------------------
Email: burnin1970-at-hotmail.com Name: Andrei Burnin
Question: Recently, we tried EDS analysis of non-carbon nanotubes (20-30 nm in diameter) in TEM microscope (Tecnai F20). The signals from compositional elements are very tiny in comparison with the cupper and carbon signals, which makes perfect sense, because we used carbon coated grids. However, in the literature such EDS spectra from a spot in one (not from a bunch), say, a ZnO nanotube exhibits signals comparable to those from Cu-C background. How it is possible, that people obtain so high intensity of compositional elements from a very small analyzed area? What should also we do to improve our results of getting EDS from a single nanotube? Is it necessary to use beryllium grids? Thank you.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kssim-at-mmu.edu.my) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Saturday, September 11, 2004 at 04:49:50 ---------------------------------------------------------------------------
I need help from anyone who know the following question:-
Basically I are going to prove that SE3 (secondary electron generated by BSEs at the pole-piece and the chamber wall from returning to the specimen) does not contribute to SE yield. The pole-piece of the SEM is covered by a 2 by 2 inch aluminum (Al) plate that has been painted with carbon paint to absorb SE3s. I need to know the bethe range of K-O range of the target which is carbon on the Al. I could not find this data from 10kev to 30kev range. I believe that with the thickness of 10um is good enough, but not a good reference. Can anyone help me on this?
But expertise says that the SE yield of both AL and carbon are not so high - refer to DC Joy journal paper. So the true SE3 contribution can be much higher. This is one of the points that I need to prove that SE3 still can be ignored.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kssim-at-mmu.edu.my) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, September 12, 2004 at 09:44:09 ---------------------------------------------------------------------------
There is a general statement at http://www.amtimaging.com/english_site/faq_e/faq_e.html#3, not specific to the 35 mm port. However, I too would like to learn more on this.
---- Divakar ------ Physical Metallurgy Section, Indira Gandhi Centre for Atomic Research. Kalpakkam, TN 603102, India
-----Original Message----- } From: Ian MacLaren [SMTP:i.maclaren-at-physics.gla.ac.uk] Sent: Monday, September 13, 2004 9:17 AM To: Microscopy-at-MSA.Microscopy.Com
Dear all, Would anyone be kind enough to comment on the relative merits of lens-coupled and fibre-optic-coupled CCD cameras for use at the wide angle (35 mm) port? I am currently investigating the best choice for the recording of diffraction patterns on a FEI Tecnai.
Thanks
-- Ian MacLaren Department of Physics and Astronomy University of Glasgow Glasgow G12 8QQ Scotland http://www.ssp.gla.ac.uk/
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 08:06:37 2004
The job below is for a TEM specialist although you have to read well through the job particulars http://www.materials.ox.ac.uk/vacancies/Job-Begbroke.pdf to find that out.
---------------------------------- UNIVERSITY OF OXFORD Begbroke Science Park Business-Development Materials Analyst Academic-Related (Research) Grade RSIa - Salary Range - £19,460 - £28,128
Applications are invited for the post of Business Development Materials Analyst for a period of up to two years. The Business Development Materials Analyst will be working for the Institute of Aerospace and Automotive Studies, in collaboration with Faraday Advance and the Oxford Materials Characterisation Service. He or she will be responsible for customer-oriented provision of industrial and analytical materials characterisation services, including provision of rapid high-quality investigations and proposals for materials related problems, and will work to broaden and strengthen the academic and commercial relationships of the OMCS.
The successful candidate is likely to have a track record in materials characterisation, particularly in operating electron microscopes and interpreting the resultant data, an appreciation of business issues including Intellectual Property Rights, budgeting, management and marketing, and the ability to communicate effectively to a range of audiences. An understanding of the Aerospace and Automotive sectors would also be beneficial.
Further particulars of the post are available from Mr Michael Sloane, Begbroke Directorate, Oxford University Begbroke Science Park, Sandy Lane, Yarnton, Oxfordshrire, OX5 1PF. email: michael.sloane-at-begbroke.ox.ac.uk.
Applications, including a covering letter indicating how the applicant meets the requirements of the post together with a detailed curriculum vitae, should reach Mr. Sloane at the above address, by no later than September 30th, 2004. Please include full contact details for three referees, of whom one must be an existing or recent employer. The post can be discussed informally with Dr Alison Crossley, alison.crossley-at-materials.ox.ac.uk, 01865 283726.
The University is an equal opportunities employer.
See also http://www.ox.ac.uk/jobs -- Chris Salter, Oxford Materials Characterisation Service, & Material Science-based Archaeology Group, & Electron Microscopy Research Support Group, Oxford University Begbroke Science Park, Sandy Lane, Yarnton, Oxford, OX5 1PF Tel 01865 283722, EPMA 283741, Mobile 07776031608
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 09:14:18 2004
I am looking for Windows software that allows me to see crystal structure with a less expensive cost. I want to use the software for quick check of number of atoms in a unit cell and distance between atoms, looking at crystal structure, and making figures for publication. Is there any recommendation?
Thank you,
Hiromi Konishi Indiana University
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 10:14:59 2004
I'm interested in hearing opinions (pro/con) from anyone that has experience with the Aspex PSEM. Please call/e-mail me off list if you wish. Thanks for your input and time.
Ed Dargelis Engineering Systems, Inc. 3851 Exchange Ave. Aurora, Il 60504 (630) 851-4566 evdargelis-at-esi-il.com
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 10:28:30 2004
ATOMS is a very good software which I used to do all kinds of work you want to do. But it is a commercial software, I do not know its exact price.
You can go to IUCr's (International Union of Crystallography)Webster and check their software page, you will find a lot of free software.
Best regards,
Ying
-----Original Message----- } From: hkonishi-at-indiana.edu [mailto:hkonishi-at-indiana.edu] Sent: Monday, September 13, 2004 9:42 AM To: Microscopy-at-MSA.Microscopy.Com
I am looking for Windows software that allows me to see crystal structure with a less expensive cost. I want to use the software for quick check of number of atoms in a unit cell and distance between atoms, looking at crystal structure, and making figures for publication. Is there any recommendation?
Note: The information contained in this message may be privileged and confidential and thus protected from disclosure. If the reader of this message is not the intended recipient, or an employee or agent responsible for delivering this message to the intended recipient, you are hereby notified that any dissemination, distribution or copying of this communication is strictly prohibited. If you have received this communication in error, please notify us immediately by replying to the message and deleting it from your computer. Thank you.
Marcus, are you out there? If anyone knows how to contact Marcus, please respond off line. Thanks, Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 11:24:25 2004
A tool developed in '99, visit www.Evactron.com actively prevents sample 'image darkening effects' from polymerized hydrocarbon contamination. The device is commercially available, improves DP & TMP system vacuum levels safely without damage to SEM/FIB components and eliminates manual iso swabbing cleanup.
XEI Scientific
-----Original Message----- } From: j.bilde-at-risoe.dk [mailto:j.bilde-at-risoe.dk] Sent: Thursday, September 02, 2004 12:16 AM To: pzou-at-feico.com Cc: microscopy-at-microscopy.com
Dear Pei Zou,
See section 9.10.6 in Goldstein et al.: Scanning electron microscopy and X-ray analysis" Plenum Press 1992. It begins: "A sample subjected to electron bombardment in a diffusion-pumped vacuum gradually becomes ccovered with a contamination layer due to polymerization, under the action of the beam, of organic matter adsorbed on the surface".
Ways to reduce the effect are: clean vacuum, clean sample, cold finger.
-----Original Message----- } From: by way of MicroscopyListserver [mailto:pzou-at-feico.com] Sent: 2. september 2004 04:16 To: microscopy-at-microscopy.com
Dear Gary,
I use 4 " sorbothane " mounts (stock number: NT35-264). Usually I use it for holography but it worked well for damping the rot. pump vibrations.
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Bases and Platforms {http://www.edmundoptics.com/onlinecatalog/displayproduct.cfm?productID=1255&search=1}
Large Workstation Platform: Hard-coat black anodized aluminum, 13"W x 18"L x 1/2"T platform with sorbothane feet and 3/4" threaded post hole. Weighs 12 lbs. Small Platform Base: Hard-coated black anod... ---
Best regards Timo Junker
Gary Gaugler schrieb:
} ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } I'm looking for sources of acoustic/anachoic } blocks of foam to reduce SEM room interference. } } These are like standard anechoic chamber panels. } I'm having trouble getting above 400KX without } mechanical noise from the scroll pump that is } located in a separate room. The acoustical } isolation from one room to the other is not } all that great, I suppose. } } Does anyone have experience with suppliers of } these panels? I'd like to glue them to drywall. } } Supplier responses are welcomed as off-line } messages. } } gary g.
I would suggest that you contact John Coyle at Unitron. You can reach him by email at : johnc-at-unitronusa.com.
DISCLAIMER: South Bay Technology distributes Unitron Microscopes and, therefore, has a vested interest in promoting their use.
Best regards-
David
-- David Henriks
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by way of Ask-A-Microscopist wrote:
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From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 12:56:44 2004
It seems that during the summer months, we have no end of troubles with our plastic not infiltrating the tissue. I've always assumed that this was because of the humidity rising quite a bit during the summer months, and the poor air conditioning.
It comes out as sections falling apart when I try to flatten them in the water boat, or breaking in the electron beam. We have always kept our epoxy resin in the freezer in 30 ml plastic syringes, but it usually only gives us a problem during the summer months. During the winter here, the air is extremely dry.
We do 4 rinses in absolute alcohol, followed by 3 rinses in propylene oxide, followed by a 50/50 mixture of epon/araldite mixed with propylene oxide over night with the caps off, for the propylene oxide evaporating overnight and slowly leaving the specimens in 100% plastic by morning. We polymerize at 70 deg C overnight, keeping a dessicant in our embedding oven to keep the air as dry as possible too.
Are there any resins which might be used for electron microscopy which are more tolerant of water, and which would give a good result with human tissue in the electron microscope?
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 13:13:39 2004
Thanks everyone. I've found him. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 13:23:55 2004
I think Mercury is a free viewer. Check out http://www.ccdc.cam.ac.uk/products/csd_system/mercury/ It is decent and works in windows.
-----Original Message----- } From: hkonishi-at-indiana.edu [mailto:hkonishi-at-indiana.edu] Sent: Monday, September 13, 2004 9:42 AM To: Microscopy-at-MSA.Microscopy.Com
I am looking for Windows software that allows me to see crystal structure with a less expensive cost. I want to use the software for quick check of number of atoms in a unit cell and distance between atoms, looking at crystal structure, and making figures for publication. Is there any recommendation?
Thank you,
Hiromi Konishi Indiana University
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 13:30:21 2004
Dear Andrei, The subject of EDS spectra in a TEM or STEM is complex and there are many different configurations of EDS detector in the TEM column. Some of these account for the differences in relative background and specific element signal. If you have a horizontal-mount detector and then tilt the sample towards the detector, this will pick up a lot of grid and carbon x-rays. I have a TEM with a high-takeoff-angle detector (68 degrees) and that results in a much lower Cu and C signal when looking at tiny particles on a carbon-coated copper grid. A 30 mm2 detector will also pick up more counts than a 10 mm2. The material in the specimen holder and the column above and below the sample also affects the relative background of your spectrum. A TEM really needs to be designed for EDS at the factory, with light-element inserts to reduce the secondary x-ray radiation that generates much of your background. The better spectra you are referring to may have come from a TEM or STEM designed or modified to give the best EDS performance, perhaps at the cost of less performance in other areas. I know my TEM was designed for analytical work, at the sacrifice of ultimate resolution. To increase your counts you can try lowering the accelerating voltage, increasing the spot size, increasing the condenser aperture size and, of course, increasing the time you count for. Make sure your objective aperture is removed, the specimen holder is specified for EDS and your EDS detector is properly aligned. Good luck, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca
----- Original Message ----- } From: "by way of MicroscopyListserver" {burnin1970-at-hotmail.com} To: {microscopy-at-microscopy.com} Sent: Saturday, September 11, 2004 2:43 PM
Hi, David
There are several sources, depending on the optical requirements. Both Chroma and Omega Filters provide an extensive line of filters. If this are fairly simple scopes, you might also try looking at Edmund Scientific.
If you have trouble finding any of the above, please contact me off-line.
Hope this is helpful.
Best regards, Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^& Need a good general text on light microscopy? MME still has copies of Optimizing Light Microscopy available, with discounts for class-sized orders (10 or more). Visit www.MicroscopyEducation.com for details. At 11:20 PM 9/9/2004, David Burton wrote:
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From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 15:51:36 2004
Gary; Your procedure is similar to the one I use here in Houston where we have very high humidity also. The difference is that my Epon/araldite: PO step is for one hour followed by one hour in pure resin then embedded in capsules in fresh resin. This sets at RT for 3-4 hours then in the oven overnight at 80 degrees C. I have been using this method for over 20 years with no problems.
Mannie Steglich Tech Dir E M Lab M D Anderson Cancer Center
Garry Burgess {GBurgess-at-exchange.hsc.mb.ca}
09/13/2004 01:24 PM
To: {Microscopy-at-msa.microscopy.com} cc:
It seems that during the summer months, we have no end of troubles with our plastic not infiltrating the tissue. I've always assumed that this was because of the humidity rising quite a bit during the summer months, and the poor air conditioning.
It comes out as sections falling apart when I try to flatten them in the water boat, or breaking in the electron beam. We have always kept our epoxy resin in the freezer in 30 ml plastic syringes, but it usually only gives us a problem during the summer months. During the winter here, the air is extremely dry.
We do 4 rinses in absolute alcohol, followed by 3 rinses in propylene oxide, followed by a 50/50 mixture of epon/araldite mixed with propylene oxide over night with the caps off, for the propylene oxide evaporating overnight and slowly leaving the specimens in 100% plastic by morning. We polymerize at 70 deg C overnight, keeping a dessicant in our embedding oven to keep the air as dry as possible too.
Are there any resins which might be used for electron microscopy which are more tolerant of water, and which would give a good result with human tissue in the electron microscope?
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 16:19:13 2004
I have two JB-4 microtomes that I would like to repair, even if only to get one to work. Neither advance. One has electronic advance and the other doesn't. I have experience in repairing older ultramicrotomes, but I'd like to have a set of service instructions, a blueprint, or even a users' manual in hand before tearing them apart!
Does anyone have these documents? More importantly, perhaps, advice?
Mahalo, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 17:29:13 2004
} } That's interesting - what was it used for in 1940? } } Lesley Weston. } } on 07/09/2004 11:03 AM, Tom Phillips at phillipst-at-missouri.edu wrote: } } } } } I have inherited some aged vials of crystalline osmium. They are a tad } } older than normal which is to say 1940! But they look quite normal and are } } sealed in glass ampules just like a more modern vintage. I have offered 30 } } x 1 gm vials so it was hard to not accept the gift; this would be a } } lifetime supply. I intend to try them out in an experiment tomorrow. If } } anyone knows why this is doomed to failure, please let me know } } asap. Otherwise I will let the list know how it works out. Tom } } } } } } } } Thomas E. Phillips, PhD
-- Dr. Steven Barlow EM Facility/Biology Dept. San Diego State University 5500 Campanile Drive San Diego CA 92182-4614 phone: (619) 594-4523 fax: (619) 594-5676 email: sbarlow-at-sunstroke.sdsu.edu http://www.sci.sdsu.edu/emfacility
Chairman, Educational Outreach subcommittee promoting microscopy instruction and increased access to microscopes Microscopy Society of America http://www.msa.microscopy.com/
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 13 18:23:35 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (adriana-at-cab.cnea.gov.ar) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 13, 2004 at 13:56:32 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] MListserver: CCD for TEM in material science
Question: Dear microscopists,
We are interested in buying a CCD camera for a Philips CM200 TEM with an ultratwin lens for HRTEM. The main use is for materials science. We would like to hear comments on the performance of the KeenView CCD camera offered by SIS. We are particularly concerned with the sensibility for HRTEM (for example typical exposure times) and with the performance of the 12 bit camera for diffraction pattern acquisition and for online astigmatism correction using FFT.
All comments will be appreciated.
Thank you for your help.
Adriana
---------------------------------------------------------- Adriana CondÛ CONICET Researcher Metals Physics Group Centro AtÛmico Bariloche 8400 San Carlos de Bariloche ARGENTINA e-mail: adriana-at-cab.cnea.gov.ar Fax: +54 2944 445299 TE: +54 2944 445290 ----------------------------------------------------------
Alfred Nobel used Osmium in the production of Explosives in the 1860's. He is best known for discovering Nitroglycerin & Dynamite (TNT). Now of course his last will & testament funds "The Nobel Prize".
Al Coritz Electron Microscopy Sciences www.emsdiasum.com
----- Original Message ----- } From: {sbarlow-at-sunstroke.sdsu.edu} To: {microscopy-at-msa.microscopy.com} Sent: Monday, September 13, 2004 6:56 PM
} } It seems that during the summer months, we have no end of } troubles with our plastic not infiltrating the tissue. I've } always assumed that this was because of the humidity rising } quite a bit during the summer months, and the poor air conditioning. } } It comes out as sections falling apart when I try to flatten } them in the water boat, or breaking in the electron beam. We } have always kept our epoxy resin in the freezer in 30 ml } plastic syringes, but it usually only gives us a problem } during the summer months. During the winter here, the air is } extremely dry. } } We do 4 rinses in absolute alcohol, followed by 3 rinses in } propylene oxide, followed by a 50/50 mixture of epon/araldite } mixed with propylene oxide over night with the caps off, for } the propylene oxide evaporating overnight and slowly leaving } the specimens in 100% plastic by morning. We polymerize at } 70 deg C overnight, keeping a dessicant in our embedding oven } to keep the air as dry as possible too.
Is dessicant effective at 70 deg C, or will it release moisture accumulated at room temperatures?
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 14 08:15:50 2004
I've been following this thread of conversation on old OsO4 with great interest. I know that there are many (?) books, articles written on the history of microscopes (esp. electron microscopes), but is there an article/book written on such interesting facts i.e. OsO4 and othere EM fixatives, etc. I would love to read more......
Peggy
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Sample Prep [mailto:sampleprep-at-earthlink.net] Sent: Monday, September 13, 2004 8:56 PM To: microscopy-at-msa.microscopy.com; sbarlow-at-sunstroke.sdsu.edu
Alfred Nobel used Osmium in the production of Explosives in the 1860's. He is best known for discovering Nitroglycerin & Dynamite (TNT). Now of course his last will & testament funds "The Nobel Prize".
Al Coritz Electron Microscopy Sciences www.emsdiasum.com
----- Original Message ----- } From: {sbarlow-at-sunstroke.sdsu.edu} To: {microscopy-at-msa.microscopy.com} Sent: Monday, September 13, 2004 6:56 PM
Hi Tina,
We manufacture the JB-4 and JB-4A here at EBS. It is our policy to provide user manuals and telephone support for all of our instruments at no charge. Please send us your contact information and we'll mail a manual right out to you. Beyond that, we can also provide you with factory parts and professional repair services, in the event that you should you need either.
Sincerely,
Michael R. Nesta General Manager Energy Beam Sciences, Inc. 800 992-9037 mnesta-at-ebsciences.com www.ebsciences.com "Adding Brilliance to Your Vision"
Tina Carvalho wrote: } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Hi, All- } } I have two JB-4 microtomes that I would like to repair, even if only to } get one to work. Neither advance. One has electronic advance and the other } doesn't. I have experience in repairing older ultramicrotomes, but I'd } like to have a set of service instructions, a blueprint, or even a users' } manual in hand before tearing them apart! } } Does anyone have these documents? More importantly, perhaps, advice? } } Mahalo, } Tina } } **************************************************************************** } * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * } * Biological Electron Microscope Facility * (808) 956-6251 * } * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* } **************************************************************************** } } }
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 14 12:52:37 2004
as the manufacturer of the KeenView system I would like to provide the following information:
1) The KeenView has a high anti-bloom factor of about 300. This means that a pixel can hold 300 times the saturation charge before blooming occurs.
2) A KeenView system is delivered with the analySIS iTEM software, which includes a Real-time FFT option for focusing and astigmatism correction.
3) Sensitivity is usually not an issue, unless you wish to do low-dose cryo-TEM. If you let me know what you plan to do, I can see if I have some images that I could send you.
Please contact me if you have further questions.
Mike
Michael Bode, Ph.D. Soft Imaging System Corp. 12596 West Bayaud Avenue Suite 300 Lakewood, CO 80228 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: adriana-at-cab.cnea.gov.ar [mailto:adriana-at-cab.cnea.gov.ar] Sent: Monday, September 13, 2004 17:52 To: microscopy-at-microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (adriana-at-cab.cnea.gov.ar) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 13, 2004 at 13:56:32 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] MListserver: CCD for TEM in material science
Question: Dear microscopists,
We are interested in buying a CCD camera for a Philips CM200 TEM with an ultratwin lens for HRTEM. The main use is for materials science. We would like to hear comments on the performance of the KeenView CCD camera offered by SIS. We are particularly concerned with the sensibility for HRTEM (for example typical exposure times) and with the performance of the 12 bit camera for diffraction pattern acquisition and for online astigmatism correction using FFT.
All comments will be appreciated.
Thank you for your help.
Adriana
---------------------------------------------------------- Adriana CondÛ CONICET Researcher Metals Physics Group Centro AtÛmico Bariloche 8400 San Carlos de Bariloche ARGENTINA e-mail: adriana-at-cab.cnea.gov.ar Fax: +54 2944 445299 TE: +54 2944 445290 ----------------------------------------------------------
I don't think water may penetrate epoxy resin easily. The picture, you described is more like you have water in your sample before plastic. As far as I could see, the 100% Et-OH step may be a problem in your recipe. 4x 100% Et-OH sounds excessive (I always do 2x20 min 100% Et-OH) - so to me it looks like that your 100% Et-ON is not a 100% (if you still have water after 4x). You may need to change the batch or use molecular sieve. You need also to use at least 1 ml of Et-OH per sample. It's better to do it in hermetic vials like 1.5 ml Eppendorf tube. If I forgot Et-OH bottle to close immediately after use, I consider it's 95% Et-OH and use it for 30-95% dehydratation steps only. 3x of PO is sounds excessive too. The trick here is that as more changes you do, as more your sample has exposed to water (when you emptied the vial). So, 1 exchange of PO (and Et-OH) is much better than 3 (you still need 2 for Et-OH) in my point of view. You need to use huge excess of the solvents - at least 1 ml per sample. As soon as your sample in PO (or epoxy) - it's protected from water by the layer of solvent (epoxy). PO and epoxy do not dissolve water, therefore it could not reach your sample. You still may have a droplets of water (from condensation for instance) but it'll generate a different picture: round empty spaces in your sample. You may try also Spurr resin - it may tolerate a few %% of water in your sample. I hope it'll help, Sergey
At 06:38 AM 9/14/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-080 Los Angeles, CA 90095
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (snyderlt-at-newpaltz.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, September 14, 2004 at 08:42:51 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] service for philips 300
Question: We have a philips 300 TEM that has been under service contract for the past 4 years. The company is no longer available and I am trying to find a service provider. We are located in New Paltz, NY, about half way between NYC and Albany.
Any suggestions would be greatly appreciated. Teresa
If the problem is caused by high humidity in the air, you could try using a water-miscible resin such as Durcupan or Aquembed. They're both a lot more expensive than Epon-substitutes and Araldite and they polymerise to a rather soft block, which may or may not matter depending on your tissue, but it would eliminate the problem if that is the cause. Another possibility is that there might still be a trace of PO left, if you don't do one more 100% in the morning before transferring to the DMP-30 mix, but I don't know why that would happen only in summer. Hope this helps.
Lesley Weston.
on 13/09/2004 11:24 AM, Garry Burgess at GBurgess-at-exchange.hsc.mb.ca wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- --} - } } } It seems that during the summer months, we have no end of troubles with our } plastic not infiltrating the tissue. I've always assumed that this was } because of the humidity rising quite a bit during the summer months, and the } poor air conditioning. } } It comes out as sections falling apart when I try to flatten them in the } water boat, or breaking in the electron beam. We have always kept our epoxy } resin in the freezer in 30 ml plastic syringes, but it usually only gives us } a problem during the summer months. During the winter here, the air is } extremely dry. } } We do 4 rinses in absolute alcohol, followed by 3 rinses in propylene oxide, } followed by a 50/50 mixture of epon/araldite mixed with propylene oxide over } night with the caps off, for the propylene oxide evaporating overnight and } slowly leaving the specimens in 100% plastic by morning. We polymerize at } 70 deg C overnight, keeping a dessicant in our embedding oven to keep the } air as dry as possible too. } } Are there any resins which might be used for electron microscopy which are } more tolerant of water, and which would give a good result with human tissue } in the electron microscope? } } This e-mail and/or any documents in this transmission is intended for the } address(s) only and may contain legally privileged or confidential } information. Any unauthorized use, disclosure, distribution, copying or } dissemination is strictly prohibited. If you receive this transmission in } error, please notify the sender immediately and return the original. }
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 14 19:37:02 2004
I have to take exception to some of the assertions made in a recent response to Garry Burgess' posting. First, propylene oxide is completely miscible with water. And second, Spurr's resin is much more likely to have polymerization problems caused by high humidity than is Epon/Araldite type resin mixtures. Spurr's should be cured either in sealed plastic capsules, or if you prefer to use flat molds, with ample fresh desiccant present. I place the flat molds in a sealed Tupperware container, along with plenty of desiccant. The first time I used Spurr's in flat molds on a humid day, I didn't use a desiccant, and the blocks were too soft to section. I repeated the embedding using a desiccant, and the same batch of chemicals, and the blocks were perfect.
We routinely cure our specimens in Epon/Araldite/DDSA without using a desiccant, and never have problems despite summer humidities of 70% or more. We use 3 x 10m changes of 100% ethanol, 3 x 10m changes in PO, infiltrate overnight in 50% resin, and 8 hours the next day in 100% resin, but always in sealed vials. Leaving vials of 50% resin/PO open to the atmosphere on a humid day will result in water absorption, and might be the cause of the problem. Also, I suspect that the propylene oxide might not be completely removed. We use commercial 100% ethanol from plastic bottles with no problems. As was discussed recently in this forum, use of a molecular sieve can result in particles embedded in your specimen, and damage to diamond knives.
In the recent discussion of the merits of BDMA I was surprised to see that people are pre-mixing their resin with DMP-30 or BDMA, and freezing it. Your resin mix will last much longer (at least a couple of months) if you leave out the accelerator. We mix up large batches of our resin, and freeze aliquots of various sizes in glass vials. When we need the resin, we warm it to room temperature, add the appropriate amount of DMP-30, and mix thoroughly. You may spend a little more time mixing in the DMP, but you won't have to worry about your resin thickening in the freezer.
Ralph Common Michigan State University Division of Human Pathology
-------------- Original posting from Garry Burgess:
It seems that during the summer months, we have no end of troubles with our plastic not infiltrating the tissue. I've always assumed that this was because of the humidity rising quite a bit during the summer months, and the poor air conditioning.
It comes out as sections falling apart when I try to flatten them in the water boat, or breaking in the electron beam. We have always kept our epoxy resin in the freezer in 30 ml plastic syringes, but it usually only gives us a problem during the summer months. During the winter here, the air is extremely dry.
We do 4 rinses in absolute alcohol, followed by 3 rinses in propylene oxide, followed by a 50/50 mixture of epon/araldite mixed with propylene oxide over night with the caps off, for the propylene oxide evaporating overnight and slowly leaving the specimens in 100% plastic by morning. We polymerize at 70 deg C overnight, keeping a dessicant in our embedding oven to keep the air as dry as possible too.
Are there any resins which might be used for electron microscopy which are more tolerant of water, and which would give a good result with human tissue in the electron microscope?
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 14 20:11:54 2004
while the thread has been very interesting, your question has been the most helpful to me. there is a book, it is Hayat's Fixation for Electron Microscopy. publication date was around 1980. it does not cover some of the history, but it is an excellent book if you can get your hands on a copy of it. the help to me....if i can just find that student who borrowed my copy in june.....
paul
Paul R. Hazelton, PhD Electron Microscope Unit University of Manitoba Department of Medical Microbiology 531 Basic Medical Sciences Building 730 William Avenue Winnipeg, Manitoba, Canada, R3E 0W3 e-mail: paul_hazelton-at-umanitoba.ca Phone:204-789-3313 Pager:204-931-954 Cell:204-781-1502 Fax:204-789-3926
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 14 20:56:29 2004
I am chronicallly plagued by humidity problems here. Besides making sure the bottles of absolute ethanol, propylene oxide and resin components, are opened as little as possible, I put my vials with samples over dessicant whenever I'm using absolute ethanol, PO, or resin. Always. I have a rotator we made years ago that accommodates film cans with dessicant into which my vials fit. I keep dessicant in the embedding oven. And, imortantly, I pre-heat my molds and labels overnight in the oven before putting in the samples. This preheating of the capsules or molds and paper labels also keep bubbles from forming so that I never have to pull a vacuum on them any more. Yes, it's totally anal, but lots of TEM is!
Good luck!
Aloha, Tina
} } It seems that during the summer months, we have no end of troubles with our } } plastic not infiltrating the tissue. I've always assumed that this was } } because of the humidity rising quite a bit during the summer months, and the } } poor air conditioning. } } } } It comes out as sections falling apart when I try to flatten them in the } } water boat, or breaking in the electron beam. We have always kept our epoxy } } resin in the freezer in 30 ml plastic syringes, but it usually only gives us } } a problem during the summer months. During the winter here, the air is } } extremely dry. } } } } We do 4 rinses in absolute alcohol, followed by 3 rinses in propylene oxide, } } followed by a 50/50 mixture of epon/araldite mixed with propylene oxide over } } night with the caps off, for the propylene oxide evaporating overnight and } } slowly leaving the specimens in 100% plastic by morning. We polymerize at } } 70 deg C overnight, keeping a dessicant in our embedding oven to keep the } } air as dry as possible too.
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 03:43:01 2004
Sergey A propylene oxide layer over the specimen cannot protect against water penetration. Water's solubility in 1,2 propylene oxide (=1,2 epoxypropane) is 14.7 wt%. Conversely, PPO's solubility in water is 40.5 wt%.
Chris
----- Original Message ----- } From: "Sergey Ryazantsev" {sryazant-at-ucla.edu} To: {Microscopy-at-microscopy.com} Sent: Tuesday, September 14, 2004 8:18 PM
How effective is molecular sieve as a desiccant at typical polymerization temperatures (60-70 oC)?? I suspect not very. Would there be a better choice?
I would also be interested to know what strategies people now recommend for drying EM solvents without getting them contaminated with bits of desiccant.
And another thing - how on earth can one determine whether the desiccant is still being effective? Can you trust the blue indicator in molecular sieves? What is the equilibrium water concentration in ethanol above fresh molecular sieve (assuming the ethanol contained some water initially), and does this increase when the MS contains, say, 10% water (half its saturation capacity)?
Best wishes Chris
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 05:35:21 2004
Each time I post a message to the list a flurry of autoforwarded replies is generated - "I am in the Bay area", I am on holiday", "I can be reached at blah blah!", "I am canoeing down the Danube", Etc. Although fascinating at a sociological level, this is not maximally useful web traffic at the EM level. Might I respectfully suggest that listers try to prevent these messages from reaching the list?
Best regards Chris
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 07:47:27 2004
Dear Listservers: We have a CM-10 we wish to do both EDS and low temp work with. We have the Be Gatan cryo-blades from a 400 but need to adapt the column phlange to fit the CM column. Neither Gatan nor FEI have been able to resolve this as yet. Does anyone have any advice or spare phlanges so we can get on with this? I am thankful for any and all advice I can get. Bob Harris
The Listserver rules ask you to NOT USE "I'm not in the office" autoreplies, instead you should unsubscribe when you go on vacation or out of town.
If you job requires you to use "out of the office" messages for your work Email then can I suggest you ask your local system guru to setup a second Email address for Microscopy which you can recieve all the message at and NOT set this function.
I will remind you that ALL postings are archived and if you miss something you can find it at:
http://www.microscopy.com/MicroscopyListserver
It is also possible to have multiple addresses subscribed, so there are alot of ways to mitigate this problem with only a few minutes of your time. The rest of the community will appreciate your thoughtfulness and consideration.
Cheers..
Nestor Your Friendly Neighborhood SysOp
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 08:39:54 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tem_iopb-at-iopb.res.in) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 15, 2004 at 05:15:20 ---------------------------------------------------------------------------
Email: tem_iopb-at-iopb.res.in Name: P. V. Satyam
Organization: Institute of Physics, Bhubaneswar,India
Question: Dear All, I would like to do TEM on (1) Metal/Semiconducting nano-particles dispersed on polymer surfaces (for example: Ge on PS) (2)Langmuir-Blodgett Films
We have JEOL 2010 Ultra High Resolution TEM operating at 200kV and LaB6 Filament.
We tried to work at 200 kV using standard double tilt holder (room temperature) but found that polymeric samples started melting upon e-beam irradiation. We dont want to coat any high Z conducting layer on our sample.
As majority of our work (in materials science) is done using 200 kV, I am concerned to work at lower energies.
Does it help having a cooling holder? If so, are there any particulars that should be taken care before purchasing so as enable us to work with polymer and LB films?
I will appreciate your suggestions either directly to me or to the list.
Best regards Satyam Institute of Physics, Bhubaneswar, India Home page: www.iopb.res.in/~tem_iopb
to avoid the bits in your solvents, try putting your molecular sieve (or whatever) into a length of dialysis tubing. You can fold the ends over and staple them to form a small sausage.
If you get any answers to the rest of your questions I, for one, would be interested to hear them.
Malcolm
Malcolm Haswell e.m. unit School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear SR1 3SD UK e-mail: malcolm.haswell-at-sunderland.ac.uk
----- Original Message ----- } From: Chris Jeffree {c.jeffree-at-ed.ac.uk}
Hi Chris,
I am out of the office and canoeing on the Current River. I will reply to your email soon, if I don't drown.
Seriously though, I asked a similar question a while back and was told that molecular sieves can be put in dialysis tubing to keep them from chunking up your solvents. Basically, we have quit using molecular sieves just in case they were responsible for what we thought was excessive wear on our diamond knives. Since we use a lot of microwave processing, we generally use acetone, rather than ethanol, and mix our dehydration series fresh each time. When I use ethanol, I usually finish up with a recently opened bottle of absolute, but after two or three uses I relegate that bottle to the 90-95% category, and open up a new one for the final steps.
This seems to work for us, as we rarely have infiltration problems, even with retinal tissue which can be problematic.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
-----Original Message----- } From: Chris Jeffree [mailto:c.jeffree-at-ed.ac.uk] Sent: Wednesday, September 15, 2004 5:40 AM To: microscopy-at-msa.microscopy.com
How effective is molecular sieve as a desiccant at typical polymerization temperatures (60-70 oC)?? I suspect not very. Would there be a better choice?
I would also be interested to know what strategies people now recommend for drying EM solvents without getting them contaminated with bits of desiccant.
And another thing - how on earth can one determine whether the desiccant is still being effective? Can you trust the blue indicator in molecular sieves? What is the equilibrium water concentration in ethanol above fresh molecular sieve (assuming the ethanol contained some water initially), and does this increase when the MS contains, say, 10% water (half its saturation capacity)?
Best wishes Chris
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 09:04:23 2004
Malcolm Thanks for this.I have heard it suggested previously, but I am sceptical about it.
1) Presumably the dialysis tubing has to be wet before it can be loaded?. OK, so you can re-dry the MS after loading. 2) I wonder what the permeability of dialysis tubing is to water vapour when very dry and immersed in ethanol. It that environment it will be in its most ultra-compact precipitated state (since ethanol precipitates polysaccharides from aqueous solution). 3) As in my original question, do you really know that it works, and if so, from what sort of evidence?
Best wishes Chris
----- Original Message ----- } From: "Malcolm Haswell" {malcolm.haswell-at-sunderland.ac.uk} To: "Microscopy MSA" {Microscopy-at-microscopy.com} Cc: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} Sent: Wednesday, September 15, 2004 3:20 PM
I'm curious about this, how was it used and what explosive was made. By the way Dynamite is not TNT. Dynamite is Nitroglycerin soaked into a diatomaceous earth called Kieselguhr (other absorbents have been used too), this is the explosive invented by Nobel that founded his munitions industry. Dynamite Nobel is still one of the worlds largest munitions makers. TNT is Trinitrotoluene, a different explosive, invented by Wilbrand. For more info on this see: http://en.wikipedia.org/wiki/TNT_(explosive) and http://en.wikipedia.org/wiki/Dynamite
James L. Roberts Firearm & Toolmark Examiner Ventura Co. Sheriff's Lab 800 S. Victoria Ave. Ventura, CA. 93009
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Alfred Nobel used Osmium in the production of Explosives in the 1860's. He is best known for discovering Nitroglycerin & Dynamite (TNT). Now of course his last will & testament funds "The Nobel Prize".
Al Coritz Electron Microscopy Sciences www.emsdiasum.com
----- Original Message ----- } From: {sbarlow-at-sunstroke.sdsu.edu} To: {microscopy-at-msa.microscopy.com} Sent: Monday, September 13, 2004 6:56 PM
Chris Molecular sieve usually used for removing water from the solvents like Et-OH etc. It has very high capacity and quite effective. Unfortunately (at my knowledge) molecular sieve does not have indicators signaling it's time to change sieve. As far as I know, if you add sieve in proportion 10% from volume, it should work effectively for couple of month (we are talking about adding molecular sieve to absolute 200 proof ethanol). You may regenerate molecular sieve at +200-250oC (24-48 h). The disadvantage of the sieve is that you may have particles in your sample (you right). There was posting on ListServer a few years ago that people used syringes with 0.22 mkm filters to remove particles. I don't remember details. In my experience, if I don't disturb solution, it's OK, no particles. Another popular desiccant is "anhydrous calcium sulfate" - this one do change the color and may be used for liquids or gases (for some reasons I did not hear that molecular sieve used for gases). All these materials are capable to remove the TRACES of water from your solution. If you have 10% water, you have to distill solvent first and then treat with few portions of desiccant. Personally, I am using 200 proof ethanol without any desiccants. I prefer to open new bottle if sample is important. Otherwise, I am using the bottle a few times as a 100% Et-OH and then use it for 30-95% dehydratation steps. Have a great day, Sergey
At 03:39 AM 9/15/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-080 Los Angeles, CA 90095
Yes it has to be wetted, but just wet the end and pour the sieve in with a funnel. Trim off the wet part and either tie or staple the ends. As for working, I also make a little packet of cupric sulfate the same way as an indicator and it turns blue indicating it has taken up moisture.
Dry it over a flame until it turns white. Recharge when blue. I generally need to do this once a year up here. all my 100% solutions (ethanol, acetone, and HMDS) are on sieve.
Scott Whittaker Laboratories of Analytical Biology Smithsonian Institution National Museum of Natural History PO Box 37012 MRC104 Washington DC 20013-7012 ***New Number202-633-0891***
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Malcolm Thanks for this.I have heard it suggested previously, but I am sceptical about it.
1) Presumably the dialysis tubing has to be wet before it can be loaded?. OK, so you can re-dry the MS after loading. 2) I wonder what the permeability of dialysis tubing is to water vapour when very dry and immersed in ethanol. It that environment it will be in its most ultra-compact precipitated state (since ethanol precipitates polysaccharides from aqueous solution). 3) As in my original question, do you really know that it works, and if so, from what sort of evidence?
Best wishes Chris
----- Original Message ----- } From: "Malcolm Haswell" {malcolm.haswell-at-sunderland.ac.uk} To: "Microscopy MSA" {Microscopy-at-microscopy.com} Cc: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} Sent: Wednesday, September 15, 2004 3:20 PM
We have been using molecular sieve in dialysis tubing for at least 20 years as a means of drying our 100% ethanol and acetone. Note that I am in Florida where the humidity is quite high for many months out of the year. Dry dialysis tubing can be opened, in order to fill it, by getting a small opening at one end and then blowing air through it to force it open all the way. A little blue indicator silica gel can be added for ethanol, but not acetone, since the dye is acetone soluble.
Greg Erdos, Hurricane Country
} Malcolm } Thanks for this.I have heard it suggested previously, but I am sceptical } about it. } } 1) Presumably the dialysis tubing has to be wet before it can be loaded?. } OK, so you can re-dry the MS after loading. } 2) I wonder what the permeability of dialysis tubing is to water vapour when } very dry and } immersed in ethanol. It that environment it will be in its most } ultra-compact precipitated state } (since ethanol precipitates polysaccharides from aqueous solution). } 3) As in my original question, do you really know that it works, and if so, } from what sort of evidence? } } Best wishes } Chris } } ----- Original Message ----- } } From: "Malcolm Haswell" {malcolm.haswell-at-sunderland.ac.uk} } To: "Microscopy MSA" {Microscopy-at-microscopy.com} } Cc: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} } Sent: Wednesday, September 15, 2004 3:20 PM } Subject: [Microscopy] Re: infiltration problems - suitable desiccants } } } } Chris } } } } to avoid the bits in your solvents, try putting your molecular sieve (or } whatever) into a length of dialysis tubing. You can fold the ends over and } staple them to form a small sausage. } } } } If you get any answers to the rest of your questions I, for one, would be } interested to hear them. } } } } Malcolm } } } } Malcolm Haswell } } e.m. unit } } School of Health, Natural and Social Sciences } } Fleming Building } } University of Sunderland } } Tyne & Wear } } SR1 3SD } } UK } } e-mail: malcolm.haswell-at-sunderland.ac.uk } } } } ----- Original Message ----- } } From: Chris Jeffree {c.jeffree-at-ed.ac.uk} } } Date: Wednesday, September 15, 2004 11:39 am } } Subject: [Microscopy] infiltration problems - suitable desiccants } } } } } } } } } } } ------------------------------------------------------------------- } } } ----------- } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } } } AmericaTo Subscribe/Unsubscribe -- } } } http://www.msa.microscopy.com/MicroscopyListserverOn-Line Help } } } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html-------- } } } ------------------------------------------------------------------- } } } ---- } } } } } } How effective is molecular sieve as a desiccant at typical } } } polymerizationtemperatures (60-70 oC)?? I suspect not very. Would } } } there be a better } } } choice? } } } } } } I would also be interested to know what strategies people now } } } recommend for } } } drying EM solvents } } } without getting them contaminated with bits of desiccant. } } } } } } And another thing - how on earth can one determine whether the } } } desiccant is } } } still being effective? } } } Can you trust the blue indicator in molecular sieves? What is the } } } equilibrium water concentration in ethanol above fresh molecular sieve } } } (assuming the ethanol contained some water initially), and does this } } } increase when the MS contains, say, 10% water (half its saturation } } } capacity)? } } } } } } Best wishes } } } Chris } } } } } } } } } } } } }
Gregory W. Erdos Ph.D. Assistant Director, Biotechnology Program Scientific Director, Electron Microscopy P.O. Box 118525 217 Carr Hall University of Florida Gainesville, FL 32611 gwe-at-ufl.edu 352-392-1295 fax- 352-846-0251
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 13:10:43 2004
I use acetone for my dehydrations (prior to SPurr's infiltration). My professor taught me years ago (and I teach my students now) to dehydrate acetone with CuSO4. Place about 50-100g of CuSO4 in an evaporating dish in a muffle furnace for about 6 hrs. The deep aqua crystals will turn to a white powder with a faint greenish cast. Allow to cool slightly (in the furnace), add to an empty bottle, add "100%" acetone from a freshly opened 500 mL bottle, shake, and allow to settle over night. As long as the copper sulfate does not change color, the acetone is assumed to be adequately desiccated. I've used this with complete success for acetone, with not apparent affect on specimens from the copper. I have no idea whether it works for EtOH.
Don
______________________________________________________________________ Donald L. Lovett e-mail: lovett-at-tcnj.edu Assoc. Professor, Dept. of Biology voice: (609) 771-2876 P.O. Box 7718 fax: (609) 637-5118 The College of New Jersey Ewing, NJ 08628-0718
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 13:39:19 2004
Hello Listers I have been reading the replies with great interest because I too have been struggling with incomplete dehydration/poor infiltration problems with 7 day zebrafish brain in very dry California. I am using a combination of microwave and bench processing. My dehydration steps are done under vacuum on the stedi-temp in the microwave(power level 3, 40 seconds each 100% 1 minute each)and I do 100% ethanol 4x (bottle used only 3 times before new one is opened, no molecular sieves.) Moving to 100% acetone and sometimes even PPO and infiltrating with acetone -resin mixture or PPO/resin. I include 1 overnight step, in 50-50 on rotator, then 100% for several hours on rotator. I have tried epon/araldite (frozen with accelerator-made fresh every 3 months) and recently Embed It (Polysciences Spurr like formulation). I have white holes or empty areas -sometimes myelin like figures. I have looked at conventionally prepared tissue that doesn't have it and was dehydrated for longer periods so I think it is a dehydration problem. The microwave fixation is far superior to the bench fixed material so I don't think it can be blamed on the microwave. I have just ordered new acetone in smaller bottles thinking that maybe my acetone had taken on water. I process in dram glass vials and always keep the tissue covered. It is only this darn fish that gives me this trouble. I.have done mouse brain, Drosophila, rat heart, 5 day fish all using this protocol with great results. There must be water somewhere but it is hard to track down. This fish brain is very dense material, along with the skin around it. I dissect away the eyes and jaw to improve penetration. Is there anyone out there who has seen this problem before? I'd love some advice. Frustrated in California, JoAnn Buchanan
Department of Molecular and Cellular Physiology Stanford University School of Medicine Stanford, CA 94305 650-723-5856
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 13:43:13 2004
On the subject of humidity causing poor infiltration ....... several of my colleagues go from 95% ethanol to Epon substitutes without any problems. No absolute alcohol, no propylene oxide. Yes, extra changes of epoxy are needed but the results are fine. Hayat's Prin. and Tech of EM, second ed., vol 1, page 154 reports that Epon is miscible with 70% ethanol. I was "raised" with the "you must get every last molecule of water out of the specimen" dogma but eperience had taught me otherwise. I suggest you look elsewhere for the cause of your difficulties.
Geoff
Chris Jeffree wrote:
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-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 13:56:24 2004
I want to buy a grinder/polisher that works with Gatan 623 Disc Grinder or SPI Precision Disc Grinder. Does anyone use "8” Grinder/Polisher" from Electron Microscopy Sciences with Disc Grinder? Is there any other recommendation? I would find a grinder around 1-2K. Thank you, Hiromi Konishi Indiana University
I've always for the last 20 years used digital balances to measure out quantities of resin components to mix up plastics, but I'm curious as to why the manufacturers always specify the amount in volume measures instead of weight, since it seems to me that it would be a tough job to clean out grad cylinders full of resin components, vs just filling up a plastic beaker on a balance, and disposing of it afterwards.
I was just curious as to how others do their measuring, and if they resort to the horrifying task of cleaning glassware of resin components, which strikes me as a particularly nasty job.
Garry Burgess
Charge Technologist Department of Pathology Health Sciences Centre Winnipeg, Canada
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 14:25:01 2004
What, specifically, are you trying to image in these samples? It may be that TEM is not the right tool. An associate of mine has been imaging LB and polymer films with AFM and AFAM (Atomic Force Acoustical Microscopy), with great success and no damage. Contact me off-line if you are interested in details.
Thanks Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^& Need a good general text on light microscopy? MME still has copies of Optimizing Light Microscopy available, with discounts for class-sized orders (10 or more). Visit www.MicroscopyEducation.com for details.
At 09:07 AM 9/15/2004, tem_iopb-at-iopb.res.in wrote:
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From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 16:40:37 2004
Garry, we measure in volume equivalents, one ml per gram. Although I recognize that the differing specific gravities of the various components means that these are not exactly equivalent measures, it works well for us.
We use 30 ml syringes sans needles to measure the components---pop out the plunger (and make sure the syringe cap is on!) and fill the body of the syringe up to the appropriate mark. Reinsert the plunger and push out the goo into a plastic mixing cup. Repeat for the other components. Not only is this quick, easy, and about as mess free as working with resins ever gets, but you get some interesting sound effects that raise eyebrows on people not in the know. As an added bonus, if you pour the mixed resin back in the syringes, they make for nice freezer storage and easy dispensing. When empty, just throw them into an oven until the remnants are polymerized and toss the used syringes.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
-----Original Message----- } From: Garry Burgess [mailto:GBurgess-at-exchange.hsc.mb.ca] Sent: Wednesday, September 15, 2004 2:38 PM To: microscopy-at-microscopy.com
I've always for the last 20 years used digital balances to measure out quantities of resin components to mix up plastics, but I'm curious as to why the manufacturers always specify the amount in volume measures instead of weight, since it seems to me that it would be a tough job to clean out grad cylinders full of resin components, vs just filling up a plastic beaker on a balance, and disposing of it afterwards.
I was just curious as to how others do their measuring, and if they resort to the horrifying task of cleaning glassware of resin components, which strikes me as a particularly nasty job.
Garry Burgess
Charge Technologist Department of Pathology Health Sciences Centre Winnipeg, Canada
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 16:56:07 2004
Dear Hiromi, Because the amount of material removed from 3.05 mm TEM discs with the Gatan Disc Grinder is very small, I use 600 grit paper stuck to a glass sheet and carefully hand grind the samples under running water, then finish off with 1200 grit the same way. It only takes a few strokes to remove 50 microns, then advance the dial and remove another 50 microns. We have spinning wheel grinders and automatic polishers, but I would not use them for this. Regards, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: {hkonishi-at-indiana.edu} To: {Microscopy-at-microscopy.com} Sent: Wednesday, September 15, 2004 12:23 PM
Dear Garry, I found that 1 oz. medicine cups, available from medical suppliers in 1000 cup batches, are disposable and marked off in ml. They hold about 30 ml and do one or two 1-inch mounts. I use the Tri-pour plastic beakers, which my Stores man assures me are inexpensive, for larger amounts. I have also heard that some people use disposable syringes. Regards, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: "Garry Burgess" {GBurgess-at-exchange.hsc.mb.ca} To: {microscopy-at-microscopy.com} Sent: Wednesday, September 15, 2004 12:37 PM
In dense plant tissues, we generally get problems in embedded tissues not because of incomplete dehydration, but due to infiltration either too fast or in steps that are too large. If cell walls are moderately impermeable, then solvent may diffuse out of the tissue faster than resin can diffuse in, causing tissue collapse and "airspaces" when remaining traces of solvent within such tissue vapourise during polymerisation. Slower infiltration, with smaller increases of % resin in solvent, and holding the tissue longer at each step, has generally solved this problem, for us, anyway.
As an example, a friend found that the single- to few-celled algal zygotes she was working on had to be infiltrated in increments of 1-2% resin per day up to 10% or so, after which the increments could be larger, then above 90% resin, the increments had to be smaller again (don't remember all the details). If this wasn't done, the zygotes looked like flat balloons after resin polymerisation.
Not sure if this applies to animal tissues but is another protocol modification to try.... cheers, Rosemary
Rosemary White rosemary.white-at-csiro.au Microscopy Centre ph. 02-6246 5475 CSIRO Plant Industry mob. 0402 835 973 GPO Box 1600 fax. 02-6246 5000 Canberra, ACT 2601
} From: JoAnn Buchanan {redhair-at-stanford.edu} } Date: Wed, 15 Sep 2004 12:06:49 -0700 } To: {Microscopy-at-msa.microscopy.com} } Subject: [Microscopy] Infiltration Problems } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } Hello Listers } I have been reading the replies with great interest because I too } have been struggling with incomplete dehydration/poor infiltration problems } with 7 day zebrafish brain in very dry California. I am using a combination } of microwave and bench processing. My dehydration steps are done under } vacuum on the stedi-temp in the microwave(power level 3, 40 seconds each } 100% 1 minute each)and I do 100% ethanol 4x (bottle used only 3 times } before new one is opened, no molecular sieves.) } Moving to 100% acetone and sometimes even PPO and infiltrating with acetone } -resin mixture or PPO/resin. I include 1 overnight step, in 50-50 on } rotator, then 100% for several hours on rotator. I have tried epon/araldite } (frozen with accelerator-made fresh every 3 months) and recently Embed It } (Polysciences Spurr like formulation). } I have white holes or empty areas -sometimes myelin like figures. I have } looked at conventionally prepared tissue that doesn't have it and was } dehydrated for longer periods so I think it is a dehydration problem. The } microwave fixation is far superior to the bench fixed material so I don't } think it can be blamed on the microwave. I have just ordered new acetone in } smaller bottles thinking that maybe my acetone had taken on water. I } process in dram glass vials and always keep the tissue covered. } It is only this darn fish that gives me this trouble. I.have done mouse } brain, Drosophila, rat heart, 5 day fish all using this protocol with great } results. There must be water somewhere but it is hard to track down. This } fish brain is very dense material, along with the skin around it. I dissect } away the eyes and jaw to improve penetration. } Is there anyone out there who has seen this problem before? I'd love some } advice. } Frustrated in California, JoAnn Buchanan } } Department of Molecular and Cellular Physiology } Stanford University School of Medicine } Stanford, CA 94305 } 650-723-5856 } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 17:58:07 2004
We have used weight measurement for many years. We have a scale in a hood and use gloves and all disposable containers so dare not too concerned about the resin components being hazardous. We mix resins as needed rather than mix large batches and freeze aliquots since it only takes a few minutes to mix the resins in the quantity we need. This works fine and we get consistent resin properties in our blocks.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building 170 S. University Street West Lafayette, IN 47907 http://www3.agriculture.purdue.edu/microscopy
On 9/15/04 2:37 PM, "Garry Burgess" {GBurgess-at-exchange.hsc.mb.ca} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } } I've always for the last 20 years used digital balances to measure out } quantities of resin components to mix up plastics, but I'm curious as to why } the manufacturers always specify the amount in volume measures instead of } weight, since it seems to me that it would be a tough job to clean out grad } cylinders full of resin components, vs just filling up a plastic beaker on a } balance, and disposing of it afterwards. } } I was just curious as to how others do their measuring, and if they resort } to the horrifying task of cleaning glassware of resin components, which } strikes me as a particularly nasty job. } } Garry Burgess } } Charge Technologist } Department of Pathology } Health Sciences Centre } Winnipeg, Canada } } This e-mail and/or any documents in this transmission is intended for the } address(s) only and may contain legally privileged or confidential } information. Any unauthorized use, disclosure, distribution, copying or } dissemination is strictly prohibited. If you receive this transmission in } error, please notify the sender immediately and return the original. }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 19:55:36 2004
The EMS Polisher you reference would certainly be suitable for use with the disc grinder. Mary Mager makes a good point that a rotating wheel may not be necessary - or even desirable - if you don't have a lot of material to remove. I can offer a few suggestions:
Rotating Wheel If you do require a rotating wheel, our Model 900 Grinder/Polisher is an enconomical solution that would be suitable for the disc grinder you describe.
Hand Polishing If you decide to go with a manual hand polishing process, I would suggest our Model 180 Lapping Tray. This is essentially a glass plate mounted in an aluminum base. It is set up to collect waste water and debris and comes with a cover that allows you to stack multiple trays each with a different grit size abrasive.
We also have a wide range of other polishing tools and supplies that may be of interest. I would be pleased to discuss your application with you in detail off-line.
DISCLAIMER: South Bay Technology produces equipment and supplies as described above and, therefore, has a vested interest in promoting their use.
Best regards-
David
hkonishi-at-indiana.edu wrote:
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From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 20:34:08 2004
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Dear Hiromi, Because the amount of material removed from 3.05 mm TEM discs with the Gatan Disc Grinder is very small, I use 600 grit paper stuck to a glass sheet and carefully hand grind the samples under running water, then finish off with 1200 grit the same way. It only takes a few strokes to remove 50 microns, then advance the dial and remove another 50 microns. We have spinning wheel grinders and automatic polishers, but I would not use them for this. Regards, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: {hkonishi-at-indiana.edu} To: {Microscopy-at-microscopy.com} Sent: Wednesday, September 15, 2004 12:23 PM
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Garry, we measure in volume equivalents, one ml per gram. Although I recognize that the differing specific gravities of the various components means that these are not exactly equivalent measures, it works well for us.
We use 30 ml syringes sans needles to measure the components---pop out the plunger (and make sure the syringe cap is on!) and fill the body of the syringe up to the appropriate mark. Reinsert the plunger and push out the goo into a plastic mixing cup. Repeat for the other components. Not only is this quick, easy, and about as mess free as working with resins ever gets, but you get some interesting sound effects that raise eyebrows on people not in the know. As an added bonus, if you pour the mixed resin back in the syringes, they make for nice freezer storage and easy dispensing. When empty, just throw them into an oven until the remnants are polymerized and toss the used syringes.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
-----Original Message----- } From: Garry Burgess [mailto:GBurgess-at-exchange.hsc.mb.ca] Sent: Wednesday, September 15, 2004 2:38 PM To: microscopy-at-microscopy.com
I've always for the last 20 years used digital balances to measure out quantities of resin components to mix up plastics, but I'm curious as to why the manufacturers always specify the amount in volume measures instead of weight, since it seems to me that it would be a tough job to clean out grad cylinders full of resin components, vs just filling up a plastic beaker on a balance, and disposing of it afterwards.
I was just curious as to how others do their measuring, and if they resort to the horrifying task of cleaning glassware of resin components, which strikes me as a particularly nasty job.
Garry Burgess
Charge Technologist Department of Pathology Health Sciences Centre Winnipeg, Canada
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 20:34:25 2004
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Garry
We have used weight measurement for many years. We have a scale in a hood and use gloves and all disposable containers so dare not too concerned about the resin components being hazardous. We mix resins as needed rather than mix large batches and freeze aliquots since it only takes a few minutes to mix the resins in the quantity we need. This works fine and we get consistent resin properties in our blocks.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building 170 S. University Street West Lafayette, IN 47907 http://www3.agriculture.purdue.edu/microscopy
On 9/15/04 2:37 PM, "Garry Burgess" {GBurgess-at-exchange.hsc.mb.ca} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } } I've always for the last 20 years used digital balances to measure out } quantities of resin components to mix up plastics, but I'm curious as to why } the manufacturers always specify the amount in volume measures instead of } weight, since it seems to me that it would be a tough job to clean out grad } cylinders full of resin components, vs just filling up a plastic beaker on a } balance, and disposing of it afterwards. } } I was just curious as to how others do their measuring, and if they resort } to the horrifying task of cleaning glassware of resin components, which } strikes me as a particularly nasty job. } } Garry Burgess } } Charge Technologist } Department of Pathology } Health Sciences Centre } Winnipeg, Canada } } This e-mail and/or any documents in this transmission is intended for the } address(s) only and may contain legally privileged or confidential } information. Any unauthorized use, disclosure, distribution, copying or } dissemination is strictly prohibited. If you receive this transmission in } error, please notify the sender immediately and return the original. }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 20:34:19 2004
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In dense plant tissues, we generally get problems in embedded tissues not because of incomplete dehydration, but due to infiltration either too fast or in steps that are too large. If cell walls are moderately impermeable, then solvent may diffuse out of the tissue faster than resin can diffuse in, causing tissue collapse and "airspaces" when remaining traces of solvent within such tissue vapourise during polymerisation. Slower infiltration, with smaller increases of % resin in solvent, and holding the tissue longer at each step, has generally solved this problem, for us, anyway.
As an example, a friend found that the single- to few-celled algal zygotes she was working on had to be infiltrated in increments of 1-2% resin per day up to 10% or so, after which the increments could be larger, then above 90% resin, the increments had to be smaller again (don't remember all the details). If this wasn't done, the zygotes looked like flat balloons after resin polymerisation.
Not sure if this applies to animal tissues but is another protocol modification to try.... cheers, Rosemary
Rosemary White rosemary.white-at-csiro.au Microscopy Centre ph. 02-6246 5475 CSIRO Plant Industry mob. 0402 835 973 GPO Box 1600 fax. 02-6246 5000 Canberra, ACT 2601
} From: JoAnn Buchanan {redhair-at-stanford.edu} } Date: Wed, 15 Sep 2004 12:06:49 -0700 } To: {Microscopy-at-msa.microscopy.com} } Subject: [Microscopy] Infiltration Problems } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } Hello Listers } I have been reading the replies with great interest because I too } have been struggling with incomplete dehydration/poor infiltration problems } with 7 day zebrafish brain in very dry California. I am using a combination } of microwave and bench processing. My dehydration steps are done under } vacuum on the stedi-temp in the microwave(power level 3, 40 seconds each } 100% 1 minute each)and I do 100% ethanol 4x (bottle used only 3 times } before new one is opened, no molecular sieves.) } Moving to 100% acetone and sometimes even PPO and infiltrating with acetone } -resin mixture or PPO/resin. I include 1 overnight step, in 50-50 on } rotator, then 100% for several hours on rotator. I have tried epon/araldite } (frozen with accelerator-made fresh every 3 months) and recently Embed It } (Polysciences Spurr like formulation). } I have white holes or empty areas -sometimes myelin like figures. I have } looked at conventionally prepared tissue that doesn't have it and was } dehydrated for longer periods so I think it is a dehydration problem. The } microwave fixation is far superior to the bench fixed material so I don't } think it can be blamed on the microwave. I have just ordered new acetone in } smaller bottles thinking that maybe my acetone had taken on water. I } process in dram glass vials and always keep the tissue covered. } It is only this darn fish that gives me this trouble. I.have done mouse } brain, Drosophila, rat heart, 5 day fish all using this protocol with great } results. There must be water somewhere but it is hard to track down. This } fish brain is very dense material, along with the skin around it. I dissect } away the eyes and jaw to improve penetration. } Is there anyone out there who has seen this problem before? I'd love some } advice. } Frustrated in California, JoAnn Buchanan } } Department of Molecular and Cellular Physiology } Stanford University School of Medicine } Stanford, CA 94305 } 650-723-5856 } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 20:41:51 2004
We have always measured our Spurr's resin components by weight, most of the recipes for this resin seem to do this. I agree most of the literature on other resins seems to quote volumes though in the latest edition of Glauerts book on Biological Specimen Preparation weight equivalents are given (proportions are close to but not identical to those using 1g = 1ml)
The reference is Biological Specimen Preparation for Transmission Electron Microscopy - Audrey M. Glauert and Peter R. Lewis Practical Methods in Electron Microscopy: Vol 17 Portland Press London 1998
The earlier version of this (1974) only seems to note volumes.
Ian
Ian Hallett HortResearch Mt Albert Research Centre, Private Bag 92 169 Auckland, New Zealand Fax +64 9 815 4201 Telephone +64 9 815 4200 EMail ihallett-at-hortresearch.co.nz
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I've always for the last 20 years used digital balances to measure out quantities of resin components to mix up plastics, but I'm curious as to why the manufacturers always specify the amount in volume measures instead of weight, since it seems to me that it would be a tough job to clean out grad cylinders full of resin components, vs just filling up a plastic beaker on a balance, and disposing of it afterwards.
I was just curious as to how others do their measuring, and if they resort to the horrifying task of cleaning glassware of resin components, which strikes me as a particularly nasty job.
Garry Burgess
Charge Technologist Department of Pathology Health Sciences Centre Winnipeg, Canada
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
The contents of this e-mail are privileged and/or confidential to the named recipient and are not to be used by any other person and/or organisation. If you have received this e-mail in error, please notify the sender and delete all material pertaining to this e-mail. ______________________________________________________
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 23:12:08 2004
It works perfectly with Et-OH and this is exactly how they teach me. Yes, CuSO4 - it was many years ago. Thanks for reminding. Sergey
At 11:37 AM 9/15/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-080 Los Angeles, CA 90095
Chris, you right. I am sorry, I was surely believe that PPO does not soluble in the water. So, my theory that PPO may "protect" sample from the water is wrong. From another hand I never used desiccants with PPO. PPO comes in 0.5 liter cans and I opened it frequently. Nevertheless, I did not have problems with water. Anyway, thanks for clarification and have a great day/night. Sergey
At 02:06 AM 9/15/2004, you wrote: } Sergey } A propylene oxide layer over the specimen cannot protect against water } penetration. } Water's solubility in 1,2 propylene oxide (=1,2 epoxypropane) is 14.7 wt%. } Conversely, PPO's solubility in water is 40.5 wt%. } } Chris } } ----- Original Message ----- } From: "Sergey Ryazantsev" {sryazant-at-ucla.edu} } To: {Microscopy-at-microscopy.com} } Sent: Tuesday, September 14, 2004 8:18 PM } Subject: [Microscopy] Re: RE: Infiltration Problems } } } } } } } } -------------------------------------------------------------------------- } ---- } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -------------------------------------------------------------------------- } ----- } } } } I don't think water may penetrate epoxy resin easily. The picture, you } } described is more like you have water in your sample before plastic. As } } far as I could see, the 100% Et-OH step may be a problem in your recipe. } 4x } } 100% Et-OH sounds excessive (I always do 2x20 min 100% Et-OH) - so to me } it } } looks like that your 100% Et-ON is not a 100% (if you still have water } } after 4x). You may need to change the batch or use molecular sieve. You } } need also to use at least 1 ml of Et-OH per sample. It's better to do it } } in hermetic vials like 1.5 ml Eppendorf tube. If I forgot Et-OH bottle to } } close immediately after use, I consider it's 95% Et-OH and use it for } } 30-95% dehydratation steps only. 3x of PO is sounds excessive too. The } } trick here is that as more changes you do, as more your sample has exposed } } to water (when you emptied the vial). So, 1 exchange of PO (and Et-OH) is } } much better than 3 (you still need 2 for Et-OH) in my point of view. You } } need to use huge excess of the solvents - at least 1 ml per sample. As } soon } } as your sample in PO (or epoxy) - it's protected from water by the layer } of } } solvent (epoxy). PO and epoxy do not dissolve water, therefore it could } } not reach your sample. You still may have a droplets of water (from } } condensation for instance) but it'll generate a different picture: round } } empty spaces in your sample. You may try also Spurr resin - it may } } tolerate a few %% of water in your sample. I hope it'll help, Sergey } } } } At 06:38 AM 9/14/2004, you wrote: } } } } } } } } --------------------------------------------------------------------------- } --- } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } } To Subscribe/Unsubscribe -- } } } http://www.msa.microscopy.com/MicroscopyListserver } } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } } --------------------------------------------------------------------------- } ---- } } } } } } } } } } } It seems that during the summer months, we have no end of } } } } troubles with our plastic not infiltrating the tissue. I've } } } } always assumed that this was because of the humidity rising } } } } quite a bit during the summer months, and the poor air conditioning. } } } } } } } } It comes out as sections falling apart when I try to flatten } } } } them in the water boat, or breaking in the electron beam. We } } } } have always kept our epoxy resin in the freezer in 30 ml } } } } plastic syringes, but it usually only gives us a problem } } } } during the summer months. During the winter here, the air is } } } } extremely dry. } } } } } } } } We do 4 rinses in absolute alcohol, followed by 3 rinses in } } } } propylene oxide, followed by a 50/50 mixture of epon/araldite } } } } mixed with propylene oxide over night with the caps off, for } } } } the propylene oxide evaporating overnight and slowly leaving } } } } the specimens in 100% plastic by morning. We polymerize at } } } } 70 deg C overnight, keeping a dessicant in our embedding oven } } } } to keep the air as dry as possible too. } } } } } } Sergey Ryazantsev Ph. D. } } Electron Microscopy } } UCLA School of Medicine } } Department of Biological Chemistry } } 10833 Le Conte Ave, Room 33-080 } } Los Angeles, CA 90095 } } } } Phone: (310) 825-1144 (office) } } (310) 206-1029 (Lab) } } FAX (departmental): (310) 206-5272 } } mailto:sryazant-at-ucla.edu } } } } } } } } } }
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-080 Los Angeles, CA 90095
I’m working with some extremely fastidious dinoflagelates, which I can’t seam to keep the flagella in place. I’m after the flagella attachment of the Pyrodinium Bahamense. On a separate batch I’m looking to break the armor to take a look inside. I have modified the dehydration process in order to see if that preserve the flagella. I’m also thinking to change the fixative from Paraformaldehyde to Glutaraldehyde. Any ideas into how to treat these organisms will be greatly appreciated.
Omayra Velez New Jersey
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From MicroscopyL-request-at-ns.microscopy.com Wed Sep 15 23:34:33 2004
I'm sorry for the previous email subject heading. It didnot make any sense. Originaly, I had an osmium question, about the dinoflagellates but I found the answer myself.
Omayra Velez New Jersey
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From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 08:01:18 2004
Just to add to you potential choices Ladd offers our L900 8" Grinder/Polisher.
JD Arnott
Disclaimer: Ladd Research is in the business of supplying your microscopy needs including the products mentioned in this thread.
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: ladres-at-att.net
----- Original Message ----- } From: {hkonishi-at-indiana.edu} To: {Microscopy-at-microscopy.com} Sent: Wednesday, September 15, 2004 3:23 PM
I want to buy a grinder/polisher that works with Gatan 623 Disc Grinder or SPI Precision Disc Grinder. Does anyone use "8" Grinder/Polisher" from Electron Microscopy Sciences with Disc Grinder? Is there any other recommendation? I would find a grinder around 1-2K. Thank you, Hiromi Konishi Indiana University
It's not hard to measure volumes when you use a syringe.
Mannie Steglich UT MDACC
Garry Burgess {GBurgess-at-exchange.hsc.mb.ca}
09/15/2004 02:37 PM
To: {microscopy-at-microscopy.com} cc:
I've always for the last 20 years used digital balances to measure out quantities of resin components to mix up plastics, but I'm curious as to why the manufacturers always specify the amount in volume measures instead of weight, since it seems to me that it would be a tough job to clean out grad cylinders full of resin components, vs just filling up a plastic beaker on a balance, and disposing of it afterwards.
I was just curious as to how others do their measuring, and if they resort to the horrifying task of cleaning glassware of resin components, which strikes me as a particularly nasty job.
Garry Burgess
Charge Technologist Department of Pathology Health Sciences Centre Winnipeg, Canada
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 08:44:56 2004
I only freeze a small amount and only use the stored frozen aliquots for the intermediate stages: the mixes with propylene oxide. I always use freshly mixed resin for embedding.
Peggy
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Debby Sherman [mailto:dsherman-at-purdue.edu] Sent: Wednesday, September 15, 2004 7:25 PM To: Garry Burgess; microscopy-at-microscopy.com
Garry
We have used weight measurement for many years. We have a scale in a hood and use gloves and all disposable containers so dare not too concerned about the resin components being hazardous. We mix resins as needed rather than mix large batches and freeze aliquots since it only takes a few minutes to mix the resins in the quantity we need. This works fine and we get consistent resin properties in our blocks.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building 170 S. University Street West Lafayette, IN 47907 http://www3.agriculture.purdue.edu/microscopy
On 9/15/04 2:37 PM, "Garry Burgess" {GBurgess-at-exchange.hsc.mb.ca} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } } I've always for the last 20 years used digital balances to measure out } quantities of resin components to mix up plastics, but I'm curious as to why } the manufacturers always specify the amount in volume measures instead of } weight, since it seems to me that it would be a tough job to clean out grad } cylinders full of resin components, vs just filling up a plastic beaker on a } balance, and disposing of it afterwards. } } I was just curious as to how others do their measuring, and if they resort } to the horrifying task of cleaning glassware of resin components, which } strikes me as a particularly nasty job. } } Garry Burgess } } Charge Technologist } Department of Pathology } Health Sciences Centre } Winnipeg, Canada } } This e-mail and/or any documents in this transmission is intended for the } address(s) only and may contain legally privileged or confidential } information. Any unauthorized use, disclosure, distribution, copying or } dissemination is strictly prohibited. If you receive this transmission in } error, please notify the sender immediately and return the original. }
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 08:51:56 2004
I prepare my resin exactly the same way, Randy, and have not had any problems. Have raised a few eyebrows with the sound effects also! I store small aliquots of resin in scintillation vials in the freezer. I don't process a high volume of tissue, so for me this method works well. I just thaw a vial and use a plastic disposable pipet to fill my molds. No fuss, no mess.
Peggy
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Tindall, Randy D. [mailto:TindallR-at-missouri.edu] Sent: Wednesday, September 15, 2004 6:07 PM To: Garry Burgess Cc: microscopy-at-microscopy.com
Garry, we measure in volume equivalents, one ml per gram. Although I recognize that the differing specific gravities of the various components means that these are not exactly equivalent measures, it works well for us.
We use 30 ml syringes sans needles to measure the components---pop out the plunger (and make sure the syringe cap is on!) and fill the body of the syringe up to the appropriate mark. Reinsert the plunger and push out the goo into a plastic mixing cup. Repeat for the other components. Not only is this quick, easy, and about as mess free as working with resins ever gets, but you get some interesting sound effects that raise eyebrows on people not in the know. As an added bonus, if you pour the mixed resin back in the syringes, they make for nice freezer storage and easy dispensing. When empty, just throw them into an oven until the remnants are polymerized and toss the used syringes.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
-----Original Message----- } From: Garry Burgess [mailto:GBurgess-at-exchange.hsc.mb.ca] Sent: Wednesday, September 15, 2004 2:38 PM To: microscopy-at-microscopy.com
I've always for the last 20 years used digital balances to measure out quantities of resin components to mix up plastics, but I'm curious as to why the manufacturers always specify the amount in volume measures instead of weight, since it seems to me that it would be a tough job to clean out grad cylinders full of resin components, vs just filling up a plastic beaker on a balance, and disposing of it afterwards.
I was just curious as to how others do their measuring, and if they resort to the horrifying task of cleaning glassware of resin components, which strikes me as a particularly nasty job.
Garry Burgess
Charge Technologist Department of Pathology Health Sciences Centre Winnipeg, Canada
This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 08:54:25 2004
Dinoflagellates are not easy to fix properly. My experience with them has been that each species (especially the lightly armored or unarmored ones)can have their own individual reqirements for fixation. When I have problems with fixation I use what has been called Parducz fixative. It was originally developed in the 1960's for work on cilliates by Bela Parducz to study the metachronal wave pattern of the cillia. It consists of a mixture of Osmium tetroxide and mercuric chloride. To prepare the solution according to Parducz, you make a 6:1(V:V) mixture of 2% aqueous Osmium tetroxide(OsO4) with saturated aqueous mercuric chloride(HgCl2). You can adjust the relative mixtures of the two components as I know that I have used 3:2 and perhaps 1:1. This fixative has been referred to as an "instantaneous fix" as it will fix the specimens very, very, rapidly. Parducz fixative was the only way that I could prepare some dinoflagellates for SEM and retain not only their morphology, but flagella as well. It is not as good a fix for TEM. Here are a couple of references that you may want to see:
Small, E.B. 1968. Scanning electron microscopy of fixed, frozen and dried } protozoa. Science. 163:1064-1065. } } and } } Bela Parducz. 1966. Ciliary Movement abd Coordination in Cilliates. } International Review of Cytology. 21:91-128.
Omayra Velez wrote:
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-- -- ================================================================== Greg Strout Electron Microscopist, University of Oklahoma WWW Virtual Library for Microscopy: http://www.ou.edu/research/electron/www-vl/ e-mail: gstrout-at-ou.edu Opinions expressed herein are mine and not necessarily those of the University of Oklahoma ==================================================================
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 09:10:45 2004
The July/August issue of PhotoTechniquesUSA has a review of the HP Photosmart 7960 printer. The reviewer, a professional photographer and printmaker named Ctein, said "it makes the best (and most permanent) black and white prints of any computer printer I've ever tried." It is an 8 ink printer that has 3 photo gray inks in addition to the usual color inks although it seems that the photo gray ink cartridge is an extra cost option? The magazine has a website http://www.phototechmag.com/ but I do not know if the review is available there.
Geoff
Gary Gaugler wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } I was a fan of Epson photo printers for quite } some time. Notable was the 890 & 980. It is a } small format printer compared to the 2200. } I recently (a year or so ago) bought a Epson Stylus } Photo 2000. It lasted about six months and } then jammed constantly. In-warranty customer } service and any idea of repair was on a wish list. } It never happened. The printer was scrapped. } } The 2200 may have solved teething problems } with large format printers. However, the 2000 } was VERY slooooow using photo paper. When it } worked, the results were stunning. Many times } (too many) it would stop printing 1/4 or 1/2 } way through the print and just die. The job } hung (Win2K Pro) and had to be restarted with } a new sheet of paper. } } The Epson and Canon small format printers seem } to do a better, more reliable job. As a result } of being burned by Epson, I now take print jobs } to a local service bureau. they do a very nice } job for not much cost. These are mostly for } 24" x 48" glossy mounted prints. Small ones } are done on my HP 4550 color laser printer. } If the color gamut is matched well between } the monitor and Photoshop, the HP does a nice job } for reports. For transparencies (not much used } any longer), the Kodak dye sub is excellent. } } Let us know what you find. There are a lot } of options. Also, check out the Ethernet print } servers that will connect a non-network printer } to a LAN and allow all to use it. HP and others } make these. they usually cost about $100 or so. } } gary g. } } } At 11:08 AM 9/2/2004, you wrote: } } } Email: jtd1-at-psu.edu } } Name: Tom Doman } } } } Organization: Penn State University } } } } Title-Subject: [Microscopy] [Filtered] MListserver: } } } } Question: Our lab is considering various photo printers (} $1K) for } } production of electron micrographs. Currently the Epson 2200 is the } } stromg favorite. Are there any recomendations for other printers } } which we should consider? What are your reasons for the recomended } } printer? } } } } Thanks in advance! } } } } Tom } } } } --------------------------------------------------------------------------- } } } } }
-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 10:44:57 2004
i wonder if the following is relevant - narcotize the critters before fixing them. see, for example:
May L. (1985) The use of procaine hydrochloride in the preparation of rotifer samples for counting. Verh.Internat.Verein.Limnol. 22, 2987-2990.
Abstract: Although many rotifer species are easily recognized when killed and fixed in formalin, some of the soft-bodied forms contract violently on contact with the chemical and become unrecognizable. In samples from Loch Leven this situation occurred most commonly with Synchaeta kitina Rousselet and several possible methods of killing the species in a relaxed from were considered (May, 1980). Narcotizing the animals with procaine hydrochloride (NH2.C6H4.COO.CH2.CH2.N(C2H5)2.HCl) before adding formalin was found to be the most succesful method. This paper describes the development of the method and examines its effect on rotifer density estimates
Chris
----- Original Message ----- } From: "Greg Strout" {gstrout-at-ou.edu} To: "Omayra Velez" {mayas003-at-yahoo.com} Cc: {Microscopy-at-MSA.Microscopy.com} Sent: Thursday, September 16, 2004 3:21 PM
I am looking for a "micrometer" that can work with optical microscope. I think that it read the difference of dial for focusing and convert to actual distance (um). I want to use it for determining the thickness of disc during disc grinding if it is not so expensive. Please advise on website or product information. Thank you,
Hiromi Konishi Indiana University
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 11:06:46 2004
I own 2 of these printers and they actually come with all 4 cartridges, color, photo, black & grey. The unit has a storage compartment for the cartridge not in use (either the grey or the black). I agree with the review it does an outstanding job of both color and B&W. When printing at its highest resolution an 8X10 print can take 11 minutes to print and will need (temporarly) 400 Mb of hard disk space. When I got mine the were selling at 300 USD, they are now selling at about 200 USD.
Love 'em Robert Schoonhoven
Geoff McAuliffe wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } Dear List: } } The July/August issue of PhotoTechniquesUSA has a review of the HP } Photosmart 7960 printer. The reviewer, a professional photographer and } printmaker named Ctein, said "it makes the best (and most permanent) } black and white prints of any computer printer I've ever tried." It is } an 8 ink printer that has 3 photo gray inks in addition to the usual } color inks although it seems that the photo gray ink cartridge is an } extra cost option? } The magazine has a website http://www.phototechmag.com/ but I do } not know if the review is available there. } } Geoff } } Gary Gaugler wrote: } } } } } } } ------------------------------------------------------------------------------ } } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } ------------------------------------------------------------------------------- } } } } } } I was a fan of Epson photo printers for quite } } some time. Notable was the 890 & 980. It is a } } small format printer compared to the 2200. } } I recently (a year or so ago) bought a Epson Stylus } } Photo 2000. It lasted about six months and } } then jammed constantly. In-warranty customer } } service and any idea of repair was on a wish list. } } It never happened. The printer was scrapped. } } } } The 2200 may have solved teething problems } } with large format printers. However, the 2000 } } was VERY slooooow using photo paper. When it } } worked, the results were stunning. Many times } } (too many) it would stop printing 1/4 or 1/2 } } way through the print and just die. The job } } hung (Win2K Pro) and had to be restarted with } } a new sheet of paper. } } } } The Epson and Canon small format printers seem } } to do a better, more reliable job. As a result } } of being burned by Epson, I now take print jobs } } to a local service bureau. they do a very nice } } job for not much cost. These are mostly for } } 24" x 48" glossy mounted prints. Small ones } } are done on my HP 4550 color laser printer. } } If the color gamut is matched well between } } the monitor and Photoshop, the HP does a nice job } } for reports. For transparencies (not much used } } any longer), the Kodak dye sub is excellent. } } } } Let us know what you find. There are a lot } } of options. Also, check out the Ethernet print } } servers that will connect a non-network printer } } to a LAN and allow all to use it. HP and others } } make these. they usually cost about $100 or so. } } } } gary g. } } } } } } At 11:08 AM 9/2/2004, you wrote: } } } } } Email: jtd1-at-psu.edu } } } Name: Tom Doman } } } } } } Organization: Penn State University } } } } } } Title-Subject: [Microscopy] [Filtered] MListserver: } } } } } } Question: Our lab is considering various photo printers (} $1K) for } } } production of electron micrographs. Currently the Epson 2200 is the } } } stromg favorite. Are there any recomendations for other printers } } } which we should consider? What are your reasons for the recomended } } } printer? } } } } } } Thanks in advance! } } } } } } Tom } } } } } } --------------------------------------------------------------------------- } } } } } } } } } } } }
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 11:05:30 2004
If a rotating platen is used with a hand grinder, the flat surface of the hand grinder is usually rounded off and can become facetted. However, you can use a polishing station, just don't turn it on; keep it stationary. I have used the South Bay Technology Lapping tray with all of the common hand grinders, South Bay's, Fischione's, and Gatan's. This is my preferred way of thinning TEM samples down to 100 um or less. I have also used the Gatan lapping tray. I found this one too small and I don't like using the PSA backing on the disks (besides the high costs of the disks). With the lapping tray, I use 8" SiC disks without the PSA backing and hold them down by hand. They don't last long enough to go through the hassle of pasting them down. In addition, the lapping tray works very well with the 3M diamond films and they can last a very long time depending on your material. For III-V compounds, I have used a single 30 um disk for months. These work well, because the water used for lubrication can be kept on the pad because they are hydrophobic. (Careful, not all diamond films are the same and other brands are not hydrophobic and have this property.) They stay put when water is used to hold them down. As in the Tripod(TM) polishing technique, you hold them down by putting a puddle of water on the plate and then carefully put the pad down moving it around so that no air bubbles get under the pad and then squeegee the water out. It will stay put like that for over 1/2 hr if you do not let any water get under it.
A word about the glass used in the lapping tray, since I work at the Glass Technology Center. Glass has two sides, the air side and the tin side. The tin side is the smoothest and is less sensitive to corrosion. You can tell the Sn side because it will fluoresce under UV. If you can determine it, this is the side that you should put your 3M papers on. You do not want water with any of the ground material to dry on the glass. If there are any silicates present, they will bond with the glass when it dries and you will not be able to remove it. I always use distilled water with the lapping trays, but that may not always be possible.
Another word of caution, don't drop your hand grinder on the glass plate! Not everyone can replace it as easily as I did.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472 Walck-at-PPG.com (412) 820-8651 (office) (412) 820-8515 (fax)
-----Original Message----- } From: Mary Mager [mailto:mager-at-interchange.ubc.ca] Sent: Wednesday, September 15, 2004 6:23 PM To: hkonishi-at-indiana.edu Cc: Microscopy
Dear Hiromi, Because the amount of material removed from 3.05 mm TEM discs with the Gatan Disc Grinder is very small, I use 600 grit paper stuck to a glass sheet and carefully hand grind the samples under running water, then finish off with 1200 grit the same way. It only takes a few strokes to remove 50 microns, then advance the dial and remove another 50 microns. We have spinning wheel grinders and automatic polishers, but I would not use them for this. Regards, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: {hkonishi-at-indiana.edu} To: {Microscopy-at-microscopy.com} Sent: Wednesday, September 15, 2004 12:23 PM
Members, In my opinion, the HP 7960 is a great photo printer. It has it all - dedicated gray level ink cartridge, tiny ink droplet size, low cost ( { $200 at the typical warehouse club stores) and has all of the various memory card slots on the front panel for your digital camera along with a color LCD display for previewing the images. Even on standard copy paper, the images really come out nice.
Disclaimer: I do not work for HP nor have any financial interest in said company. I do however, bundle this printer (at the end user's request) with the SEM Digital Image Capture Systems that I do sell.
Gary M. Easton Scanners Corporation Third Party SEM Service/Digital Imaging/EDS Sales 410-857-7633 x102
----- Original Message ----- } From: "Geoff McAuliffe" {mcauliff-at-umdnj.edu} To: "Gary Gaugler" {gary-at-gaugler.com} Cc: "by way of MicroscopyListserver" {jtd1-at-psu.edu} ; "MSA listserver" {Microscopy-at-MSA.Microscopy.Com} Sent: Thursday, September 16, 2004 1:33 PM
Recently I acquired an HP 6540 printer, not a photo printer as such (no built in card reader or LCD screen), but I have been extremely impressed with its quality and speed in printing my SEM images (formerly used an HP 970). It is very quick - prints two 4x5" photos on one sheet of HP Premium inkjet paper in 30 seconds (standard print quality setting). I compared the output quality with printing at the top quality setting and found little if any visible improvement for the doubling in output time.
It also takes gray or photo color cartridges in place of the black one. I have not been impressed with either of these, the colors seem weak and don't match the ones on my monitor, which are gray scale. So for all that I'm sticking with the standard black and color cartridges for the best results here.
I had been looking at the HP 7960, but purchasing wrinkles pointed me to the 6540 and I'm happy they did. Was a bit cheaper too ($150).
Richard Shalvoy Arch Chemicals, Inc. 350 Knotter Drive Cheshire, CT 06410 (203) 271-4394 rbshalvoy-at-archchemicals.com
-----Original Message----- } From: rschoon [mailto:rschoon-at-email.unc.edu] Sent: Thursday, September 16, 2004 12:32 PM To: Geoff McAuliffe Cc: Gary Gaugler; by way of MicroscopyListserver; MSA listserver
Hi Omayra, We have used Parducz since 1967 and it works wonders on bacteria, protozoa, and many other critters especially those with flagella and cilia. Two things that we have found important in its use are: 1. Make it FRESH just before use We keep a saturated solution of HgCl2 around in a brown 50 ml bottle which has been as old as one year. To make it we put a bit of HgCl2 into distilled water until we see that it is saturated i.e. with a ppt on the bottom. We also keep a bottle of 2% aq OsO4 in the refrigerator (usually double jarred to prevent vapors from escaping). JUST before use then we mix 6 ml of 2% OsO4 and 1 ml of the saturated HgCl2 (taken from the top of the bottle). We then fix whatever we are fixing for an hour or so. Obviously if more fix is needed, larger amounts can be made. 2. Wash WELL after the Parducz fixation. Otherwise you get starfish shaped crystals on the surface of your prep. They may look neat but obviously are not part of the specimen. Other than that, it is a great hardening fix for SEM.
Good Luck, Judy Murphy Stockton, CA
Omayra Velez wrote:
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From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 13:30:40 2004
Greetings Hiromi, A good solution would be a linear encoder for upright microscopes with a digital readout. We use a unit from Hiedenhain (www.heidenhain.com/product.html) on our stereology rig. There may be a way to set one up for inverted bases as well. Digital length gauge systems or linear encoders can measure z position with very high precision. You may wish to contact the folks at MicroBrightfield (www.microbrightfield.com) as they sell such solutions specifically for microscopes. Simply calibrating the markings on focus knobs to displacement of the stage in z may not be very precise in some instances. Older axiovert bases have a friction based focusing system which can slip depending on the speed or distance through which the knob is turned. Other bases may have similar issues with the focus mechanism. Calibration strategies in which the top and bottom of a z calibration standard are imaged to determine endpoints will have to take into account the uncertainty due to depth of focus of a particular objective lens as well. The linear encoder avoids this uncertaintly. Regards, Karl
hkonishi-at-indiana.edu wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-- Karl Garsha Light Microscopy Specialist Imaging Technology Group Beckman Institute for Advanced Science and Technology University of Illinois at Urbana-Champaign 405 North Mathews Avenue Urbana, IL 61801 Office: B650J Phone: 217.244.6292 Fax: 217.244.6219 Mobile: 217.390.1874 www.itg.uiuc.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 14:08:21 2004
We just purchased a Canon i9900. The initial test prints are beautiful. It uses eight cartridges and when using Canon's Pro Paper, you would not know that they are computer prints rather than high quality true photographs. We elected to have a larger format capacity (13" x 19") as we like to decorate our lab and office walls with our favorites.
Al Stone ASTON Metallurgical Services
At 01:02 PM 9/16/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 16:20:53 2004
I really enjoy being on this forum and am educated a lot. Now I have a question about how often we need to flash our field emission SEM's tip.
We have a brand new Hitachi 4700 which has a cold field emission tip. When it was first installed, the service engineer told us to record the initial extraction voltage each time right after flashing, during the use if the increase of extraction voltage is more than 1.4 KV, we need to do a flash. We follow this rule strictly. Several weeks ago Hitachi's application engineer visited us and said by doing that we flashed too much and would short our tip's lifetime. We do not need to flash until we see the tip noise. I never see any tip noise in my image by following his suggestion so I am forced to flash by the instrument setting (every 48 hours).
Could you please kindly share your experience with me?
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I have a BioScan Ted Pella microwave with a cold-spot and vacuum chamber that I would like to use for fixation and embedding insect eggs and larvae for TEM.
The project includes mosquito larvae, pupae and fly eggs. I worked on this same project many years ago using traditional benchtop methods. The eggs and larvae were difficult, if not impossible, to infiltrate. Now that I have a MW with vacuum I am trying this project again.
I would greatly appreciate any suggestions, advice or direct me to references?
My first attempt at MW didn't go so well. I MW fixed the larvae with 2% glut in cacodylate in 1.5ml centrifuge tubes at 20mmHg vacuum -at- 250W 1' on, 1' off. Repeated several times removing the vacuum to observe the larvae. At first when vacuum was applied the larvae floated to the top and squirmed around a bit, then after several attempts at that they finally all stopped moving and didn't float. Buffer washed on benchtop then used the vacuum again at the same settings for the buffered osmium tetroxide. Just the ends of the larvae turned black. I tried it again without any change. I finally got tired of messing with them and placed them into a KFeCN + OsO4 overnight at 4C and have had them stored in buffer. ( I have used the overnight KFeCN + OsO4 with success for hard to infiltrate nematodes). Dehydrated with MW 40s -at- 250W no vacuum 40 s on, 1' off, infiltrated 50% acetone/Embed no vac at 250W 3', 100% Embed -at- 450W with vac (20mm Hg). The larvae had areas of collapse after the 100% with vacuum so I'm concerned that they are not infiltrating as I had hoped.
Karen L. Kelley ICBR Electron Microscopy Manager University of Florida ICBR Electron Microscopy Core Lab Bartram Hall Room 214 Box 118525 Gainesville Florida Lab: 352-392-1184 fax: 352-846-0251 Email: klk-at-biotech.ufl.edu Southeastern Microscopy Society Treasurer http://www.biotech.ufl.edu/EM/
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 17:18:35 2004
Dear colleague, I remember some discussion about CD-R storage. Some CD are not good at all and in many cases some CD become unreadable after 3 years. I would like to know what is the best brand or type of CD for storing images
Keep care and be of good cheer
Regards
(name) Vratislav Richard Eugene Maria John Baptist (surname) of Bejsak (Bayshark)-Colloredo-Mansfeld
Tenebrionidae of the World, incl. Alleculinae and Lagriinae, higher taxonomy, Australian beetles. websites: http://www.coleoptera.org. and http://www.egroups.com/group/coleoptera
University of Sydney The Wentworth Bldg., B 62 NSW 2006 AUSTRALIA phone : +61 414 540 465 email: ricardo-at-ans.com.au vratislav-at-bigfoot.com ICQ: 13610107
Only after the last tree has been cut down, only after the last river has been poisoned, only after the last fish has been caught, only then will you find that money can not be eaten.' CREE INDIAN PROPHECY.
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From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 17:23:42 2004
We have been using a digital length gauge from Heidenhain. the readout unit has an RS-232 port so that we can record depth with a computer. It uses an optical encoder that has been clamped in a holder so that it can be placed below the stage.
Our cost was about $1500, but that was about 1993.
Regards, Glen On Sep 16, 2004, at 9:11 AM, hkonishi-at-indiana.edu wrote:
} } } ----------------------------------------------------------------------- } ------- } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ----------------------------------------------------------------------- } -------- } } I am looking for a "micrometer" that can work with optical microscope. } I think } that it read the difference of dial for focusing and convert to actual } distance (um). I want to use it for determining the thickness of disc } during } disc grinding if it is not so expensive. } Please advise on website or product information. } Thank you, } } Hiromi Konishi } Indiana University } } } Glen MacDonald Core for Communication Research Virginia Merrill Bloedel Hearing Research Center Box 357923 University of Washington Seattle, WA 98195-7923 USA (206) 616-4156 glenmac-at-u.washington.edu
************************************************************************ ****** The box said "Requires Windows 95 or better", so I bought a Macintosh. ************************************************************************ ******
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 17:32:22 2004
Can you make an estimate how much a high quality print, letter size in full color and b/w costs (probably just ink and paper)?
I'd be interested also for the HP that was mentioned in the original post.
mike
Michael Bode, Ph.D. Soft Imaging System Corp. 12596 West Bayaud Avenue Suite 300 Lakewood, CO 80228 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: Alan Stone [mailto:as-at-astonmet.com] Sent: Thursday, September 16, 2004 13:36 To: microscopy-at-microscopy.com
We just purchased a Canon i9900. The initial test prints are beautiful. It uses eight cartridges and when using Canon's Pro Paper, you would not know that they are computer prints rather than high quality true photographs. We elected to have a larger format capacity (13" x 19") as we like to decorate our lab and office walls with our favorites.
Al Stone ASTON Metallurgical Services
At 01:02 PM 9/16/2004, you wrote:
} --------------------------------------------------------------------------- --- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 18:53:41 2004
IMO, Mitsui is the absolute best for CD-R or DVD-R:
http://www.mitsuicdr.com/
gary g.
At 03:46 PM 9/16/2004, you wrote:
} Dear colleague, } I remember some discussion about CD-R storage. Some CD are not good at all } and in many cases some CD become unreadable after 3 years. } I would like to know what is the best brand or type of CD for storing } images } } Keep care and be of good cheer } } Regards } } (name) Vratislav Richard Eugene Maria John Baptist } (surname) of Bejsak (Bayshark)-Colloredo-Mansfeld
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 20:26:25 2004
} Dear colleague, } I remember some discussion about CD-R storage. Some CD are not good at all } and in many cases some CD become unreadable after 3 years. } I would like to know what is the best brand or type of CD for storing } images } } Keep care and be of good cheer } } Regards } } (name) Vratislav Richard Eugene Maria John Baptist } (surname) of Bejsak (Bayshark)-Colloredo-Mansfeld
I would recommend Mitsui. Remember, the ideal conditions for storage of CDs is very similar to negatives: cool, dark, and dry.
It is essential to remember that a CDR is -absolutely not- an archival copy of your data.
Brent
-- Brent Neal, Ph.D. Reindeer Graphics, Inc. brent-at-reindeergraphics.com
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 16 22:43:50 2004
DO NOT use sharpie markers or any other acid base pen to label cd's or dvd's. The acid in these pens etches through the top of the disk destroying your data.
Dave Crone B.E. (Mechanical) Engineer-in-Training Department Assistant Metallurgical Lab Mechanical Engineering College of Engineering University of Saskatchewan 57 Campus Drive Saskatoon, SK S7N 5A9 Phone: (306) 966-5461 Fax: (306) 966-5427 E-mail: dgc132-at-mail.usask.ca -----Original Message----- } From: Brent Neal [mailto:brent-at-reindeergraphics.com] Sent: Thursday, September 16, 2004 7:54 PM To: Vr.R.E.M.J..-B. BEJSAK-COLLOREDO-MANSFELD Cc: MICROSCOPY
} Dear colleague, } I remember some discussion about CD-R storage. Some CD are not good at all } and in many cases some CD become unreadable after 3 years. } I would like to know what is the best brand or type of CD for storing } images } } Keep care and be of good cheer } } Regards } } (name) Vratislav Richard Eugene Maria John Baptist } (surname) of Bejsak (Bayshark)-Colloredo-Mansfeld
I would recommend Mitsui. Remember, the ideal conditions for storage of CDs is very similar to negatives: cool, dark, and dry.
It is essential to remember that a CDR is -absolutely not- an archival copy of your data.
Brent
-- Brent Neal, Ph.D. Reindeer Graphics, Inc. brent-at-reindeergraphics.com
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 01:24:07 2004
We use on an old OPL inverted microscope (~1950) a Mitutoyo digital workshop micrometer (Digimat IDF 543 serie, ref 543-511) witch gives 1 micron as last digit. It's easy to put in on inverted microscope, a bit more complicate for a upright one. Very useful and reliable enough for all samples thinning job for TEM (before tripod polishing or dimpler thinning). I don't remember the price, we bought it 6-7 years ago, but it's à workshop stuf, not a laboratory one. So it should not be much expensive. Some models can be interface to record measurements.
No interest in that manufacturer, just satisfied from that product.
J. Faerber IPCMS-GSI (Institut de Physique et Chimie des Matériaux de Strasbourg Groupe Surface et Interfaces) 23, rue de Loess ; BP43 67034 Strasbourg CEDEX 2 France
On Thu, 16 Sep 2004 hkonishi-at-indiana.edu wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } I am looking for a "micrometer" that can work with optical microscope. I think } that it read the difference of dial for focusing and convert to actual } distance (um). I want to use it for determining the thickness of disc during } disc grinding if it is not so expensive. } Please advise on website or product information. } Thank you, } } Hiromi Konishi } Indiana University } }
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 06:30:11 2004
For archival quality: you should always finalise the disk and don't bother with multisession, consider copying at a slower than maximum speed for the drive and disk. Run the disk check software and maybe check some images on the CD (or just the disk directory) because problems may happen at the copying stage. I have used those special CD markers (I know TDK make some) for about two to three years without problem.
Make 2 (or 3) copies instead of one and store separately and check one periodically - at least if one fails you have a reasonable chance that the other(s) may survive.
Make sure that you will be able to read them in the next few years - with the advance of technologies such as 'Blu-ray' DVD writers how long will CD compatibility be maintained? You only have to look at 8, 5 1/4, & 3 inch floppies and some of the older magneto optical formats to realize that the format may have a shorter useful lifetime than its archival quality.
Malcolm
PS I don't always practice what I preach.
Malcolm Haswell e.m. unit School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear SR1 3SD UK e-mail: malcolm.haswell-at-sunderland.ac.uk
----- Original Message ----- } From: Dave Crone {dgc132-at-mail.usask.ca}
So what should be used to archive electronic data?
Richard Doelle Dofasco Inc. richard_doelle-at-dofasco.ca
-----Original Message----- } From: Brent Neal [mailto:brent-at-reindeergraphics.com] Sent: Thursday, September 16, 2004 9:54 PM To: Vr.R.E.M.J..-B. BEJSAK-COLLOREDO-MANSFELD Cc: MICROSCOPY
HI Garry, I too, mix by weight but I have a friend who uses disposable syringes to measure by volume. No cleaning, since she uses each syringe once and discards it (adding to the trash load...we just can't win). She sets up her resin component bottles with stoppers and tubing that she can seal off when not measuring and which she can just hook up to the syringes and then draw out the volume she needs. I know that she got this method from her first employer and that between them the method has worked beautifully for 30 years or more. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 08:02:17 2004
At 4:47 PM -0500 9/16/04, Ying.Shi wrote: } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Ying Shi,
We have had a Hitachi S4700 since 10/1996. We typically only flash the tip only once in the morning each day. If I remember correctly, the instrument was originally programmed to make you flash about every eight hours. We had the service engineer change the required flash time to 16 hours. This allows us to have a stable beam current for XEDS line scans and maps in the afternoon. We record the current at each flash, when this gets above a certain level it indicates to the service engineer that the column needs to be baked, typically twice a year. We are still using our original tip!! Good luck with your S4700.
-- David R. Hull NASA Glenn Research Center at Lewis Field Advanced Metallics Branch Mail Stop 49-1 21000 Brookpark Road Cleveland, OH 44135
I think you would be safe in doing as the applications engineer tells you, i.e. every 48 hrs. Of course, if you have samples that outgas, this may need to be more frequent. If the emission current is stable, and the extraction voltage is stable, just let it run.
We have 2 4700's and a 4500 here at Agere, Allentown, PA.
Hope that is of some help.
Peter Tomic Agere System
-----Original Message----- } From: Ying.Shi [mailto:Ying.Shi-at-delphi.com] Sent: Thursday, September 16, 2004 5:48 PM To: Microscopy-at-MSA.Microscopy.com
Dear All EM Specialists:
I really enjoy being on this forum and am educated a lot. Now I have a question about how often we need to flash our field emission SEM's tip.
We have a brand new Hitachi 4700 which has a cold field emission tip. When it was first installed, the service engineer told us to record the initial extraction voltage each time right after flashing, during the use if the increase of extraction voltage is more than 1.4 KV, we need to do a flash. We follow this rule strictly. Several weeks ago Hitachi's application engineer visited us and said by doing that we flashed too much and would short our tip's lifetime. We do not need to flash until we see the tip noise. I never see any tip noise in my image by following his suggestion so I am forced to flash by the instrument setting (every 48 hours).
Could you please kindly share your experience with me?
Note: The information contained in this message may be privileged and confidential and thus protected from disclosure. If the reader of this message is not the intended recipient, or an employee or agent responsible for delivering this message to the intended recipient, you are hereby notified that any dissemination, distribution or copying of this communication is strictly prohibited. If you have received this communication in error, please notify us immediately by replying to the message and deleting it from your computer. Thank you.
We have a Hitachi S4700, too, and were told by our service engineer to flash about 1.3 KV above the starting extraction voltage. We have had the same tip since installation in 1999 and since then the starting extraction voltage (i.e., right after flashing) has increased from 4.1 to 4.3 KV at 5.0 KV accelerating voltage at 10ua beam current.
My impression from this is that the tip is relatively robust and long-lasting, even under frequent flashing. This included one period of VERY frequent flashing due to a bad connection to the gun which prevented flashing from occurring at the proper strength. When this was discovered and repaired, the first flash was so powerful it gave us a reading of 72, rather the recommended 20, before we dialed down the flash intensity! We were afraid that we'd taken the end right off the tip with this one, but there was no apparent harm and resolution has remained fine since then.
Whether or not this is better than flashing only when the software tells you to is unclear to me, but this is how we were told to do it.
Hope this helps.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
-----Original Message----- } From: Ying.Shi [mailto:Ying.Shi-at-delphi.com] Sent: Thursday, September 16, 2004 4:48 PM To: Microscopy-at-MSA.Microscopy.com
Dear All EM Specialists:
I really enjoy being on this forum and am educated a lot. Now I have a question about how often we need to flash our field emission SEM's tip.
We have a brand new Hitachi 4700 which has a cold field emission tip. When it was first installed, the service engineer told us to record the initial extraction voltage each time right after flashing, during the use if the increase of extraction voltage is more than 1.4 KV, we need to do a flash. We follow this rule strictly. Several weeks ago Hitachi's application engineer visited us and said by doing that we flashed too much and would short our tip's lifetime. We do not need to flash until we see the tip noise. I never see any tip noise in my image by following his suggestion so I am forced to flash by the instrument setting (every 48 hours).
Could you please kindly share your experience with me?
Note: The information contained in this message may be privileged and confidential and thus protected from disclosure. If the reader of this message is not the intended recipient, or an employee or agent responsible for delivering this message to the intended recipient, you are hereby notified that any dissemination, distribution or copying of this communication is strictly prohibited. If you have received this communication in error, please notify us immediately by replying to the message and deleting it from your computer. Thank you.
We have a Hitachi S-900 field-emission SEM, and I agree with the 2nd engineer's comment. Flash when you see noise, not by hours or extraction voltage. If the extraction voltage gets excessive, then I'd consider flashing, but "excessive" depends on your instrument and operating variables. 1.4kV may not be "excessive". For us it would be -- but we rarely seen that much increase in extraction V.
Phil
} Dear All EM Specialists: } } I really enjoy being on this forum and am educated a lot. Now I have } a question about how often we need to flash our field emission SEM's } tip. } } We have a brand new Hitachi 4700 which has a cold field emission } tip. When it was first installed, the service engineer told us to } record the initial extraction voltage each time right after } flashing, during the use if the increase of extraction voltage is } more than 1.4 KV, we need to do a flash. We follow this rule } strictly. Several weeks ago Hitachi's application engineer visited } us and said by doing that we flashed too much and would short our } tip's lifetime. We do not need to flash until we see the tip noise. } I never see any tip noise in my image by following his suggestion so } I am forced to flash by the instrument setting (every 48 hours). } } Could you please kindly share your experience with me? } } Thanks } } Ying Shi } } Analytic Scientist } Delphi Catalyst } ying.shi-at-delphi.com } } } **************************************************************************************** } } Note: The information contained in this message may be privileged } and confidential and thus protected from disclosure. If the reader } of this message is not the intended recipient, or an employee or } agent responsible for delivering this message to the intended } recipient, you are hereby notified that any dissemination, } distribution or copying of this communication is strictly } prohibited. If you have received this communication in error, please } notify us immediately by replying to the message and deleting it } from your computer. Thank you. } } ****************************************************************************************
-- Philip Oshel Supervisor, BBPIC microscopy facility Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 08:33:55 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (bgorman-at-unt.edu) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, September 16, 2004 at 09:59:31 ---------------------------------------------------------------------------
We are dismantling our H-9000 TEM, and have a brand new (never installed) gun housing with retro fit kit available to anyone interested. Please contact me for more details.
If anyone wants other parts of this microsope, please contact me as well.
Regards, Brian
Brian P. Gorman, PhD Assistant Professor Dept. of Materials Science and Engineering University of North Texas Denton, TX 76203 bgorman-at-unt.edu ph: 940-891-6778 fax: 940-565-4824
Depending on your definition of archival, Mitsui DVD-R media ought to work for you. Their media is rated at 75 year retention. Their Gold Archival CD-R is rated at 100 years. Obviously, don't expect these results if you leave them on the rear deck of your car for twenty years.
I believe that this topic has been discussed before and archived. If not, it will get archived now.
My belief and strategy is to have four, redundant backups. The first backup is either dual CD-R or dual DVD-R. Second backup is dual hard drives. These are typically 120GB IDE/ATAPI. Third backups are two SNAP/NAS servers, 340GB-1TB. Fourth backup are two identical Ultrium 1 or Ultrium 2 LTO tapes.
The hard drives are in removeable bays and are real time storage and retrieval media. The SNAP/NAS servers are about the same but are slower due to LAN speed. Once a directory of data on a drive will fill a CD or DVD, it is written to the optical media. At this point, the last backup is to the optical media. The first backup is to hard drive, then to NAS, then to LTO and finally, optical.
The LTO 1 media will hold 100GB of un-compressed data. The LTO 2 holds 200GB. Since the image data is either TIFF or JPEG, it does not really compress much more at all. LTO is backward compatable to LTO 1. The drives are wide SCSI and work with most all operating systems. NovaStore and Veritas make backup software for these drives.
A most important aspect of backing up is to actually do it.
gary g.
At 06:05 AM 9/17/2004, you wrote:
} So what should be used to archive electronic data? } } Richard Doelle } Dofasco Inc. } richard_doelle-at-dofasco.ca } } } -----Original Message----- } } From: Brent Neal [mailto:brent-at-reindeergraphics.com] } Sent: Thursday, September 16, 2004 9:54 PM } To: Vr.R.E.M.J..-B. BEJSAK-COLLOREDO-MANSFELD } Cc: MICROSCOPY } Subject: [Microscopy] Re: Brand / type of CD-R disk for sImages } } } } } I would recommend Mitsui. Remember, the ideal conditions for storage of } CDs is very similar to negatives: cool, dark, and dry. } } It is essential to remember that a CDR is -absolutely not- an archival } copy of your data. } } Brent } } -- } Brent Neal, Ph.D. } Reindeer Graphics, Inc. } brent-at-reindeergraphics.com
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 10:42:03 2004
For all you aficionados of classic, operating ultramicrotomes---this is your chance! We are disposing of four LKB Ultratome III's (control and main units), which, when last assembled, comprised three operating units plus one for spare parts. Included are numerous separate spare parts, including chucks, belts, fuses, etc., etc., etc.
We also have a operating Pyramitome 11800 and an LKB Historange 2218 Microtome.
The catch: whoever gets these units is responsible for supplying packing materials and shipping costs. The machines themselves are free.
We had been using this equipment for all of our ultramicrotomy until the last couple years and had planned to keep using them as teaching units, but we are desperately short on space. We hope we can find a good home for them out there in EM land, because our alternative is surplusing them with our beat up filing cabinets and broken computer monitors.
Any takers?
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 12:30:56 2004
On Sep 16, 2004, at 3:46 PM, Vr.R.E.M.J..-B. BEJSAK-COLLOREDO-MANSFELD wrote:
} I remember some discussion about CD-R storage. Some CD are not good at } all } and in many cases some CD become unreadable after 3 years. } I would like to know what is the best brand or type of CD for storing } images } } Keep care and be of good cheer } Dear Vratislav, We are using Imation disks. Since the cost is small, we get the best combination of brand name and price. If someone like Computer Shopper has done a test for the reliability of various brands of CD-R or DVD-R disks, I'd be interested in the results. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 14:25:05 2004
Agree on this interest point. I've bought brand-name CD-R that show a rating of "48X Multispeed" and found that my recorder (rated up to 24X) can only record at 4X on these CDs. Curiously, other CD-R from the same manufacturer with nominally the same rating and price can go at the full 24X of my recorder. I asked the fellow at our local computer shop and he gave me the blank stare of ignorance followed by a shoulder shrug.
If anyone can explain how I can determine which CD-R will record "fast" (and as Vratislav suggests, will also have a long life) I'm all ears. I suppose speed ratings on CDs are like mileage ratings on cars - your results may vary from the sticker...
Bill William A. Heeschen, Ph.D. Microscopy, Digital Imaging The Dow Chemical Company Midland, MI 48674 waheeschen-at-dow.com
-----Original Message----- } From: Bill Tivol [mailto:tivol-at-caltech.edu] Sent: Friday, September 17, 2004 2:07 PM To: microscopy-at-msa.microscopy.com
Wow, there's a big demand for Ultratome III's out there. I think I have enough takers for now, so early next week I'll sort through the replies and see what I can do, starting with the first ones.
Correction: The Historange microtome is not included, after all. I almost gave away my boss's machine by mistake....
Thanks for responding.
Randy
-----Original Message----- } From: Tindall, Randy D. [mailto:TindallR-at-missouri.edu] Sent: Friday, September 17, 2004 1:31 PM To: microscopy-at-microscopy.com Cc: Katz, Martin
For all you aficionados of classic, operating ultramicrotomes---this is your chance! We are disposing of four LKB Ultratome III's (control and main units), which, when last assembled, comprised three operating units plus one for spare parts. Included are numerous separate spare parts, including chucks, belts, fuses, etc., etc., etc.
We also have a operating Pyramitome 11800 and an LKB Historange 2218 Microtome.
The catch: whoever gets these units is responsible for supplying packing materials and shipping costs. The machines themselves are free.
We had been using this equipment for all of our ultramicrotomy until the last couple years and had planned to keep using them as teaching units, but we are desperately short on space. We hope we can find a good home for them out there in EM land, because our alternative is surplusing them with our beat up filing cabinets and broken computer monitors.
Any takers?
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.emc.missouri.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 17:39:54 2004
I don't think the terms "Archival" and CD-R/DVD-R technologies are compatible and should not be used in the same sentence unless associated with the phrase "do not count on".
Yes, manufacturers will offer ratings of 75 or 100 years.
If the media is not exposed to _ANY_ variations in temperature, humidity, etc. and never handled, I expect you could see that life. I know of very few places on earth that meet those requirements.
There are a number of potential things that will render the media unusable. This includes not only the recordable, but also the "pre-recorded" or "permanent" versions.
The permanent versions especially are susceptible to the aluminum layer (the LABEL side) oxidizing, and then falling apart.
As the disks go through thermal cycling, there can also be problems with de-lamination.
Scratches on the "bottom" side of the media are relatively harmless. Some media is VERY susceptible to scratching and damage on the top or label side. In addition to avoiding the acidic "Sharpies", avoid pencils, ball point pens, etc. I would also caution against trying to "undo" any pressure sensitive label.
DVD-R may prove to be more stable then their CD-R counterparts. I would still be cautious. Many of the vulnerabilities of CDs (permanent and recordable) were not realized until recently. There may be unknown vulnerabilities to DVD-Rs that we don't know about yet either.
Probably the most reliable "archive" media is still magnetic tape. I have know people that have taken a tape from a "fireproof" safe, where the tape cartridge was melted beyond use (due to longer than rated exposure), and even appeared "melted into" the tape. The tape was sent out to a recovery specialist and 90% plus of the data was recovered. The stuff takes a beating and keeps on winding.
John W. Raffensperger, Jr. IT Manager Apache Stainless
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 17:55:04 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (bwadgaobkar-at-mail.mcg.edu) from http://www.msa.microscopy.org/Ask-A-Microscopist/Ask-A-Microscopist.html on Friday, September 17, 2004 at 13:15:12 ---------------------------------------------------------------------------
Question: We are studying a polymer (poly e-caprolactone)thats semicrystelline. We were wondering how to stain just the crystelline lamella to eatimate the crystillinity and study the morphology.
I concur with what all that was said. But still cd's are cheap, 25 cents a piece here, and at 40x or faster you are looking at 5 to 6 min burn. A non-acidic marker is $2, so backup your data and if you are unsure back it up again in 6 months. If it is dvd-r they are $0.46 and you can fit 4.2 GB on them, so back up often! If you have problems with data backup feel free to contact me I will help you out as much as I can.
Dave Crone B.E. (Mechanical) Engineer-in-Training Department Assistant Metallurgical Lab Mechanical Engineering College of Engineering University of Saskatchewan 57 Campus Drive Saskatoon, SK S7N 5A9 Phone: (306) 966-5461 Fax: (306) 966-5427 E-mail: dgc132-at-mail.usask.ca -----Original Message----- } From: Chiphead [mailto:chiphead-at-sbcglobal.net] Sent: Friday, September 17, 2004 5:07 PM To: 'Microscopy MSA'
I don't think the terms "Archival" and CD-R/DVD-R technologies are compatible and should not be used in the same sentence unless associated with the phrase "do not count on".
Yes, manufacturers will offer ratings of 75 or 100 years.
If the media is not exposed to _ANY_ variations in temperature, humidity, etc. and never handled, I expect you could see that life. I know of very few places on earth that meet those requirements.
There are a number of potential things that will render the media unusable. This includes not only the recordable, but also the "pre-recorded" or "permanent" versions.
The permanent versions especially are susceptible to the aluminum layer (the LABEL side) oxidizing, and then falling apart.
As the disks go through thermal cycling, there can also be problems with de-lamination.
Scratches on the "bottom" side of the media are relatively harmless. Some media is VERY susceptible to scratching and damage on the top or label side. In addition to avoiding the acidic "Sharpies", avoid pencils, ball point pens, etc. I would also caution against trying to "undo" any pressure sensitive label.
DVD-R may prove to be more stable then their CD-R counterparts. I would still be cautious. Many of the vulnerabilities of CDs (permanent and recordable) were not realized until recently. There may be unknown vulnerabilities to DVD-Rs that we don't know about yet either.
Probably the most reliable "archive" media is still magnetic tape. I have know people that have taken a tape from a "fireproof" safe, where the tape cartridge was melted beyond use (due to longer than rated exposure), and even appeared "melted into" the tape. The tape was sent out to a recovery specialist and 90% plus of the data was recovered. The stuff takes a beating and keeps on winding.
John W. Raffensperger, Jr. IT Manager Apache Stainless
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 20:18:09 2004
(9/17/04 9:01) Gary Gaugler {gary-at-gaugler.com} wrote:
} Depending on your definition of archival, Mitsui DVD-R } media ought to work for you. Their media is rated at } 75 year retention. Their Gold Archival CD-R is rated } at 100 years. Obviously, don't expect these results } if you leave them on the rear deck of your car for } twenty years. } } I believe that this topic has been discussed before } and archived. If not, it will get archived now. } } My belief and strategy is to have four, redundant backups. } The first backup is either dual CD-R or dual DVD-R.
} } The hard drives are in removeable bays and are real time } storage and retrieval media. The SNAP/NAS servers are \ } } The LTO 1 media will hold 100GB of un-compressed data.
} } A most important aspect of backing up is to actually do it. } } gary g.
Of course, in 20 years, no drives will exists that will read any of the above media, so it really doesn't matter how long the supposed "archival" DVDs last, now does it. How many DECtape readers have -you- seen recently?
Brent
-- Brent Neal, Ph.D. Reindeer Graphics, Inc. brent-at-reindeergraphics.com
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 17 23:55:33 2004
} From: "Brent Neal" {brent-at-reindeergraphics.com} : : (9/17/04 9:01) Gary Gaugler {gary-at-gaugler.com} wrote: : : } Depending on your definition of archival, Mitsui DVD-R : } media ought to work for you. Their media is rated at : } 75 year retention. Their Gold Archival CD-R is rated : } at 100 years. Obviously, don't expect these results : } if you leave them on the rear deck of your car for : } twenty years. : } : } I believe that this topic has been discussed before : } and archived. If not, it will get archived now. : } : } My belief and strategy is to have four, redundant backups. : } The first backup is either dual CD-R or dual DVD-R. : : } : } The hard drives are in removeable bays and are real time : } storage and retrieval media. The SNAP/NAS servers are : \ : } : } The LTO 1 media will hold 100GB of un-compressed data. : : } : } A most important aspect of backing up is to actually do it. : } : } gary g. : : : : Of course, in 20 years, no drives will exists that will read any of the above media, so it really doesn't matter how long the supposed "archival" DVDs last, now does it. How many DECtape readers have -you- seen recently? : If you want to preserve your work, data, photos, notes and such forever there is no computer media up to the task. Only a system that relies on copies of the data sorted in geographically diverse location that are recopied to new media and updated media as it becomes available. This quickly becomes a very large difficult problem much like putting one grain of wheat on the first square of a chess board, 2 on the second, 4 on the third, ..... and you exceed the world capacity to grow wheat before the chess board is full. It is more practical with computer data but not a lot and any of us the are truly confident of the integrity our back ups are few and far between.
I still contend that black and white silver halide film is the least expensive longest lived option available. We can all do it and afford it and the technology is well tested an proven. Building a device to put digital information on 35 mm film would be trivial and we all have the equipment for our images. Adding a copy stand covers notes and printed matter. With today's technology that we can use that is proven to have a 150 year life stored in a shoe box. Modern film substrates probably fixed and toned stored in a controlled atmosphere should last a very long time indeed.
Digital images, data and writing are sure a lot more convenient but they are sure not as reliable or as long lived as conventional photography.
Gordon Gordon Couger gcc-at-couger.com
I collect links on information related to light microscopes. http://www.couger.com/microscope/links/gclinks.html Please forward any links or information you think might be useful to others. Microscope Manual at www.science-info.org
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 18 07:40:53 2004
} : Of course, in 20 years, no drives will exists that will read any } of the above media, so it really doesn't matter how long the } supposed "archival" DVDs last, now does it. How many DECtape readers } have -you- seen recently?
I don't know that I'd agree. Today's optical media and drives appear to offer a means for total backward compatibility ... e.g., DVD drive which can read CDs. This is of course up to the manufacturers sticking to it and supporting its application toward archiving.
That said ... I believe this thread should be as much about the dependability of the software. It isn't so much about the media ... e.g., if an archive was written today, what software would be able to restore it in 20 years? What can I use with confidence? ... which will write archives identical to the original directory structure, without splitting files, and verify the archive integrity??? Will the software maintain any database characteristics of the original collection of files?? What software would support and manage redundant archives across multiple devices (as Gary Gaugler suggested as required)??
cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com (in progress)
From MicroscopyL-request-at-ns.microscopy.com Sat Sep 18 08:29:08 2004
I have a DEC TK-70, circa 1990. I don't have any VAX systems any more so the drive is not really being used. That said, it is a DLT-II tape unit. This format is still offered, and media is still available. It is not a huge repository of data. So from a hardware standpoint, this is a viable option. But the point of selecting a long lived platform is valid.
The longer the media type and drives stay around, it is all that much longer before the data needs to be migrated to newer methods. If one picks reliable and solid backup software from either NovaStore or Veritas, they very much seem to be fully backwards compatible regardless of the OS that was used to host the app. Furthermore, these apps will backup to drives, tapes, CDs and DVDs. So, it gets back to the hardware media set. Once written, the app's format does not change. The new app will have new features but it allows reading older media that used earlier versions of the app.
The most long lasting standard as I recall is SCSI. If the drive works, it can still be read today. SCSI has evolved to SCSI-II and more exotic types like LVD (low voltage differential). Even Adaptec LVD adapters will work with single ended narrow SCSI devices. They have made dual channel adapters so each side can work with either type of media.
SCSI is fast but it is costly. Hence, this is why I prefer the IDE/ATAPI drives at about 1/3 the cost of a similar SCSI drive. MFM was around at first but gave way to IDE, and IDE is still here. What is happening now is that the parallel IDE/ATAPI is evolving to serial ATA (SATA). They are not plug compatible. Even if mother boards drop parallel IDE, there are SATA-to-IDE adapters. In fact, current versions of SATA drives include a SATA-to-IDE adapter since true SATA drives are just now coming to market.
My selection of backup was carefully made to support archiving tens of thousands of digital images. Many of these are 30MB-250MB TIFF each. As was pointed out, technology moves on. Consequently, a one time archive is not going to last indefinitely into the future. But it potentially could if properly designed. I'm not taking that chance.
How many specimens do you all prepare and archive? What about this facet of preservation? If the captured data was lost, having the specimen would allow re-doing the imaging. A pain for sure but not fatal. Properly prepared specimens stored in a vacuum seem to hold up for a long, long time with no discernable effects. So, archive the specimens (this can get overwhelming for sure) and archive the image results.
BTW, I use a fine point Sharpie to write on CDs and DVDs. I never have a problem. The big felt markers are a no-no.
gary g.
At 06:08 AM 9/18/2004, you wrote:
} Gordon Couger writes ... } } } : Of course, in 20 years, no drives will exists that will read any } } of the above media, so it really doesn't matter how long the } } supposed "archival" DVDs last, now does it. How many DECtape readers } } have -you- seen recently? } } I don't know that I'd agree. Today's optical media and drives appear to } offer a means for total backward compatibility ... e.g., DVD drive which can } read CDs. This is of course up to the manufacturers sticking to it and } supporting its application toward archiving. } } That said ... I believe this thread should be as much about the } dependability of the software. It isn't so much about the media ... e.g., } if an archive was written today, what software would be able to restore it } in 20 years? What can I use with confidence? ... which will write archives } identical to the original directory structure, without splitting files, and } verify the archive integrity??? Will the software maintain any database } characteristics of the original collection of files?? What software would } support and manage redundant archives across multiple devices (as Gary } Gaugler suggested as required)?? } } cheerios ... shAf :o)
From MicroscopyL-request-at-ns.microscopy.com Sun Sep 19 12:03:15 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kssim-at-mmu.edu.my) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, September 19, 2004 at 11:37:16 ---------------------------------------------------------------------------
Question: Does anyone know tell me whether there is stigmatism problem or auto focusing problem at high resolution on optical misroscope? How to solve this problem?? Let me know ...
Autofocusing on a digital microscope works fine, even at high magnification (63x, 1.4 NA) in both brightfield as well as fluorecence microscopy, as was published in:
Geusebroek J.M., Cornelissen F., Smeulders A. W. M., and Geerts H., Robust autofocusing in microscopy. Cytometry, 36(1), pp. 1-9, (2000).
Regardless of which algortihm you use, it is important to sample your Z-stack at the appropriate interval (see Geusebroek et al.).
Regards,
Peter
---------------------------------------------- Peter Van Osta
Director Imaging MAIA SCIENTIFIC (formerly Union Biometrica NV) Cipalstraat 3 B-2440 Geel, Belgium Tel.: +32 (0)14 570 620 Mobile: +32 (0)497 228 725 Fax.: +32 (0)14 570 621 Email: pvosta-at-maia-scientific.com Website: www.maia-scientific.com A Harvard Bioscience Company ----------------------------------------------
by way of MicroscopyListserver wrote: } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kssim-at-mmu.edu.my) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, September 19, 2004 at 11:37:16 } --------------------------------------------------------------------------- } } Email: kssim-at-mmu.edu.my } Name: kssim } } Organization: mmu } } Title-Subject: [Microscopy] [Filtered] MListserver: } } Question: Does anyone know tell me whether there is stigmatism problem or auto focusing problem at high resolution on optical misroscope? How to solve this problem?? Let me know ... } } Many thanks } Ks }
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 20 08:06:05 2004
Peter Van Osta provided a reference to a very helpful article. Sadly, there was a typo in the reference. The correct citation is
Geusebroek J.M., Cornelissen F., Smeulders A. W. M., and Geerts H., Robust autofocusing in microscopy. Cytometry, 39(1), pp. 1-9, (2000).
The article is worth the look...
I also found the following article helpful: A. Santos et. al., "Evaluation of autofocus functions in molecular cytogenic analysis," J. Microsc., 188(3), 264-272 (1997)
In particular, I found the functions "F4" and "F5", attributed to Vollath, to be robust.
Hope this helps.
John Minter
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 20 17:13:11 2004
That's a good point about the software needed to access these archived CD's remaining available. I now have a number of CD's of data that were made using Roxio's "Direct CD" that I do not have on my new computer. And Win XP's CD handler won't access the files. The data isn't so important for me to take the trouble to find Direct CD and load it onto my new computer just to off load data from old CD's. But it made me aware of the fact that Win XP's CD handler isn't going to be around forever either. When the new "Big Thing" shows up, we need to be ready to transfer anything that's important to the new media. Come to think of it, I have a bunch of stuff on Jazz cartridges too.
Joiner Cartwright, Jr., Ph.D. Baylor College of Medicine Houston, Texas U.S.A.
-----Original Message----- } From: michael shaffer [mailto:michael-at-shaffer.net] Sent: Saturday, September 18, 2004 7:09 AM To: MSA listserver
Gordon Couger writes ...
} : Of course, in 20 years, no drives will exists that will read any of } the above media, so it really doesn't matter how long the supposed } "archival" DVDs last, now does it. How many DECtape readers have -you-
} seen recently?
I don't know that I'd agree. Today's optical media and drives appear to offer a means for total backward compatibility ... e.g., DVD drive which can read CDs. This is of course up to the manufacturers sticking to it and supporting its application toward archiving.
That said ... I believe this thread should be as much about the dependability of the software. It isn't so much about the media ... e.g., if an archive was written today, what software would be able to restore it in 20 years? What can I use with confidence? ... which will write archives identical to the original directory structure, without splitting files, and verify the archive integrity??? Will the software maintain any database characteristics of the original collection of files?? What software would support and manage redundant archives across multiple devices (as Gary Gaugler suggested as required)??
cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com (in progress)
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 20 19:22:33 2004
Previously I worked for a GLP accredited CRO, and we had some guidelines on using CDs for backups. At that time I was the Systems Administrator (Bask in Zoology, go figure...) and wrote most of their Standard Operating Procedures. Generally this can be summed up:
- Buy gold or silver CD-Rs, and buy a reputable brand. 'Gold' or 'silver' CDs usually use a reactive metal coating to store data, rather than a photoreactive dye as seen in 'green' or 'blue' CDs. The metal based CD-R is more stable and (usually) better quality. I'd trust these for 5 years. Good brands are Kodak, Imation, Verbatium, TDK, Sony, Mitsubishi. Other brands may be good too, if you trust their VHS or Audio tapes then you can trust their CD-Rs.
- Never ever CR-RW. Untrustworthy and expensive. CD-Rs are cheap, use it, store it, throw it away. Better than losing data.
- Buy a good CD writer. This is hard for those not savvy with PCs. Buy a good brand, and not all the good brands are well-known brands. Asus, Sony, Pioneer, and Plextor are considered the best. Never use HP CD writers, though this is mostly from poor personal experience. Finally, buy a new one every two years, or more often if you use it lots. Make sure that it has a 2MB buffer or bigger (8MB is best) and some form of 'underrun protection'.
- Get good software and use it. Forget the widgets built into Windows 2000/XP, too easy to stuff up. Don't use DirectCD. Don't use ROXIO (some versions of Roxio software had some serious bugs that put your whole hard drive at risk! I'm not forgiving enough to start using their software again). I recommend Nero Burning ROM. It's cheap, easy, and often comes bundled with a CD-Writer. CloneCD is a good package for just copying CDs.
- Keep your CDs safe. A cool dry dark place. A cupboard is fine. Do not keep CDs in the fridge or freezer, some brands get very brittle at low temperatures and will shatter in the CD drive. Keep them out of direct sunlight and don't store them in a car.
Finally for the really paranoid. Don't use CDs, use tapes, or even better arrange for a reputable archiving company to do your backups for you and store them off site. If you have to use CDs, make multiple copies and keep one set off site. Use more than one brand of CD-R and alternate, so if one brand packs it in, the next month/week CD will still be Ok.
More will probably come to me later.
Aaron Hicks Electron Microscopy Preparation Technician
Comparative Physiology and Anatomy Institute of Veterinary, Animal, and Biomedical Sciences Massey University
PN-412 Private Bag 11 222 Palmerston North New Zealand
Phone +64 06 350 4470
At 03:46 PM 9/16/2004, you wrote:
} Dear colleague, } I remember some discussion about CD-R storage. Some CD are not good at } all and in many cases some CD become unreadable after 3 years. } I would like to know what is the best brand or type of CD for storing
} images } } Keep care and be of good cheer } } Regards } } (name) Vratislav Richard Eugene Maria John Baptist } (surname) of Bejsak (Bayshark)-Colloredo-Mansfeld
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 20 22:33:34 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (patljg-at-gwumc.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 20, 2004 at 11:47:56 ---------------------------------------------------------------------------
Question: Recently, I have been having problems with my thick sections lying flat. Nothing has been changed in the protocol; section, stain, and rinse. No matter how carefully I place the section on the slide, it will not be flat after the slide has dried.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jameen-at-rohmhaas.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 20, 2004 at 16:05:08 ---------------------------------------------------------------------------
Email: jameen-at-rohmhaas.com Name: Joseph G. Ameen
Organization: Rohm and Haas Electronic Materials
Title-Subject: [Microscopy] [Filtered] MListserver: electronic Magnetic Cancellation Systems
Question: Does any one know if the Electromagnetic Compensation Systems or Magnetic Active Compensation systems work on an SEM? Our company is about to purchase a new SEM and the site survey showed AC fields in the 11 - 29 milliGauss range and DC fields in the 2 mG range. Vibration testing was fine. The stray fields are at least an order of magnitude higher then the recommended levels. Has anyone had experience with these cancellation systems?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (cmeyer911-at-yahoo.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 20, 2004 at 22:23:28 ---------------------------------------------------------------------------
Email: cmeyer911-at-yahoo.com Name: Chris Meyer
Organization: Boeing
Title-Subject: [Microscopy] [Filtered] [TEM] Selected Area Diffraction ZA Deconvolution
Question: I would like to hear any suggestions as to what is the best free software or web-based solutions site, for de-convoluting zone axis for selected-area diffraction patterns based on angles and length ratios. I'm specifically looking for some help beyond the FCC, BCC and HCP (with Ti c/a ratio)that I find in textbooks.
RE: Plastic Membrane Boxes, Gel Sticky Box, Wafer Containers (see website below)
I am looking for containers of ion-milling samples and small pieces in progress of sample preparation. I found three types of containers: Membrance Box, Stickey gel box, and just small containers. Is there any suggestion, regarding advantage or disadvantage. I am using this when I trip by car or airplane.
I know that some people use TEM grid boxes that come with carbon films when you buy it. I don't like this because sometimes I break ion-thinned samples when I close the cover.
Lesley, I presume that you have already stained your semi-thin sections at that point, when you notice that the section is no longer flat. One thing that I have noticed, is that a person needs to wait a bit longer after the section has dried on the hot plate before they start staining. So when the section is drying, I am busy typing my slide labels, but even then, when the slide looks dry and ready to go, it isn't in fact ready to go, and you need to give it a few more minutes, otherwise you won't have a good bond of the section to the slide, and it will start to lift off during the staining, and especially the washing after the staining. This could be the most frustrating thing in the universe if this happens.
If on the other hand, you give your slide some time to BOND to the slide, (a few minutes) even after it has dried down, then the sections will refrain from lifting up and folding over during the staining process.
If you allow it this extra time, you can be quite rough with the slide during the staining and washing, and those sections won't lift off and fold. And not only that, but you won't have to use coated slides to make sure that the sections won't wash off, because they won't be going anywhere.
Garry
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (patljg-at-gwumc.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 20, 2004 at 11:47:56 ---------------------------------------------------------------------------
Question: Recently, I have been having problems with my thick sections lying flat. Nothing has been changed in the protocol; section, stain, and rinse. No matter how carefully I place the section on the slide, it will not be flat after the slide has dried.
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From MicroscopyL-request-at-ns.microscopy.com Tue Sep 21 13:11:54 2004
Hello, I have some Kodak EM film SO-163 and the expiration date was July 2003. It has been stored in the freezer. What are the odds that it is still useable? Any advice would be appreciated. thanks in advance, Beth
********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
"Between the two evils, I always pick the one I never tried before". Mae West (1893-1980) *******************************************************************
"And it's only the giving that makes you what you are". Wond'ring Aloud, Jethro Tull (Aqualung)
Hello Lesley, When I cut resin thick sections (0.5 - 5.0 microns) I transfer the sections to a drop of water on a slide and set the slide on a slide warmer under a large Petri dish cover with a cotton swab dipped in toluene. The toluene vapor softens and smooths the resin sections so that they flatten and adhere to the slide.
At 10:32 PM 9/20/2004 -0500, you wrote: } Email: patljg-at-gwumc.edu } Name: lesley graham } Organization: george washington university } Title-Subject: [Microscopy] [Filtered] MListserver: } Question: Recently, I have been having problems with my thick sections } lying flat. Nothing has been changed in the protocol; section, stain, and } rinse. No matter how carefully I place the section on the slide, it will } not be flat after the slide has dried. } Lesley Graham
Dean Abel Biological Sciences 143 BB University of Iowa Iowa City IA 52242-1324
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 21 14:55:49 2004
----- Original Message ----- } From: "Beth Richardson" {beth-at-plantbio.uga.edu} To: "microscopy microscopy" {microscopy-at-msa.microscopy.com} Sent: Tuesday, September 21, 2004 7:10 PM
On Sep 21, 2004, at 11:10 AM, Beth Richardson wrote:
} I have some Kodak EM film SO-163 and the expiration date was July } 2003. It has been stored in the freezer. What are the odds that it is } still useable? Any advice would be appreciated. } thanks in advance,
Dear Beth, When I was at Albany, we had both 4489 and LoDose films that had been refrigerated for some years when I got there, so was from the 70s or before. When I used either film and developed it according to instructions, there was no loss of contrast compared to fresh film, and the fog level--especially with the LoDose, which is about 1.5 orders of magnitude more sensitive than SO163--was not measurable; i.e., one could detect only a few grains in a 10 x 10 um field, and the microdensitometer reading was 0.000 OD. Unless the quality control has gotten much worse, I expect that the SO163 should be indistinguishable in response from brand new film. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 21 16:19:06 2004
Dear Hiromi, I use flat silicon rubber plates to hold ion-milled samples. These are flat slabs of silicon rubber that fit in a petri plate and the pieces or grids stick to them, then release them cleanly. They are available from Canemco (Canada) and probably other EM suppliers and are marked off in numbered squares. I have also made my own out of the castable silicon rubber kits. Good luck, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: {hkonishi-at-indiana.edu} To: {Microscopy-at-microscopy.com} Sent: Tuesday, September 21, 2004 9:10 AM
Hiromi: I like the membrane boxes for thin, non-mechanically strong things. They hold tightly but when you take the lid off, the parts can be handled w/ ease. I love the sticky-gel-type boxes for most everything else but things do STICK in them and can be damaged when trying to remove them. For what you're doing, I recommend the membrane boxes.
hkonishi-at-indiana.edu wrote:
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From MicroscopyL-request-at-ns.microscopy.com Tue Sep 21 17:46:49 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (donaldawbrey-at-texashealth.org) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, September 21, 2004 at 09:05:20 ---------------------------------------------------------------------------
Email: donaldawbrey-at-texashealth.org Name: Donald G. Awbrey HT(ASCP) QIHC
Organization: Harrris Methodist Hospital
Title-Subject: [Microscopy] [Filtered] Printers for diagnostic quality images.
Question: Dear fellow mico netters,
We are checking into purchasing a monochrome laser printer that will be used to print EM photos for diagnostic purposes.
The ones we are looking at are at 1200 X 1200 dpi and have a 16-32MB expanded memory. We are not neccessarily interested in print speed.
We will be using a 2K camera on our TEM.
Will this printer give us the quality for photographs that will be used for pathology diagnosis in a clinical setting? Will these prints be of good quality to pass any laboratory inspection (i.e. CAP)?
Hi Karen, There was some discussion of microwave protocols for insect larvae in January 2004. Try the Listserver archives for these. Myself, I found that much trial and error was necessary. Some things you may want to try include:
- Karnovsky's fixative (glutaraldehyde & paraformaldehyde in phosphate buffer) - injecting the fixative into the specimen - using the vacuum (I draw the vacuum to 20" Hg and leave it for 5 minutes before MWing. MW with vacuum on.)
Using cooled Karnovsky's, I draw up a vacuum, leave it for 5 min and then MW with an initial sequence at 100 W (2 min 0% power, 2 min 100% power, 3 min 0% power), cool my specimen to 20 deg C, draw up a vacuum and MW for a second sequence at 450 W (30 sec 0% power, 30 sec 100% power, 30 sec 0% power). I repeat the 450 W sequence 3 times. After I let my specimens sit in fix for 5 min.
For my osmium step, with a vacuum and cooled solution, I MW at 100 W for a sequence of 2 min 0% power, 2 min 100% power, 3 min 0% power. I also let the samples sit in osmium for a few minutes (10-20 min).
I've used this on larvae larger than mosquitos (specifically, diamond back moth and Colorado potato beetle) and it has yielded good results. I've also found that processing several samples will yield some with good quality fixation and the others can be discarded.
If you have any questions, feel free to contact me. I'd be interested in hearing how things worked out for you.
Shannan
Shannan Little Research Technician/Technicien de recherche Electron Microscopy and Image Analysis /Microscopie électronique et Analyse d'images Agriculture and Agri-Food Canada/Agriculture et Agroalimentaire Canada Telephone/Téléphone: 403-317-3446 Facsimile/Télécopieur: 403-382-3156 P.O. Box 3000 / CP 3000 Lethbridge, Alberta T1J 4B1 littlesm-at-agr.gc.ca http://res2.agr.ca/lethbridge/emia/index_e.htm
-----Original Message----- } From: Karen Kelley [mailto:klk-at-biotech.ufl.edu] Sent: Thursday, September 16, 2004 4:06 PM To: Microscopy-at-MSA.Microscopy.Com
Hello All,
I have a BioScan Ted Pella microwave with a cold-spot and vacuum chamber that I would like to use for fixation and embedding insect eggs and larvae for TEM.
The project includes mosquito larvae, pupae and fly eggs. I worked on this same project many years ago using traditional benchtop methods. The eggs and larvae were difficult, if not impossible, to infiltrate. Now that I have a MW with vacuum I am trying this project again.
I would greatly appreciate any suggestions, advice or direct me to references?
My first attempt at MW didn't go so well. I MW fixed the larvae with 2% glut in cacodylate in 1.5ml centrifuge tubes at 20mmHg vacuum -at- 250W 1' on, 1' off. Repeated several times removing the vacuum to observe the larvae. At first when vacuum was applied the larvae floated to the top and squirmed around a bit, then after several attempts at that they finally all stopped moving and didn't float. Buffer washed on benchtop then used the vacuum again at the same settings for the buffered osmium tetroxide. Just the ends of the larvae turned black. I tried it again without any change. I finally got tired of messing with them and placed them into a KFeCN + OsO4 overnight at 4C and have had them stored in buffer. ( I have used the overnight KFeCN + OsO4 with success for hard to infiltrate nematodes). Dehydrated with MW 40s -at- 250W no vacuum 40 s on, 1' off, infiltrated 50% acetone/Embed no vac at 250W 3', 100% Embed -at- 450W with vac (20mm Hg). The larvae had areas of collapse after the 100% with vacuum so I'm concerned that they are not infiltrating as I had hoped.
Karen L. Kelley ICBR Electron Microscopy Manager University of Florida ICBR Electron Microscopy Core Lab Bartram Hall Room 214 Box 118525 Gainesville Florida Lab: 352-392-1184 fax: 352-846-0251 Email: klk-at-biotech.ufl.edu Southeastern Microscopy Society Treasurer http://www.biotech.ufl.edu/EM/
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 21 19:12:31 2004
I also swear by the membrane boxes. I use the small 1 x 1 size. For very brittle samples, such as II-Vi and III-V compounds, I don't put the membrane boxes together with full force. Just tight enough so that they can't go from edge to edge. I also re-use them. I use 1" white Post-it tape and tear off long enough to use as a label.
One thing that you must be very careful with using these and that is that the sample must be thoroughly dried from any solvents that you may have used, especially acetone. If the sample is even slightly damp when you put it in, it will stick to the membrane.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472 Walck-at-PPG.com (412) 820-8651 (office) (412) 820-8515 (fax)
-----Original Message----- } From: hkonishi-at-indiana.edu [mailto:hkonishi-at-indiana.edu] Sent: Tuesday, September 21, 2004 12:11 PM To: Microscopy-at-microscopy.com
RE: Plastic Membrane Boxes, Gel Sticky Box, Wafer Containers (see website below)
I am looking for containers of ion-milling samples and small pieces in progress of sample preparation. I found three types of containers: Membrance Box, Stickey gel box, and just small containers. Is there any suggestion, regarding advantage or disadvantage. I am using this when I trip by car or airplane.
I know that some people use TEM grid boxes that come with carbon films when you buy it. I don't like this because sometimes I break ion-thinned samples when I close the cover.
When we use our TEM at low magnification (below 4,000X), we sometimes get a bright spot in the middle of pictures we take. I always thought the microscope setting was the only thing responsible for this problem. But when I checked into a latest episode, I found pictures from one particular specimen (thin sections on Formvar coated grid) had it, but others from a different specimen (negative staining on Farmvar/carbon coated grid) did not, even though all pictures were taken with the same scope setting. Has anyone else experienced the same? Can anyone explain to me thoroughly how this problem occurs, and how to avoid it. Thank you very much in advance.
Hong Emory EM
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 22 07:45:41 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dieter-at-genetik.uni-bielefeld.de) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 22, 2004 at 06:14:11 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] MListserver: Which Deconvolution Software?
Question: Hello Listers!
We are currently searching for a deconvolution software. Mainly we look at GFP-fusions in single cells, monolayers (Sf9) or cell aggregates (tobacco BY2), up to 100 µm on a Olympus IX81.
What are your experiences with such a software tool? How are the results compared to confocal images of the same specimen (...okay, deconvolution can be applied to confocal images too)? Is it worth the money?
The two major competitors in the market are SVI/Huygens and AutoQuant. Which product did you choose and why? Is there a side by side test of both softwares available? Do they differ in image quality? We only need deconvolution, since our imaging software is doing well. Do you have other recommendations/suggestions?
Any comment on this issue will be warmly welcomed! Thank you in advance,
Pacific Northwest Microscopy Society (PNMS) announces its annual meeting held this year in Hillsboro, OR during two half-day sessions on September 30 and October 1st.
The FEI company generously offered its new facility for the meeting location, and largely contributed to its program. Two parallel platform sessions will run in biological and material sciences. Besides the scientific program, demonstrations of the FIB and a Technai TEM will be available, as well as the FEI factory tour. The evening social at the historical Cornelius House and Brewery will include a barbecue dinner, and it is covered with the registration. The second day morning biological session will take place in the OSHU Primate Center including its tour. Due to the space limitation (50 people), we require advanced registration. Registration is free of charge to the current PNMS members, and $10 for non-members.
Program:
Fall 2004 Annual Meeting When: Thursday Sept. 30th, and Friday October 1st, 2004 Where: FEI Company, Hillsboro, Oregon OHSU National Primate Research Center
Thursday Sept 30, 2004 11:30am - Arrival to FEI, registration 12:00 - Lunch 12:30 - Welcome note - 2 keynote speakers provided by FEI Speaker I - DualBeam Applications in Biological and Materials Science Speaker II - Tomography Applications in Biological and Materials Science 1:45pm - break
2:00 - Demonstrations of DualBeam Focused Ion Beam (FIB) instrument, and the Tomography capability on a Technai TEM (split into 2 groups) 2:45 - break
3:00 - Split into 2 sessions - Biological and Material Science platforms will run in parallel.
Material Science: 3:00 - Jun Jiao, Portland State University 3:20 - Joe Robinson Ascend 3:40 - David Basile, HP 4:00 Break 4:10 - Barbara Miner, Intel 4:30 - Eric Sanchez, Portland State U 4:50 - Jeff McDowell, Rontec
Biological Sciences: 3:00 - Elaine Humphrey, University of British Columbia - "Microwave Processing In a Modern Microscopy Facility" 3:30 - Glen MacDonald, University of Washington Seattle, Virginia Merrill Bloedel Hearing Research Center 4:00 - mini workshop: "Coloring Electron Micrographs With PhotoShop" presented by the image processing guru Jim Young
5:15 -6:00 - Poster session We are soliciting poster presentations in both areas, and strongly encouraging students to participate in the poster session. 6:00 - Dinner social at the Cornelius Pass Road House and Brewery (McMenamins). Poster winners announcement.
Friday Oct. 1, 2004 9:00 Arrival at FEI 9:15 - FEI factory tour (!!!), talk: Emitters and Optics by FEI Beam Technology. 10:15am Depart from FEI, commute (only minutes drive) to the OSHU's Primate Research Center. 10:30 - 11am Tour of the Primate Center. 11am -2pm Biological session at the conference room, tour of Imaging and Morphology Core Facility (Anda Cornea lab director). Lunch will be arranged in the cafeteria within the PC. 2pm - departure.
Alice Dohnalkova PNMS President Environmental Microbiology Pacific Northwest National Laboratory Richland, WA 99352 (509) 372-0692 office (509) 376-3654 TEM lab
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 22 12:20:41 2004
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I work for JEOL in the UK. We use them fairly frequently and they generally work pretty well.
Normally, the standard systems only cancel AC fields - DC cancellation is usually an option, although it is very rare that DC fields need to be cancelled - the SEM alignment does that for you unless the DC field is very high.
You need to look at the wave form of the AC fields - a coil and an oscilloscope is all you need. If the wave form is fairly stable and changes only relatively slowly, then cancellation will work well. If it is changing a lot, then you may have problems - the control box on the system we use has a LED to indicate when the field is cancelled or if it is still calculating the wave form. In some cases, it will help if you have two sensor boxes - these need to be placed as near to the sample as possible, usually on either side of the chamber. -- Larry Stoter PLEASE NOTE 1. Any mail other than plain text will be automatically deleted. 2. Any mail, legitimate or not, apparently or actually from hotmail, netscape, yahoo or excite will automatically be deleted. 3. Mail with no subject or without a clear subject will be ignored :-)
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 22 12:20:23 2004
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
If you want top quality photo-standard prints, you really need a dye-sublimation printer. Quite a bit more expensive to purchase and to run than a laser printer but the results are much better. Codonics make several printers aimed specifically at the medical imaging market. -- Larry Stoter PLEASE NOTE 1. Any mail other than plain text will be automatically deleted. 2. Any mail, legitimate or not, apparently or actually from hotmail, netscape, yahoo or excite will automatically be deleted. 3. Mail with no subject or without a clear subject will be ignored :-)
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 22 13:15:45 2004
} When we use our TEM at low magnification (below 4,000X), we } sometimes get a bright spot in the middle of pictures we take. I } always thought the microscope setting was the only thing responsible } for this problem. But when I checked into a latest episode, I found } pictures from one particular specimen (thin sections on Formvar } coated grid) had it, but others from a different specimen (negative } staining on Farmvar/carbon coated grid) did not, even though all } pictures were taken with the same scope setting. Has anyone else } experienced the same? Can anyone explain to me thoroughly how this } problem occurs, and how to avoid it. Thank you very much in advance. } } Hong } Emory EM
The most likely cause is removal of the plastic due to the beam -- resulting in an area of lowered density. This is quite common (and can sometimes be used to enhance the contrast of sectioned specimens). What is probably happening is that someone is investigating the specimen at high magnification with the beam condensed to a small spot. When you then go to a lower magnification, you will see the area where the beam spot has etched away the plastic. Also, some people focus the image with the condenser reduced in size (since it much brighter). If too much time is spent in this condition, you will see the etching of the plastic after the beam is spread.
One work-around would be to record the images at low mag FIRST and then do the high mag work (where the condenser is reduced in size) AND avoid focusing at crossover.
We sometimes use this with specimens of low contrast (LR White and Spurr's, for example). As the plastic is removed by the beam, the remaining specimen appears with greater contrast. In fact, our TEM even has this capability programmed in by the manufacturer (the beam is traversed over the specimen to "stabilize" the plastic--and it also improves contrast). -- ############################################################## John J. Bozzola, Ph.D., Director I.M.A.G.E. (Integrated Microscopy & Graphics Expertise) 750 Communications Drive - MC 4402 Southern Illinois University Carbondale, IL 62901 U.S.A. Phone: 618-453-3730 Email: bozzola-at-siu.edu Web: http://www.siu.edu/~image/ ##############################################################
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 22 13:17:35 2004
} When we use our TEM at low magnification (below 4,000X), we } sometimes get a bright spot in the middle of pictures we take. I } always thought the microscope setting was the only thing responsible } for this problem. But when I checked into a latest episode, I found } pictures from one particular specimen (thin sections on Formvar } coated grid) had it, but others from a different specimen (negative } staining on Farmvar/carbon coated grid) did not, even though all } pictures were taken with the same scope setting. Has anyone else } experienced the same? Can anyone explain to me thoroughly how this } problem occurs, and how to avoid it. Thank you very much in advance. } } Hong } Emory EM
The most likely cause is removal of the plastic due to the beam -- resulting in an area of lowered density. This is quite common (and can sometimes be used to enhance the contrast of sectioned specimens). What is probably happening is that someone is investigating the specimen at high magnification with the beam condensed to a small spot. When you then go to a lower magnification, you will see the area where the beam spot has etched away the plastic. Also, some people focus the image with the condenser reduced in size (since it much brighter). If too much time is spent in this condition, you will see the etching of the plastic after the beam is spread.
One work-around would be to record the images at low mag FIRST and then do the high mag work (where the condenser is reduced in size) AND avoid focusing at crossover.
We sometimes use this with specimens of low contrast (LR White and Spurr's, for example). As the plastic is removed by the beam, the remaining specimen appears with greater contrast. In fact, our TEM even has this capability programmed in by the manufacturer (the beam is traversed over the specimen to "stabilize" the plastic--and it also improves contrast). -- ############################################################## John J. Bozzola, Ph.D., Director I.M.A.G.E. (Integrated Microscopy & Graphics Expertise) 750 Communications Drive - MC 4402 Southern Illinois University Carbondale, IL 62901 U.S.A. Phone: 618-453-3730 Email: bozzola-at-siu.edu ##############################################################
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 23 05:45:26 2004
FYI, here is a current job opening at the GE Global Research Center in Niskayuna, NY. To apply, please go to www.gecareers.com {http://www.gecareers.com} and enter job #372689.
Materials Scientist/ Microscopist
Business Unit:
GE Global Research
Function:
Engineering/Technology
Location:
NISKAYUNA, NY
Job #:
372689 Posted: Aug 12, 2004
Job Description: Responsibilities
The Microstructural and Surface Sciences Laboratory is involved in research into the structure and composition of materials in support of development programs both at GEGR and at GE businesses. Staff members are expected to work independently with a high level of expertise, and to become involved with a number of major project teams. The successful candidate will execute research projects using advanced electron beam analysis of bulk materials (e.g., HRSEM, EDS, EBSD, FIB) to study the structure and composition of materials, including metals, ceramics, composites, polymers, and electronic materials. Lead in the development of new advanced techniques for materials characterization. Participate in or lead project teams with the goal of developing new processes and materials.
Qualifications
BS required, MS preferred in Materials Science, Chemistry, or Physics. Extensive training and experience in cutting-edge materials characterization techniques, particularly electron beam techniques. Demonstrated technical leadership capabilities with excellent teaming and communication skills. THIS POSITION REQUIRES UNRESTRICTED U.S. WORK AUTHORIZATION (US citizen or permanent resident status required).
General We offer a competitive salary, outstanding benefits package and the professional advantages of an environment that supports your development and recognizes your achievements. We are an Equal Opportunity Employer.
**************************************************************************** ************************************** Katharine Dovidenko, Ph.D. Materials Scientist Microstructural and Surface Sciences Laboratory GE Global Research Center K-1-2C12, 1 Research Circle Niskayuna, NY 12309 Phone: 518-387-4759 Fax: 518-387-6972 Email: dovidenk-at-research.ge.com {mailto:dovidenk-at-research.ge.com}
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 23 08:34:28 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (rdesilva-at-ipn.mx) from on Wednesday, September 22, 2004 at 08:59:35 ---------------------------------------------------------------------------
Email: rdesilva-at-ipn.mx Name: Claudia Douriet
Organization: CIIDIR
Education: Graduate College
Location: Guasave, Sinaloa, MÈxico
Question: Hi! I just started to use a Leica stereomicroscope for my Bachelor degree work and unfortunately I never used one before. I am taking pictures of zooplanktonic organisms and I dont know how much magnification does a picture has. I am not sure if you have to multiply the zomm, by the objective and by the ocular numbers. Can you help me? Thank you very much.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (zerfasp-at-ors.od.nih.gov) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 22, 2004 at 15:24:37 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] MListserver: Bone Fixation
Question: Does anyone have a protocol for fixing mouse bone tissue? The investigator does not have a microwave nor does she plan to profuse the animals. Thanks.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (john.dumont-at-ctel.net) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 22, 2004 at 16:26:34 ---------------------------------------------------------------------------
Question: I'm presently refurbishing an older Olympus Model EHT with a DO head. I require a lower and upper lamp assemblies, as well as an owner/operator's manual.
Thank you for your input. I definitely have seen this removal of the plastic phenomenon on scope. But I am not convinced that is all what was happening here. The spot was there even if we moved away from etched area. Also, the spot sometimes was so intense that it could be seen on phosphor screen. Someone suggested that it has something to do with the distance between the pole piece and the objective aperture. I think I agree, but it seemed that the type of sample prep played a role in it too.
Thank you all.
Hong
On Sep 22, 2004, at 2:14 PM, John J. Bozzola wrote:
} } } ----------------------------------------------------------------------- } ------- } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ----------------------------------------------------------------------- } -------- } } } When we use our TEM at low magnification (below 4,000X), we } } sometimes get a bright spot in the middle of pictures we take. I } } always thought the microscope setting was the only thing responsible } } for this problem. But when I checked into a latest episode, I found } } pictures from one particular specimen (thin sections on Formvar } } coated grid) had it, but others from a different specimen (negative } } staining on Farmvar/carbon coated grid) did not, even though all } } pictures were taken with the same scope setting. Has anyone else } } experienced the same? Can anyone explain to me thoroughly how this } } problem occurs, and how to avoid it. Thank you very much in advance. } } Hong } } Emory EM } } The most likely cause is removal of the plastic due to the beam -- } resulting in an area of lowered density. This is quite common (and can } sometimes be used to enhance the contrast of sectioned specimens). } What is probably happening is that someone is investigating the } specimen at high magnification with the beam condensed to a small } spot. When you then go to a lower magnification, you will see the area } where the beam spot has etched away the plastic. Also, some people } focus the image with the condenser reduced in size (since it much } brighter). If too much time is spent in this condition, you will see } the etching of the plastic after the beam is spread. } } One work-around would be to record the images at low mag FIRST and } then do the high mag work (where the condenser is reduced in size) AND } avoid focusing at crossover. } } We sometimes use this with specimens of low contrast (LR White and } Spurr's, for example). As the plastic is removed by the beam, the } remaining specimen appears with greater contrast. In fact, our TEM } even has this capability programmed in by the manufacturer (the beam } is traversed over the specimen to "stabilize" the plastic--and it also } improves contrast). } -- } ############################################################## } John J. Bozzola, Ph.D., Director } I.M.A.G.E. (Integrated Microscopy & Graphics Expertise) } 750 Communications Drive - MC 4402 } Southern Illinois University } Carbondale, IL 62901 U.S.A. } Phone: 618-453-3730 } Email: bozzola-at-siu.edu } Web: http://www.siu.edu/~image/ } ############################################################## } } ====================== Hong Yi Emory School of Medicine Microscopy Core Emory University 6215 Woodruff Memorial Research Building 101 Woodruff Circle Atlanta, GA 30322
} } When we use our TEM at low magnification (below 4,000X), we } } sometimes get a bright spot in the middle of pictures we take. I } } always thought the microscope setting was the only thing } } responsible for this problem. But when I checked into a latest } } episode, I found pictures from one particular specimen (thin } } sections on Formvar coated grid) had it, but others from a } } different specimen (negative staining on Farmvar/carbon coated } } grid) did not, even though all pictures were taken with the same } } scope setting. Has anyone else experienced the same? Can anyone } } explain to me thoroughly how this problem occurs, and how to avoid } } it. Thank you very much in advance. } } Hong } } Emory EM } ============= } The most likely cause is removal of the plastic due to the beam -- } resulting in an area of lowered density. This is quite common (and } can sometimes be used to enhance the contrast of sectioned } specimens). What is probably happening is that someone is } investigating the specimen at high magnification with the beam } condensed to a small spot. When you then go to a lower } magnification, you will see the area where the beam spot has etched } away the plastic. Also, some people focus the image with the } condenser reduced in size (since it much brighter). If too much time } is spent in this condition, you will see the etching of the plastic } after the beam is spread. } } One work-around would be to record the images at low mag FIRST and } then do the high mag work (where the condenser is reduced in size) } AND avoid focusing at crossover. } } We sometimes use this with specimens of low contrast (LR White and } Spurr's, for example). As the plastic is removed by the beam, the } remaining specimen appears with greater contrast. In fact, our TEM } even has this capability programmed in by the manufacturer (the beam } is traversed over the specimen to "stabilize" the plastic--and it } also improves contrast). } } John J. Bozzola, Ph.D., Director } I.M.A.G.E. (Integrated Microscopy & Graphics Expertise) } 750 Communications Drive - MC 4402 } Southern Illinois University } Carbondale, IL 62901 U.S.A. } Phone: 618-453-3730 } Email: bozzola-at-siu.edu } ######################## Hong, I knew what you were referring to on the low power magnification and just checked it out on my old Philips 200.
The spot is evident on the screen when in low mag. and hence would be on the negative if I took a picture. It even shows up on the grid bar when I scan ONLY when the image is out of focus!
If you have a WOBBLER on your scope use it to get into focus and see if the light spot disappears. If you do not have a wobbler, put in a holey grid or of course a thicker section that has a hole or two and focus on the hole to the point where it is in focus and look for the spot. I bet it is no longer there.
Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu ***********************
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 23 13:26:26 2004
Greetings all, I'm helping with a project which requires dehydration/clearing of the specimen for observation using laser scanning using both reflected light and fluorescence detection. Also, we need to minimize mounting agent shrinkage in order to preserve the z axis aspect ratio of the specimen. I've suggested using an ethanol dehydration followed by clearing and mounting in clove oil, but I'm not sure what problems we might encounter from auto-fluorescence of the clove oil. Oil of wintergreen would work also, I've just read that clove oil is a little more tolerant of incomplete dehydration (and seeing as I'm not performing the dehydration I figured a little margin for error would be a good idea). Any input on the relative merits of clove oil/oil of wintergreen or even immersion oil as a penetrating mounting agent would be appreciated. Thanks. -Karl
-- Karl Garsha Light Microscopy Specialist Imaging Technology Group Beckman Institute for Advanced Science and Technology University of Illinois at Urbana-Champaign 405 North Mathews Avenue Urbana, IL 61801 Office: B650J Phone: 217.244.6292 Fax: 217.244.6219 Mobile: 217.390.1874 www.itg.uiuc.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 23 14:42:10 2004
We use clove oil routinely to clear spider genitalia that is preserved in ethanol 75%, without any previous dehydration. It works very well. Larger pieces take longer, but I guess you will be working with very small preparations.
Other labs use lactic acid or methyl salicylate, but I prefer clove oil.
Martín J. Ramírez División Aracnología Museo Argentino de Ciencias Naturales Av. Angel Gallardo 470 C1405DJR Buenos Aires Argentina tel +54 11 4982-8370 int. 168 fax +54 11 4982-4494
At 03:24 PM 9/23/2004, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Thu Sep 23 21:47:43 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Tammy-at-kbwiz.com) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Thursday, September 23, 2004 at 16:19:04 ---------------------------------------------------------------------------
Email: Tammy-at-kbwiz.com Name: Tammy Schwalb
Organization: Schwalb Research
Education: Graduate College
Location: Irvine, CA
Question: I am lookig for an embedding media for a delicate tissue membrane encasing a small sensor surface. The device material is epo fix. I would prefer a hydrophilic material that is easy to cut. Any advice?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (spradhan-at-siu.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Thursday, September 23, 2004 at 12:30:09 ---------------------------------------------------------------------------
Question: i have been thinking about building an experimental setup to test the mechanical properties of thin films under loading using speckle pattern interferometry with electron microscopy...i would like to observe the speckle patters using the electron microscopy facility we have here(S-2460N hitachi)...it seems however that i might have problems placing the loading unit inside the chamber primarily owing to the contract they have here regarding the maintainence of the microscope with hitachi and also due to the fact that there arent supposed to be any EM fields or unapproved active devices inside the chamber...in this regard, i would be glad if you could help me out by providing ways to circumvent the problems involved....have people done that kind of work before? if so then how....how can u replace the motor, for example, to load the specimen? i shall look forward to your reply...
We are currently using the Slidebook by Inteligent Imaging Inc. (3I) with an Olympus IX-81. [http://www.intelligent-imaging.com/slidebook/overview.php ]
It integrates and operates the IX-81 fully, comes with all its available camera drivers, and does little other than deconvolution. 3I seems to be readily working with the camera vendors and Olympus to continually update the drivers and software. I think it maybe a little less costly than the others but came as a larger package deal. I've not tried the others - Slide book seems to work well - makes use of multiple processors and available memory (4GB RAM) to run pretty fast.
Deconvolution: We're just basic users in deconvolution but it does seem to me that there has been plenty of published justification that the transition from confocal to deconvolution is at 15-micrometers, i.e. below 15 deconvolution is better, thicker than 15 confocal is better. For 100um samples you maybe far better served by confocal.
} } Email: dieter-at-genetik.uni-bielefeld.de } Name: Dieter Kapp } } Organization: Academic/University } } Title-Subject: [Microscopy] [Filtered] MListserver: Which Deconvolution } Software? } } Question: Hello Listers! } } We are currently searching for a deconvolution software. } Mainly we look at GFP-fusions in single cells, monolayers (Sf9) or cell } aggregates (tobacco BY2), up to 100 µm on a Olympus IX81. } } What are your experiences with such a software tool? } How are the results compared to confocal images of the same specimen (...okay, } deconvolution can be applied to confocal images too)? Is it worth the money? } } The two major competitors in the market are SVI/Huygens and AutoQuant. Which } product did you choose and why? Is there a side by side test of both softwares } available? Do they differ in image quality? We only need deconvolution, since } our imaging software is doing well. Do you have other } recommendations/suggestions? } } Any comment on this issue will be warmly welcomed! } Thank you in advance, } } Dieter } } ------------------------------------------------------------ } Dr. Dieter Kapp } Lehrstuhl fuer Genetik } Postfach 100131 } Universitaet Bielefeld PHONE: +49 (0)521 106 5620 } D-33501 Bielefeld FAX: +49 (0)521 106 5626 } GERMANY Email: dieter-at-genetik.uni-bielefeld.de } } ------------------------------------------------------------ } } } } --------------------------------------------------------------------------- } }
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Director 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 08:10:01 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (John H. Cross-at-lmco.com) from http://www.microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, September 23, 2004 at 13:31:31 ---------------------------------------------------------------------------
Email: John H. Cross-at-lmco.com Name: John H. Cross, CIH
Organization: Lockheed Martin Space Operations
Title-Subject: [Microscopy] [Filtered] MListserver: Occupational Health Aspects of Microscope Usage
Question: We have a group of technicians using microscopes to solder electronic components 10-12 hours per day 6-7 days a week.
Does any professional, academic, or industry organization publish occupational health guidelines for the use of microscopes? A sample of questions I need to address are: How often must an individual take a break from looking through a microscope. Is exposure to light coming through the eyepieces a problem? Has anyone described ergonomically-correct work stations for microscopists?
All comments will be welcome. I particularly need objective standards that I can present to management to justify work practice adjustments.
When doing semi-thin sectioning I face off my block to cut, when the specimen comes across the knife it takes an obscenely thick section and ruins my block face. I have had problems before where the first section is a little too thick, but simply backing the knife off a little bit helped. No matter what I do or change the section is still cutting too thick, are there any suggestions before I go crazy!
Thanks,
Todd M. Hamm Oklahoma Medical Research Foundation 825 NE 13th St OKC, OK 73132 tmhamm09-at-yahoo.com
what sort of microtome are you using? If it's an LKB/Reichert ultramicrotome you could be aligning specimen and knife when the knife is retracted - this is part of the normal cutting cycle to prevent the block snagging the knife when it returns to the top of its travel. When you complete the first cutting cycle the microtome knife stage jumps forward at least 10s of microns. So when setting up for semi-thin sectioning make sure that this isn't happening - it's quite easy to do when you know how.
I haven't gone into detail in case this is not your problem, but it should be easy enough to check. Let me know if you need more info.
Malcolm
Malcolm Haswell e.m. unit School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear SR1 3SD UK e-mail: malcolm.haswell-at-sunderland.ac.uk
----- Original Message ----- } From: todd hamm {ripbnowell-at-hotmail.com}
Dear Claudia,
The best way to define magnification is to set-up the microscope to a prescribed magnification and zoom (one which is reproducible) then to take a picture of either a ruler with mm or a stage micrometer. If you are viewing the image on a screen, you can lay an overhead transparency on the screen and draw the length of some convenient "marker bar" (ex: 50 micrometers). You will then always have the magnification available. Also, if you are going to capture digital images, most software gives you the ability to calibrate the magnification.
Because of the nature of both the focusing and zoom mechanism on the stereo microscope, it is more difficult to establish exact magnification, so you may need to repeat this calibration for each test run.
Good hunting! Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^& Need a good general text on light microscopy? MME still has copies of Optimizing Light Microscopy available, with discounts for class-sized orders (10 or more). Visit www.MicroscopyEducation.com for details.
At 08:33 AM 9/23/2004, rdesilva-at-ipn.mx wrote:
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From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 12:22:53 2004
The best way to define magnification is to set-up the microscope to a prescribed magnification and zoom (one which is reproducible) then to take a picture of either a ruler with mm or a stage micrometer. If you are viewing the image on a screen, you can lay an overhead transparency on the screen and draw the length of some convenient "marker bar" (ex: 50 micrometers). You will then always have the magnification available. Also, if you are going to capture digital images, most software gives you the ability to calibrate the magnification.
Because of the nature of both the focusing and zoom mechanism on the stereo microscope, it is more difficult to establish exact magnification, so you may need to repeat this calibration for each test run.
Good hunting! Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^& Need a good general text on light microscopy? MME still has copies of Optimizing Light Microscopy available, with discounts for class-sized orders (10 or more). Visit www.MicroscopyEducation.com for details.
At 08:33 AM 9/23/2004, rdesilva-at-ipn.mx wrote:
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From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 12:29:05 2004
When doing semi-thin sectioning I face off my block to cut, when the specimen comes across the knife it takes an obscenely thick section and ruins my block face. I have had problems before where the first section is a little too thick, but simply backing the knife off a little bit helped. No matter what I do or change the section is still cutting too thick, are there any suggestions before I go crazy! ______________________________________
Todd, Could you give a little more info?
What microtome are you using? The "newer" ones (less than 20 years old) have back lighting that makes approaching the block so much easier.
I know that this may be a silly question, but are you sure that everything is tight? (knife stage, knife holder, specimen chuck, specimen arm)
How are you approaching the block face? I normally creep up on it slowly using the knife advance while watching through the binoculars until I'm shaving off little bits. Then I set up my section thickness and cut a few thicks manually (rather than with the motor drive).
When you bring your knife up to the block are you sure that the specimen arm is in the cutting stroke and not in the retracted phase of the cycle?
Have you had your microtome checked to be sure that the calibration for the thick sectioning mode is OK?
That pretty much covers all of the mistakes I've made over the years... Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 12:30:50 2004
Again, this might be better done as a scanning probe experiment rather than an SEM experiment. You can readily measure force, adhesion, and viscoelasticity directly.
Contact me offline for details.
Best regards, Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^&^& Need a good general text on light microscopy? MME still has copies of Optimizing Light Microscopy available, with discounts for class-sized orders (10 or more). Visit www.MicroscopyEducation.com for details.
At 09:47 PM 9/23/2004, spradhan-at-siu.edu wrote:
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From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 13:56:58 2004
_________________________________________________________________ Get ready for school! Find articles, homework help and more in the Back to School Guide! http://special.msn.com/network/04backtoschool.armx
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 14:07:15 2004
Sorry I was a little vague before, the ultramicrotome is a fairly new LKB model. I found that when I take the semi-thins manually the problem isn't as bad. Is it possible that the belt that controls the specimen advancement has been worn, I thought that might be possible since the thickness varies considerably from section to section. But like I said when section manually it is much less of a problem.
Todd Hamm
_________________________________________________________________ Don’t just search. Find. Check out the new MSN Search! http://search.msn.click-url.com/go/onm00200636ave/direct/01/
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 14:39:01 2004
We're looking at mouse embryos in the early stages using quadrature tomographic microscopy. What I want to do is rotate an embryo in specific increments and image at these increments at various time points (total ‰ 48 hrs). This will be done in a Bioptechs chamber (no affiliation) with additional atmospheric control.
My first thought is to "attach" the embryo to a "tube" via a syringe, with slight negative pressure. Then, attach a goniometer somehow to the tube or the syringe.
These lists have been extremely useful in the past and I thank you all in advance for your input.
Best,
Gary --
Gary Laevsky, Ph.D. Keck Facility Manager, CenSSIS Northeastern University 302 Stearns 360 Huntington Ave. Boston, MA 02115
voice (617) 373-8570 fax (617) 373-7783
From MicroscopyL-request-at-ns.microscopy.com Fri Sep 24 17:51:40 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (timandcathy-at-mho.com ) from http://www.msa.microscopy.com/Ask-A-Microscopist/Ask-A-Micro-Form.html on Friday, September 24, 2004 at 13:04:20 ---------------------------------------------------------------------------
Question: Hi I am looking for a microscope -at- 1600x to look at bacteria, I have seen oil immersion does the microscope need a oil lense or can you just use the oil on the slides ? and any ideas on good brands for the max power ?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (necipunlu-at-yahoo.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, September 24, 2004 at 08:51:18 ---------------------------------------------------------------------------
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (rangari0-at-yahoo.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, September 24, 2004 at 22:21:53 ---------------------------------------------------------------------------
The header is a text field which describes how the data is formated. You should be able to read the entire spectrum in any text editor. XY columns is only one of many possible formats which are legal forms for the data set to be output.
The program that is exporting the data into the MSA-MAS file format should ask the individual how many columns and what format to export the file into. It sounds like the person requested 4 column format. I'd suggest that you read the header closely it should explain how the data is output. You can then edit it as appropriate.
Alternatively have the data re-exported into XY format.
Title :EMSAMASFF Keywords :XEDS,EELS,AES,WDS,CLS,GAM,XRF,PES Computer :IBM, MAC, DEC Operating System :ALL Programming Language :Fortran 77 Hardware Requirements :None Author(s) :EMSA/MAS TASK FORCE Ray Egerton ,Charles E. Fiori ,John A. Hunt ,Mike S. Isaacson, Earl J. Kirkland ,Nestor J. Zaluzec Correspondence Address : R.F. EGERTON CHAIRMAN University of Alberta Dept. of Physics Alberta, Canada, Abstract:
A simple format for the exchange of digital spectral data is presented, and proposed as an EMSA/MAS standard. This format is readable by both humans and computers and is suitable for transmission through various electronic networks (BITNET, ARPANET), the phone system (with modems) or on physical computer storage devices (such as floppy disks). The format is not tied to any one computer, programming language or computer operating system. The adoption of a standard format would enable different laboratories to freely exchange spectral data, and would help to standarize data analysis software. If equipment manufacturers were to support a common format, the microscopy and microanalysis community would avoid duplicated effort in writing data-analysis software. This version of EMSAMASFF contains two subroutines which read and write spectral data files Version 1.0 data format. The data are stored as simple ASCII characters at a user defined number of columns per line for the length of the data file. The spectral data is preceeded by a series of header lines, which tell the user about the parameters of the spectrum. The header lines are identified by the first character in the line being the symbol (#) followed by a descriptor and if appropriate its units. An example of a data file format can be found in the EMSAMASFF.DOC file.
} } Hi: } } Can anyone fill me in on the details of the EMSA file format for EDS spectra. } } Mostly I need to know how to help someone strip out the header so they can } look at it in Excel. } } The file I see when I look at it has the header info, followed by what } looks like 4 columns of numbers. I was expecting 2 columns, x and y. What's } up? } } Thanks } } Jonathan Krupp } Microscopy & Imaging Lab } University of California } Santa Cruz, CA 95064 } (831) 459-2477 } jmkrupp-at-cats.ucsc.edu
From MicroscopyL-request-at-ns.microscopy.com Sun Sep 26 10:59:11 2004
There is a new technology coming from a company called AETOS which is probably just what you need. Contact Sam Lawrence at 334-749-0134. The technology is called "CytoViva". I had a chance to run it last week and was imaging particles at the 100nm level (I think a combo of both detection and resolution).
Hope this is helpful.
Best regards, Barbara Foster Microscopy/Microscopy Education
We've Moved! 313 S Jupiter Rd, Suite 100 Allen, TX 75002 P: 972-954-8011 F: 972-954-8018
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At 10:48 PM 9/24/2004, timandcathy-at-mho.com wrote:
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From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 06:42:23 2004
Hi Todd, I had a similar problems while cutting semi-thin sections. What I've noticed it always happend after rough triming. I talked to Leica representative and he expalained to me that the problem was how I've been trimming blocks. I advanced knife towards the block and always at the same time I moved the block in the specimen arm up and down, but I was not making a full rotation with the wheel. He told me that even though I do not rotate the specimen arm and it is not advancing it builds up the distance, meaning that every time I moved block up and down it advanced the specimen setting without advancing it physically.( I am not sure if I explained well.) Then when I switch to an automatic mode all this "build up" adds up and microtome jumps the specimen forward cutting a big chunk. I changed my method of trimming and I do not have any problems anymore. I hope my explanation will be of help to you. Dorota
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 07:51:46 2004
Greetings, We are submitting a purchase requisition for an image capture system for our SEM that has an older Microspec WDS spectrometer (pre-Oxford). Due to budget requirements we are unable to upgrade the WDS system and the need to capture the data stream from the spectrometer and bring it into our image capture database is great. Are there any suggestions to accomplish this capture of the data stream from the spectrometer and bring it into the computer for further processing?
Thanks for any help. Robb
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 08:55:45 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kssim-at-mmu.edu.my) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 27, 2004 at 08:30:04 ---------------------------------------------------------------------------
Can we do real-time signal-to-noise ratio value of SEM image ? What is the advantages after obtaining SNR value? If you have the solutions, please kindly email me.
I just ran across this reference in our local newspaper about the care and storage of CDs and DVDs. It is copublished by CLIR (Council on Library and Information Resources) and NIST, so it should be reliable info.
From my quick run through of the article, it appears that DVDs may be inherently more stable than CDs.
Cheers, Henk
Hendrik O. Colijn colijn.1-at-osu.edu Campus Electron Optics Facility Ohio State University (614) 292-0674 http://www.ceof.ohio-state.edu Time is that quality of nature which keeps events from happening all at once. Lately it doesn't seem to be working.
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 10:34:19 2004
Hi all, Despite pending budget cuts, I've been asked to submit an equipment wish list. Go figure;-) So wishfully thinking, I'd really like to hear your thoughts on the various microwave ovens that are available from the EM supply companies. Any great machines out there? Giving great results! thanks in advance, Beth
********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
"Between the two evils, I always pick the one I never tried before". Mae West (1893-1980) *******************************************************************
"And it's only the giving that makes you what you are". Wond'ring Aloud, Jethro Tull (Aqualung)
I am in a similar situation... SEM: Hitachi S-3500H EDS: IXRF Systems (Full) WDS: Microspec 2A w/ Kevex Sesame24 Controller
My WDS controller has an analog ratemeter output for line scans and a "digital" output for mapping. These signals are input to the "x-ray_1 and x-ray_2 external inputs on the SEM. Maps and line scans are captured and stored directly using Hitachi/Quartz software. FWIW: Image brightness/contrast manipulation may be required if your SEM works like my Hitachi. In the mapping mode, I must use slow recursive imaging to have sufficient integration time (other modes too fast) for most work. The 3500 software does not handle this gracefully. The finished map is usually totally black because each "dot" luminosity is divided (I infer) by the number of (recursive) passes and the data summed for presentation. Since dots may not occur at the same exact location for each pass, they are too dim to see. Thus, contrast/brightness expansion, using Photoshop, IrFan View, or the like, is required to see the result. In the linescan mode dynamic range is about half of what would be expected. The software limits graphical amplitude excursions of half (or less) of the vertical range of the crt image.
I often run "spectrum - scans" also, generating data (spectrum graphic) within a limited wavelength range. My Sesame system has a serial data port which will allow me to upload the wavelength positions and x-ray intensities to my EDS PC. Any serial terminal program should work. The saved data can be sent to a spreadsheet, or wherever.
If you have a 3PC? instead of the Kevex Sesame, I cannot be so specific...
Hope this is helpful.
Woody
-----Original Message----- } From: Robb Westby [mailto:robbw-at-mxim.com] Sent: Monday, September 27, 2004 8:51 AM To: microscopy-at-msa.microscopy.com
Greetings, We are submitting a purchase requisition for an image capture system for our SEM that has an older Microspec WDS spectrometer (pre-Oxford). Due to budget requirements we are unable to upgrade the WDS system and the need to capture the data stream from the spectrometer and bring it into our image capture database is great. Are there any suggestions to accomplish this capture of the data stream from the spectrometer and bring it into the computer for further processing?
Thanks for any help. Robb
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 11:06:06 2004
I think there was an article in some magazine months ago on how reliable CD-R are for archival. Depending on the brand (or no-brand if you prefer) it can fail after less than 2 years. That was much less than the 20-100 years promised by the maker. People were recomending to use CD-RW for archival purpose instead. Since it doesn't use a dye for recording the data should be less affected by the problem. I suppose the same applies for DVD+/-R and DVD+/-RW. Regards,
Carlos
On Mon, 27 Sep 2004, Hendrik O. Colijn wrote:
} } Hi all, } } I just ran across this reference in our local newspaper about the care and } storage of CDs and DVDs. It is copublished by CLIR (Council on Library and } Information Resources) and NIST, so it should be reliable info. } } http://www.clir.org/pubs/reports/pub121/contents.html } } From my quick run through of the article, it appears that DVDs may be } inherently more stable than CDs. } } Cheers, } Henk } }
o-------------------------------------------------------o | Carlos Kazuo Inoki | | Department of Physics - University at Albany | | 1400 Washington Ave.- Albany - NY - 12222 | o-------------------------------------------------------o
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 12:26:47 2004
That's an extremely informative and useful article. Thank you!
It was interesting to learn that CD RW and DVD RW (bits encoded in metal structure by a phase change from crystalline to amorphous) are less stable over time than CD R and DVD R (bits encoded in degradable organic dyes). And that RW discs that are "cycled" a lot (many re-writes) will degrade faster.
The best advice seems to be to use R (write once) discs (CD or DVD) and store them vertically in a cool, low-light environment. Of course, transferring to fresh media every year or so couldn't hurt...
Mike
"Hendrik O. Colijn" wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Hi all, } } I just ran across this reference in our local newspaper about the care and } storage of CDs and DVDs. It is copublished by CLIR (Council on Library and } Information Resources) and NIST, so it should be reliable info. } } http://www.clir.org/pubs/reports/pub121/contents.html } } From my quick run through of the article, it appears that DVDs may be } inherently more stable than CDs. } } Cheers, } Henk } } Hendrik O. Colijn colijn.1-at-osu.edu } Campus Electron Optics Facility Ohio State University } (614) 292-0674 http://www.ceof.ohio-state.edu } Time is that quality of nature which keeps events from happening all at } once. Lately it doesn't seem to be working.
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 12:54:34 2004
} Name: John H. Cross, CIH } Organization: Lockheed Martin Space Operations } Title-Subject: [Microscopy] [Filtered] MListserver: Occupational } Health Aspects of } Microscope Usage } Question: We have a group of technicians using microscopes to solder } electronic } components 10-12 hours per day 6-7 days a week. } } Does any professional, academic, or industry organization publish } occupational health } guidelines for the use of microscopes? A sample of questions I need } to address are: } How often must an individual take a break from looking through a microscope. } Is exposure to light coming through the eyepieces a problem? } Has anyone described ergonomically-correct work stations for microscopists? } } All comments will be welcome. I particularly need objective } standards that I can present } to management to justify work practice adjustments. } } John Cross Email: John H. Cross-at-lmco.com ============== John, I am not aware of the reference that you requested. However now that I am in my mid-50's , I recently have gotten several work related physical problems.
When I am using any scope for an extended period of time my eyes fail to focus properly for distance. This I first noticed after nearly a whole day on my TEM, doing a lot of scanning for a particular cell type in sections. When I left the building for the night I could not see properly anything that was more than a few yards in front of me. I realized that this had been happening over an extended period of time but it was not so disturbing as it had become. I consulted my eye doctor and he said that this was not abnormal for people my age. The eye muscles are not as elastic as they were a few years ago hence they do not snap back like they used to. I need to stop "scoping" every 15 minutes or so and give my muscles a workout by focusing on a particular object in the distance for a short while. This has been working great for me after I got used to the short interruptions and my concentration was improved because of the short distraction from what can be very boring, when what I am looking for is difficult to find.
A second problem is the nerve that goes through my elbow. Everyone has heard of carpel tunnel syndrome. There is a similar area in the elbow. The test for this is to extend the arms straight out to the front, at shoulder height with the backs of the hands facing upwards. Make a fist, flex the wrist to raise just the fist upwards. If there is no pain, it is OK, but if you are truly having a problem the pain can be quite intense. I would suggest that one contact the doctor about this. My case was already very painful when I went to the doctor, thinking that it would get better itself - mistake! I received several shots into the area over an 8 week period because I can not take oral anti-inflammatory drugs. I also had to do exercises several times a day to strengthen the surrounding muscles. In the mean time I took several older mouse-pads, cut them in half, stacked them so that my lower arm/elbow is at least as high as my hand when using the controls. When using the hands to solder, one may just need more cushioning under the elbow. I have seen advertised scopes that claim to be more ergonomical but at the moment I can not recall the make.
If this is not already being done, I should expect that the fumes from the solder should be sucked away from the scope and go through a suitable filter so that they are not inhaled by the tech.
Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 13:16:21 2004
Dear John, I am the Ergonomics Representative for our department at UBC and the guidelines I was given for people working at computer terminals all day was: a one minute break every ten minutes and a ten-minute break every hour. The short break was just to ease the eye strain of looking at a fixed focus for extended periods and just consists of focussing out the window or at a longer distance and relaxing arms, hands, neck, etc. The longer hour break was to stand up, stretch, move around and relax. These were just a guideline. There are also things about the setup to check, such as the angle of arms, wrists, hips, knees, the height of chair and height of monitor. Your case is special, but you should be able to work out, with the people doing the work, a checklist of things to adjust to make the work ergonomically sound or as least-damaging as possible. Good support chairs with lots of adjustment, microscopes that can adjust eyepiece spacing, armrests are just some of the things that might help. The light coming from the microscope should not be a problem, but should be controlled by the user so it can be adjusted to a comfortable level. The breaks are important to relieve eye-strain. Good luck, Mary Mager Electron Microscopist Department of 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-interchange.ubc.ca ----- Original Message ----- } From: "by way of MicroscopyListserver" {John.H.Cross-at-lmco.com} To: {microscopy-at-microscopy.com} Sent: Friday, September 24, 2004 6:08 AM
Pat;
Have you tried contacting OSHA [Occupational Safety and Health Aministration]? OSHA is a US Gov't agency. http://www.osha.gov
Peter Tomic Agere Systems Allentown, PA
-----Original Message----- } From: Pat Connelly [mailto:psconnel-at-sas.upenn.edu] Sent: Monday, September 27, 2004 1:54 PM To: by way of MicroscopyListserver Cc: microscopy-at-msa.microscopy.com
} Name: John H. Cross, CIH } Organization: Lockheed Martin Space Operations } Title-Subject: [Microscopy] [Filtered] MListserver: Occupational } Health Aspects of } Microscope Usage } Question: We have a group of technicians using microscopes to solder } electronic } components 10-12 hours per day 6-7 days a week. } } Does any professional, academic, or industry organization publish } occupational health guidelines for the use of microscopes? A sample of
} questions I need to address are: } How often must an individual take a break from looking through a microscope. } Is exposure to light coming through the eyepieces a problem? } Has anyone described ergonomically-correct work stations for microscopists? } } All comments will be welcome. I particularly need objective standards } that I can present to management to justify work practice adjustments. } } John Cross Email: John H. Cross-at-lmco.com ============== John, I am not aware of the reference that you requested. However now that I am in my mid-50's , I recently have gotten several work related physical problems.
When I am using any scope for an extended period of time my eyes fail to focus properly for distance. This I first noticed after nearly a whole day on my TEM, doing a lot of scanning for a particular cell type in sections. When I left the building for the night I could not see properly anything that was more than a few yards in front of me. I realized that this had been happening over an extended period of time but it was not so disturbing as it had become. I consulted my eye doctor and he said that this was not abnormal for people my age. The eye muscles are not as elastic as they were a few years ago hence they do not snap back like they used to. I need to stop "scoping" every 15 minutes or so and give my muscles a workout by focusing on a particular object in the distance for a short while. This has been working great for me after I got used to the short interruptions and my concentration was improved because of the short distraction from what can be very boring, when what I am looking for is difficult to find.
A second problem is the nerve that goes through my elbow. Everyone has heard of carpel tunnel syndrome. There is a similar area in the elbow. The test for this is to extend the arms straight out to the front, at shoulder height with the backs of the hands facing upwards. Make a fist, flex the wrist to raise just the fist upwards. If there is no pain, it is OK, but if you are truly having a problem the pain can be quite intense. I would suggest that one contact the doctor about this. My case was already very painful when I went to the doctor, thinking that it would get better itself - mistake! I received several shots into the area over an 8 week period because I can not take oral anti-inflammatory drugs. I also had to do exercises several times a day to strengthen the surrounding muscles. In the mean time I took several older mouse-pads, cut them in half, stacked them so that my lower arm/elbow is at least as high as my hand when using the controls. When using the hands to solder, one may just need more cushioning under the elbow. I have seen advertised scopes that claim to be more ergonomical but at the moment I can not recall the make.
If this is not already being done, I should expect that the fumes from the solder should be sucked away from the scope and go through a suitable filter so that they are not inhaled by the tech.
Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 15:38:57 2004
JOHN WROTE: ------------------------------------------------------------------------------
The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} Name: John H. Cross, CIH } Organization: Lockheed Martin Space Operations } Title-Subject: [Microscopy] [Filtered] MListserver: Occupational } Health Aspects of } Microscope Usage } Question: We have a group of technicians using microscopes to solder } electronic } components 10-12 hours per day 6-7 days a week. } } Does any professional, academic, or industry organization publish } occupational health } guidelines for the use of microscopes? A sample of questions I need } to address are: } How often must an individual take a break from looking through a microscope. } Is exposure to light coming through the eyepieces a problem? } Has anyone described ergonomically-correct work stations for microscopists? } } All comments will be welcome. I particularly need objective } standards that I can present } to management to justify work practice adjustments. } } John Cross Email: John H. Cross-at-lmco.com
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 19:50:02 2004
Can someone tell me if such a data base exist? I am hoping to get information on rates and annual budget of university EM facilities in the US. Specifically, I am hoping to know the percentage of charge based income vs. university support. If there is not such a data base, would you EM facility directors mind emailing me and giving me some idea about the case in your facility. Thank you all very much in advance.
Hong Emory EM.
From MicroscopyL-request-at-ns.microscopy.com Mon Sep 27 20:56:57 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (walter.bobrowski-at-pfizer.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 27, 2004 at 15:31:59 ---------------------------------------------------------------------------
Email: walter.bobrowski-at-pfizer.com Name: Walt Bobrowski
Question: A colleague is asking if anyone has permanently removed buffered osmium tetroxide out of the routine processing of mammalian tissues for EM, and substituted with another, equally good reagent? I thought it was still the standard post-fix, but maybe I'm behind the times! TIA.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mdawes-at-scu.edu.au) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, September 27, 2004 at 19:33:49 ---------------------------------------------------------------------------
Question: We have a Reflected light fluorescence attachment CX-RFL which fits onto our Olympus Compound microscope CX 40. The Cube Dichroic Mirror/Filter combinations are: a). CX-DMB (Mirror cube), Exciter filter BP475, Barrier filter 05151F and b). CX-DMG (Mirror cube), Exciter filter BP545, Barrier filter 0590. My question is: We are trying to stain some pollen tubes to photograph the pollen tube growth - all papers that I have read seem to use the fluorescing stain Analine Blue which stains the cellulose ñ we do not have a Mirror cube to view this stain and after doing some research with our suppliers we cannot use Aniline blue with the set up we have. We can use the fluorochrome FITC . As there seem to be several stains Fluoresceine diacetate FDA (which appears to stain for protein) and Fluoresceine isothiocyanates FITC (which also appears to stain for protein), my question is can we use either of these stains with our system to photograph pollen tube growth and which one is the easier of the two and is there a tried and tested procedure that someone might be willing to share with us.
Does anyone have an experience or some information about how to realize a sharp cutting edge by hand working an iron piece? I’d like to use it for cutting wax in an hand microtome. I’d like to know the cutting angle, the abrasive powders to use and the operating steps. Thank you. Kindly Regards,
Massimo
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 08:19:14 2004
Maxine, It was probably a typo in your post, but analine blue stains callose (as well as some other polysaccharides), not cellulose. You probably also know that Biosupplies in Mebourne sells a cleaned up version of the dye called Sirofluor that is more speciific for callose, and often gives nicer images, but is still uv excited.
Have you tried imaging analine blue stained stuff with your blue cube? It just might work--I think that analine blue has a pretty broad excitation spectrum.
HTH, Tobias
} } } Email: mdawes-at-scu.edu.au } Name: Maxine Dawes } } Organization: Southern Cross University } } Title-Subject: [Microscopy] [Filtered] Fluorochrome stain - Light Microscope } } Question: We have a Reflected light fluorescence attachment CX-RFL } which fits onto our Olympus Compound microscope CX 40. The Cube } Dichroic Mirror/Filter combinations are: a). CX-DMB (Mirror cube), } Exciter filter BP475, Barrier filter 05151F and b). CX-DMG (Mirror } cube), Exciter filter BP545, Barrier filter 0590. My question is: } We are trying to stain some pollen tubes to photograph the pollen } tube growth - all papers that I have read seem to use the } fluorescing stain Analine Blue which stains the cellulose ñ we do } not have a Mirror cube to view this stain and after doing some } research with our suppliers we cannot use Aniline blue with the set } up we have. We can use the fluorochrome FITC . As there seem to be } several stains Fluoresceine diacetate FDA (which appears to stain } for protein) and Fluoresceine isothiocyanates FITC (which also } appears to stain for protein), my question is can we use either of } these stains with our system to photograph pollen tube growth and } which one is the easier of the two and is there a tried and tested } procedure that someone might be willing to share with us. } } Regards } Maxine } } } ---------------------------------------------------------------------------
Your service engineer should monitor radiation leaking during operation. After 23yrs our SEM has just started to leak (at 30kV only). I am having a lead shield fitted to block the leak.
Dave
On Tue, 28 Sep 2004 11:21:35 +0300 (EEST) Kart Padari {kartp-at-ut.ee} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } } I´m worried about my health too. } My question: Is there any harmful radiation working with } JEOL 100-S electron microscope? } } Kärt Padari } University of Tartu } kartp-at-ut.ee } } } } } } } This incoming email to UWE has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
This email has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 08:45:09 2004
Hong, You can see the rates for my 2 facilities (EM and Optical Microscopy) on the school's website at: http://www.med.cornell.edu/research/cores/index.html
We are subsidized by the College to help cover staff salaries, we are not charged "overhead" costs. Otherwise, the 2 facilities that I'm involved with cover all other costs.
Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 09:10:54 2004
Kärt Padari wrote: ===================================================== I´m worried about my health too. My question: Is there any harmful radiation working with JEOL 100-S electron microscope? ===================================================== My first exposure to an EM was in 1963, the round screen "horizontal" Philips 100 TEM. I was told (but don't know from my own knowledge that this was true) that some of these earliest models were manufactured with ordinary instead of lead glass. I also had contact with an early RCA TEM, I think it was an EMU-1 (or something close to that). Those are the only two EMs I have ever heard about that had inherent radiation problems and once the problem was recognized, retrofits were installed, taking away the problems. I have met operators of those early RCA instruments who literally did their work wearing a lead apron. My sense if that there are none of these old generation EMs in use any more.
Instruments manufactured after that era were designed such that the mass of the column itself would be sufficiently massive and absorbing of x-rays, that raditation just could not escape during operation. True, one could envision that a column could be sufficiently misaligned that x-rays could possibly get out, but the instruments seem to be designed in a way that if they really were misaligned by that extent, there would be a vacuum leak and the high voltage would automatically shut down. Some very many years ago, we tried this with a JEOL 100CX and could not get the microscope sufficiently misaligned to allow x-rays to get out. With a JSM-U3 SEM, at that same time, the column would lose vacuum before it could be misaligned sufficiently to allow x-rays to come through.
But there are at least several circumstances in which radiation can escape:
a) Someone breaks the lead glass viewing screen of a TEM and thinking they are saving money, replaces the broken glass with ordinary glass. I have heard of this very thing happening, on occasion over the years. Such a replacement of the glass would be picked up instantly during any kind of a radiation survey, however. Without such a survey, it is not possible to differentiate, by appearance, lead vs. non-lead glass.
b) Adjustment knobs for the lenses seem to be slightly (x-ray) "leaky" in some instruments, but the exposure is localized to the area of the knobs, and therefore also to the fingers. With modern instruments where this is done by the computer, this risk is much less and might no longer be present. I have heard of people wrapping lead foil around their several fingers when adjusting the knobs but I don't know if that is necessarily a "recommended" procedure. But a general room survey would not necessarily reveal this exposure risk.
c) Home made or radically altered commercial instruments which do not have the "safety engineering" built into them that would otherwise be found in a commerical instrument. I have always thought that this type of instrument should be surveyed quite frequently because of the frequency that changes seem to be made to the basic instrument.
Much of the motivation for those who do radiation surveys involving commercially purchase instruments is to protect the organization from lawsuits from former or present employees who make the claim that some medical condition was either caused or aggrivated by their operation of their TEM or SEM. Considering all the persons who have used both SEMs and TEMs over the years, if there was some health issue, I would have thought that it would have shown up clinically by now. When considering all the risks in an EM loboratory, I have always thought that exposure to the embedding resin chemicals would represent a far greater health risk than operating a column instrument.
This is a very important topic and if there are other perspectives on this, I would surely like to hear them.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
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From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 10:15:45 2004
I would hope that these rates are not for the general public and are restricted to in-house use. Those of us in the private commercial labs are not subsidized by our own tax dollars. This is always an issue for us with faculty taking on outside work and using school resources.
My $0.02 as a local professor was soliciting his services only yesterday's. My associate was a upset with the lack of awareness and sensitivity to the unfair tax advantage.
Alan Stone ASTON
At 08:45 AM 9/28/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Alan Stone ASTON Metallurgical Services Co., Inc. 200 Larkin Drive Ste A Wheeling, IL 60090 847/353-8100 www.astonmet.com
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 10:43:01 2004
Dave, Could you please elaborate? I'm having a little trouble imagining what could change (other than physical modifications) that would allow leakage.
Is the newest survey perhaps simply more sensitive than previous surveys?
What brand and model?
Where has it started leaking?
What kind of intensity?
I service a lot of older scopes (as much as 10 years older than yours) and need to know if this should be more closely watched.
Thanks for your time and help.
Ken Converse
owner QUALITY IMAGES Servicing Scanning Electron Microscopes Since 1981 16 Creek Rd. Delta, PA 17314 717-456-5491 Fax 717-456-7996 kenconverse-at-qualityimages.biz qualityimages.biz
-----Original Message----- } From: Patton, David [mailto:David.Patton-at-uwe.ac.uk] Sent: Tuesday, September 28, 2004 9:20 AM To: Kart Padari Cc: microscopy-at-msa.microscopy.com
Your service engineer should monitor radiation leaking during operation. After 23yrs our SEM has just started to leak (at 30kV only). I am having a lead shield fitted to block the leak.
Dave
On Tue, 28 Sep 2004 11:21:35 +0300 (EEST) Kart Padari {kartp-at-ut.ee} wrote:
} } } ---------------------------------------------------------------------------- -- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- --- } } } } I´m worried about my health too. } My question: Is there any harmful radiation working with } JEOL 100-S electron microscope? } } Kärt Padari } University of Tartu } kartp-at-ut.ee } } } } } } } This incoming email to UWE has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
This email has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
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From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 10:54:28 2004
Dave, Could you please elaborate? I'm having a little trouble imagining what could change (other than physical modifications) that would allow leakage.
Is the newest survey perhaps simply more sensitive than previous surveys?
What brand and model?
Where has it started leaking?
What kind of intensity?
I service a lot of older scopes (as much as 10 years older than yours) and need to know if this should be more closely watched.
Thanks for your time and help.
Ken Converse
owner QUALITY IMAGES Servicing Scanning Electron Microscopes Since 1981 16 Creek Rd. Delta, PA 17314 717-456-5491 Fax 717-456-7996 kenconverse-at-qualityimages.biz qualityimages.biz
-----Original Message----- } From: Patton, David [mailto:David.Patton-at-uwe.ac.uk] Sent: Tuesday, September 28, 2004 9:20 AM To: Kart Padari Cc: microscopy-at-msa.microscopy.com
Your service engineer should monitor radiation leaking during operation. After 23yrs our SEM has just started to leak (at 30kV only). I am having a lead shield fitted to block the leak.
Dave
On Tue, 28 Sep 2004 11:21:35 +0300 (EEST) Kart Padari {kartp-at-ut.ee} wrote:
} } } ---------------------------------------------------------------------------- -- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- --- } } } } I´m worried about my health too. } My question: Is there any harmful radiation working with } JEOL 100-S electron microscope? } } Kärt Padari } University of Tartu } kartp-at-ut.ee } } } } } } } This incoming email to UWE has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
This email has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
___________________________________________________________ $0 Web Hosting with up to 120MB web space, 1000 MB Transfer 10 Personalized POP and Web E-mail Accounts, and much more. Signup at www.doteasy.com
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 12:36:09 2004
Alan, and others My rates are for grant-funded "in house" personnel...we add a whopping surcharge to outside users to compensate for the fact that they don't contribute to the overhead costs of the College as do our grant-funded people. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 13:00:53 2004
At 12:37 PM 9/28/2004, you wrote: } Alan, and others } My rates are for grant-funded "in house" personnel...we add a whopping } surcharge to outside users to compensate for the fact that they don't } contribute to the overhead costs of the College as do our grant-funded people. } Lee } -- } Leona Cohen-Gould, M.S. } Sr. Staff Associate } Director, Electron Microscopy Core Facility } Manager, Optical Microscopy Core Facility } Joan & Sanford I. Weill Medical College } of Cornell University } voice (212)746-6146 } fax (212)746-8175
Alan Stone ASTON Metallurgical Services Co., Inc. 200 Larkin Drive Ste A Wheeling, IL 60090 847/353-8100 www.astonmet.com
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 13:58:37 2004
} IŽm worried about my health too. } My question: Is there any harmful radiation working with } JEOL 100-S electron microscope? } Dear Kärt, There should not be any radiation problems; however, not all EMs have been properly maintained, so it is best to check. The best ways are to use a hand-held ion chamber detector to survey the area while the scope is operated in the most extreme conditions, such as putting a Pt aperture in the stage and hitting the aperture with the most intense beam you can produce, moving the shift and tilt controls through the full extent of their range, etc. Another way is to place film badges or thermoluminescent detectors around the room and measure the background on a month-by-month schedule. You should decide where a radiation leak is most likely by looking at the location of shielding pieces and thin spots in the column wall, and you need to measure the areas where a radiation leak, however unlikely, would do the most damage--obviously where the operator sits, where specimens are loaded, or anywhere that someone is expected to spend a fair amount of time. Remember that x-rays can penetrate walls and floors, so check for radiation that could harm someone in the lab above, below, across the hall, or other possible locations. Thick concrete is a much better shielding material than wood or sheet metal, so more care is required, if your scope is surrounded by the latter. Take the time to make this check, then you will be able to operate with greater peace of mind. Good luck. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 16:43:57 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (donaldawbrey-at-texashealth.org) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, September 28, 2004 at 09:22:18 ---------------------------------------------------------------------------
Email: donaldawbrey-at-texashealth.org Name: Donald G. Awbrey HT(ASCP) QIHC
Organization: Harris Methodist Hospital
Title-Subject: [Microscopy] [Filtered] TEM Laser Printer
Question: Dear Micro netters,
We are looking for a high resolution printer for our TEM workstation. I've seen monochrome lazer printers that have a 1200 X 1200 dpi resolution. Our digital camera will be a 2K camera. Will this printer print photos of diagnostic quality? Or would we have to go with a dye sublimation printer? I've noticed that these sub printers are very expensive, but have a "spatial" resolution of around 320 dpi. Is this the same as regular resolution? If so, then the laser printer would have better resolution. This does not make sense. Is 1200 X 1200 dpi good enough for CAP inspection.
Any information on diagnostic quality printers for TEM will be greatfully appreciated.
Chuck; Chuck; When I was in grad school, we purchased a used JSM-U3 through JEOL that had been in previous use at the GM tech center. The stage door was EXTREMELY leaky under normal use conditions (it was different than our other JSM-U3). I eventually installed a lead lining in the door.
John Mardinly Intel
-----Original Message----- } From: Garber, Charles A. [mailto:cgarber-at-2spi.com] Sent: Tuesday, September 28, 2004 8:12 AM To: MICROSCOPY BB
-- [ From: Garber, Charles A. * EMC.Ver #3.1 ] --
Kärt Padari wrote: ===================================================== I´m worried about my health too. My question: Is there any harmful radiation working with JEOL 100-S electron microscope? ===================================================== My first exposure to an EM was in 1963, the round screen "horizontal" Philips 100 TEM. I was told (but don't know from my own knowledge that this was true) that some of these earliest models were manufactured with ordinary instead of lead glass. I also had contact with an early RCA TEM, I think it was an EMU-1 (or something close to that). Those are the only two EMs I have ever heard about that had inherent radiation problems and once the problem was recognized, retrofits were installed, taking away the problems. I have met operators of those early RCA instruments who literally did their work wearing a lead apron. My sense if that there are none of these old generation EMs in use any more.
Instruments manufactured after that era were designed such that the mass of the column itself would be sufficiently massive and absorbing of x-rays, that raditation just could not escape during operation. True, one could envision that a column could be sufficiently misaligned that x-rays could possibly get out, but the instruments seem to be designed in a way that if they really were misaligned by that extent, there would be a vacuum leak and the high voltage would automatically shut down. Some very many years ago, we tried this with a JEOL 100CX and could not get the microscope sufficiently misaligned to allow x-rays to get out. With a JSM-U3 SEM, at that same time, the column would lose vacuum before it could be misaligned sufficiently to allow x-rays to come through.
But there are at least several circumstances in which radiation can escape:
a) Someone breaks the lead glass viewing screen of a TEM and thinking they are saving money, replaces the broken glass with ordinary glass. I have heard of this very thing happening, on occasion over the years. Such a replacement of the glass would be picked up instantly during any kind of a radiation survey, however. Without such a survey, it is not possible to differentiate, by appearance, lead vs. non-lead glass.
b) Adjustment knobs for the lenses seem to be slightly (x-ray) "leaky" in some instruments, but the exposure is localized to the area of the knobs, and therefore also to the fingers. With modern instruments where this is done by the computer, this risk is much less and might no longer be present. I have heard of people wrapping lead foil around their several fingers when adjusting the knobs but I don't know if that is necessarily a "recommended" procedure. But a general room survey would not necessarily reveal this exposure risk.
c) Home made or radically altered commercial instruments which do not have the "safety engineering" built into them that would otherwise be found in a commerical instrument. I have always thought that this type of instrument should be surveyed quite frequently because of the frequency that changes seem to be made to the basic instrument.
Much of the motivation for those who do radiation surveys involving commercially purchase instruments is to protect the organization from lawsuits from former or present employees who make the claim that some medical condition was either caused or aggrivated by their operation of their TEM or SEM. Considering all the persons who have used both SEMs and TEMs over the years, if there was some health issue, I would have thought that it would have shown up clinically by now. When considering all the risks in an EM loboratory, I have always thought that exposure to the embedding resin chemicals would represent a far greater health risk than operating a column instrument.
This is a very important topic and if there are other perspectives on this, I would surely like to hear them.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 17:43:58 2004
Peter; I'm not sure what good OSHA is. They require records of film badges to be kept for 30 years but don't require film badges, so guess what my employer did? They elimininated film badges!
John Mardinly Intel
-----Original Message----- } From: Tomic, Peter (Peter) [mailto:ptomic-at-agere.com] Sent: Monday, September 27, 2004 12:06 PM To: Pat Connelly; by way of MicroscopyListserver Cc: microscopy-at-msa.microscopy.com
Pat;
Have you tried contacting OSHA [Occupational Safety and Health Aministration]? OSHA is a US Gov't agency. http://www.osha.gov
Peter Tomic Agere Systems Allentown, PA
-----Original Message----- } From: Pat Connelly [mailto:psconnel-at-sas.upenn.edu] Sent: Monday, September 27, 2004 1:54 PM To: by way of MicroscopyListserver Cc: microscopy-at-msa.microscopy.com
} Name: John H. Cross, CIH } Organization: Lockheed Martin Space Operations } Title-Subject: [Microscopy] [Filtered] MListserver: Occupational } Health Aspects of } Microscope Usage } Question: We have a group of technicians using microscopes to solder } electronic } components 10-12 hours per day 6-7 days a week. } } Does any professional, academic, or industry organization publish } occupational health guidelines for the use of microscopes? A sample of
} questions I need to address are: } How often must an individual take a break from looking through a microscope. } Is exposure to light coming through the eyepieces a problem? } Has anyone described ergonomically-correct work stations for microscopists? } } All comments will be welcome. I particularly need objective standards } that I can present to management to justify work practice adjustments. } } John Cross Email: John H. Cross-at-lmco.com ============== John, I am not aware of the reference that you requested. However now that I am in my mid-50's , I recently have gotten several work related physical problems.
When I am using any scope for an extended period of time my eyes fail to focus properly for distance. This I first noticed after nearly a whole day on my TEM, doing a lot of scanning for a particular cell type in sections. When I left the building for the night I could not see properly anything that was more than a few yards in front of me. I realized that this had been happening over an extended period of time but it was not so disturbing as it had become. I consulted my eye doctor and he said that this was not abnormal for people my age. The eye muscles are not as elastic as they were a few years ago hence they do not snap back like they used to. I need to stop "scoping" every 15 minutes or so and give my muscles a workout by focusing on a particular object in the distance for a short while. This has been working great for me after I got used to the short interruptions and my concentration was improved because of the short distraction from what can be very boring, when what I am looking for is difficult to find.
A second problem is the nerve that goes through my elbow. Everyone has heard of carpel tunnel syndrome. There is a similar area in the elbow. The test for this is to extend the arms straight out to the front, at shoulder height with the backs of the hands facing upwards. Make a fist, flex the wrist to raise just the fist upwards. If there is no pain, it is OK, but if you are truly having a problem the pain can be quite intense. I would suggest that one contact the doctor about this. My case was already very painful when I went to the doctor, thinking that it would get better itself - mistake! I received several shots into the area over an 8 week period because I can not take oral anti-inflammatory drugs. I also had to do exercises several times a day to strengthen the surrounding muscles. In the mean time I took several older mouse-pads, cut them in half, stacked them so that my lower arm/elbow is at least as high as my hand when using the controls. When using the hands to solder, one may just need more cushioning under the elbow. I have seen advertised scopes that claim to be more ergonomical but at the moment I can not recall the make.
If this is not already being done, I should expect that the fumes from the solder should be sucked away from the scope and go through a suitable filter so that they are not inhaled by the tech.
Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Sep 28 21:56:08 2004
Chuck's mention of the horizontal Philips 100 TEM brought to mind a story that very likely was true. At the beginning of my microscopy career I worked in the lab of Humberto Fernandez-Moran (of pointed filament and diamond knife fame). Moran started using TEMS in Europe in the late 40's including working on the earliest versions of the Philips 100 TEM. He had a a large scar on his nose from surgery to remove a skin cancer. He blamed the cancer on long hours with his face pressed close to the unleaded window of the TEM and the resultant exposure to x-rays. Being rather sensitive to this problem, he was quite careful to have later instruments tested. When I worked in his lab we had Siemens Elmiskop 1 and 1a TEMs that tested fine.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building 170 S. University Street West Lafayette, IN 47907 http://www3.agriculture.purdue.edu/microscopy
On 9/28/04 5:39 PM, "Mardinly, John" {john.mardinly-at-intel.com} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } Chuck; } Chuck; } When I was in grad school, we purchased a used JSM-U3 through JEOL that had } been in previous use at the GM tech center. The stage door was EXTREMELY leaky } under normal use conditions (it was different than our other JSM-U3). I } eventually installed a lead lining in the door. } } John Mardinly } Intel } } -----Original Message----- } } From: Garber, Charles A. [mailto:cgarber-at-2spi.com] } Sent: Tuesday, September 28, 2004 8:12 AM } To: MICROSCOPY BB } Subject: [Microscopy] Safety around a column instrument } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------} - } } -- [ From: Garber, Charles A. * EMC.Ver #3.1 ] -- } } Kärt Padari wrote: } ===================================================== } I´m worried about my health too. } My question: Is there any harmful radiation working with } JEOL 100-S electron microscope? } ===================================================== } My first exposure to an EM was in 1963, the round screen "horizontal" } Philips 100 TEM. I was told (but don't know from my own knowledge that } this was true) that some of these earliest models were manufactured with } ordinary instead of lead glass. I also had contact with an early RCA TEM, } I think it was an EMU-1 (or something close to that). Those are the only two } EMs I have ever heard about that had inherent radiation problems and once } the problem was recognized, retrofits were installed, taking away the } problems. I have met operators of those early RCA instruments who } literally did their work wearing a lead apron. My sense if that there are } none of these old generation EMs in use any more. } } Instruments manufactured after that era were designed such that the mass of } the column itself would be sufficiently massive and absorbing of x-rays, } that raditation just could not escape during operation. True, one could } envision that a column could be sufficiently misaligned that x-rays could } possibly get out, but the instruments seem to be designed in a way that if } they really were misaligned by that extent, there would be a vacuum leak and } the high voltage would automatically shut down. Some very many years ago, } we tried this with a JEOL 100CX and could not get the microscope } sufficiently misaligned to allow x-rays to get out. With a JSM-U3 SEM, at } that same time, the column would lose vacuum before it could be misaligned } sufficiently to allow x-rays to come through. } } } But there are at least several circumstances in which radiation can escape: } } a) Someone breaks the lead glass viewing screen of a TEM and thinking they } are saving money, replaces the broken glass with ordinary glass. I have } heard of this very thing happening, on occasion over the years. Such a } replacement of the glass would be picked up instantly during any kind of a } radiation survey, however. Without such a survey, it is not possible to } differentiate, by appearance, lead vs. non-lead glass. } } b) Adjustment knobs for the lenses seem to be slightly (x-ray) "leaky" in } some instruments, but the exposure is localized to the area of the knobs, } and therefore also to the fingers. With modern instruments where this is } done by the computer, this risk is much less and might no longer be present. } I have heard of people wrapping lead foil around their several fingers when } adjusting the knobs but I don't know if that is necessarily a "recommended" } procedure. But a general room survey would not necessarily reveal this } exposure risk. } } c) Home made or radically altered commercial instruments which do not have } the "safety engineering" built into them that would otherwise be found in a } commerical instrument. I have always thought that this type of instrument } should be surveyed quite frequently because of the frequency that changes } seem to be made to the basic instrument. } } } Much of the motivation for those who do radiation surveys involving } commercially purchase instruments is to protect the organization from } lawsuits from former or present employees who make the claim that some } medical condition was either caused or aggrivated by their operation of } their TEM or SEM. Considering all the persons who have used both SEMs and } TEMs over the years, if there was some health issue, I would have thought } that it would have shown up clinically by now. When considering all the } risks in an EM loboratory, I have always thought that exposure to the } embedding resin chemicals would represent a far greater health risk than } operating a column instrument. } } This is a very important topic and if there are other perspectives on this, } I would surely like to hear them. } } Chuck } } =================================================== } Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 } President 1-(800)-2424-SPI } SPI SUPPLIES FAX: 1-(610)-436-5755 } PO BOX 656 e-mail: cgarber-at-2spi.com } West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com } } } Look for us! } ############################ } WWW: http://www.2spi.com } ############################ } ================================================== } } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 29 01:00:02 2004
As has been mentioned previously on this listserver, at first nearly every state, if not all of them, required regular surveys of EMs. This was sparked by at least one manufacturer that released some TEMs with non-leaded glass (problems mainly restricted to the facial area) and I believe at least one other instance of TEM camera assemblies that leaked (a number of early TEM operators took to clipping their film badges over their genitals). Over the years, most states have dropped this requirement as few surveys ever detect anything. However, a good knowledge of the potential radiation and the detecting methods used is really required.
Any change in radiation leakage in an EM over time is probably an indication that either something has been improperly changed in the EM or the instruments being used to detect a problem have changed.
In the first case, the basic design of a modern EM (and by that I mean anything produced in the last 30 years) makes them inherently safe in most respects. The standard design of shrouding electromagnetic lenses in soft iron (ferromagnetics) in order to use those shrouds to focus the fields in narrow spaces means that the majority of the column is well shielded. In areas where these lenses aren't used, the outer structures are very thick metal just to support the electron gun or upper lenses and keep them in alignment. Where these sections are mated, they normally have close fitting, concentric overlaps for alignment. In other sealing areas, such as vacuum manifolds, gauges, port covers and door seals, sufficient overlap is provided to prevent a direct path for x-rays.
For example, a flat door with an o-ring sealing up against flat metal chamber in an SEM generally has a half an inch to an inch of overlap. Yes, x-rays will leak, but due to the repeated, zigzag path they would have to take to get out, you won't see primary x-rays. Instead, you will get the result of multiple fluorescence events, with each generation producing x-rays of lower energy and fewer photons that can make it out. There could, however, be structures within the chamber that are always, or occasionally, in the right position to provide a fluorescence to the primary sample x-rays with a portion going directly out between the door and the chamber, through the o-ring.
That brings us to the equipment used to detect these leaks. Geiger tubes of various shapes and sizes have been the standard over the years. There are two general varieties - compensated and uncompensated. The uncompensated tube is a bare tube. Because of the design, the uncompensated Geiger tube is extremely sensitive to x-rays, the lower the energy the more it reports. As long as some x-rays can penetrate the outer glass tube, or cause a secondary fluorescence in it, the lower energy x-rays stand a much better chance of ionizing the gas within.
Because a wide range of radiation detection is desired for most Geiger counters, various means are used to 'compensate' them. This basically means shielding them from lower energy x-rays so that they aren't swamped by them and thus higher energy x-rays and gamma rays can be more effectively detected. Other types of radiation detectors can have a response similar to the uncompensated Geiger tube - they can be far more sensitive to low energy x-rays than other radiation, but they are often calibrated to high energy x-rays or gamma rays (the most interesting form of radiation detection for many uses) and perhaps compensated in some way.
Leaving the technical behind, a change over time in the x-ray emissions of an EM could also be the result of a change in instrumentation to detect it. If the detector doesn't have a linear calibration that includes the low x-ray energies, then it may produce results on an EM that don't represent reality. Since most survey instruments can't differentiate radiation sources by energy, appropriate adjustment has to be made to their readings. An SEM running at 30KV won't leak 30KV x-rays, and if that assumption is made the reading will be off. More than likely any emitted radiation would be an order of magnitude or more lower in energy. At those energies, the survey meter would likely have a much different sensitivity, perhaps several orders of magnitude.
In regards to vacuum integrity, generally an instrument column misaligned enough to allow x-ray leakage from the column would not be capable of maintaining a vacuum and thus the instrument would itself prevent the application of the electron beam. Avoiding the issue of properly operating vacuum interlocks, I confess I can think of an instrument or two that could have such misalignments and still operate. These would be visibly obvious to anyone familiar with the instrument and extremely unlikely to occur even from the most inexperienced service personnel. Yet, the possibility exists.
User modification of an instrument is one I watch for carefully. I have had customers who have done things like replace a sample chamber metal port cover with a Plexiglas panel so they could peer in and see the sample positioning. Of course, they would fashion a light tight cover to prevent swamping of the secondary electron detector in operation, but it would usually be of a thin aluminum construction. Great, we now have an aluminum fluorescence source external to the chamber. Don't try this at home, kiddies.
Localized, point sources such as lens adjustment knobs present little problem. First, remember that radiation rates fall off as the inverse square of the distance. If you measure a dose rate at 5mm from the external exit, at 10mm it is 1/4 as strong, at 20mm 1/16 the strength and at 40mm (just 1.6") 1/64 as strong.
You also have to consider the extent of the exposure. We probably all agree that we'd like to minimize exposure to radiation as much as possible, but what areas of the body are exposed have a large effect on any probable future problems. While not fully understood, radiation effects are generally agreed to affect different parts of the body to differing extents. At the low end of things is exposure of peripheral areas - hands, arms, feet and legs. These areas don't contain the essential organs that the thorax does, don't have the faster regenerating cells or the high blood supply that can support rapid cellular mutations. While there are always exceptions, you rarely hear of cancers that originate in the fingers or toes, arms or legs.
My own personal feeling is that Chuck is right, you stand a much greater chance of long term damage from the various chemicals used in sample preparation, not to mention common solvents and the samples themselves, used in an EM lab than you do from possible radiation from an EM.
Allen R. Sampson Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174
On Tuesday, September 28, 2004 10:12 AM, Garber, Charles A. [SMTP:cgarber-at-2spi.com] wrote: } } } ------------------------------------------------------------------------ ------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------ ------- } } -- [ From: Garber, Charles A. * EMC.Ver #3.1 ] -- } } Kart Padari wrote: } ===================================================== } I?m worried about my health too. } My question: Is there any harmful radiation working with } JEOL 100-S electron microscope? } ===================================================== } My first exposure to an EM was in 1963, the round screen "horizontal" } Philips 100 TEM. I was told (but don't know from my own knowledge that } this was true) that some of these earliest models were manufactured with } ordinary instead of lead glass. I also had contact with an early RCA TEM, } I think it was an EMU-1 (or something close to that). Those are the only two } EMs I have ever heard about that had inherent radiation problems and once } the problem was recognized, retrofits were installed, taking away the } problems. I have met operators of those early RCA instruments who } literally did their work wearing a lead apron. My sense if that there are } none of these old generation EMs in use any more. } } Instruments manufactured after that era were designed such that the mass of } the column itself would be sufficiently massive and absorbing of x-rays, } that raditation just could not escape during operation. True, one could } envision that a column could be sufficiently misaligned that x-rays could } possibly get out, but the instruments seem to be designed in a way that if } they really were misaligned by that extent, there would be a vacuum leak and } the high voltage would automatically shut down. Some very many years ago, } we tried this with a JEOL 100CX and could not get the microscope } sufficiently misaligned to allow x-rays to get out. With a JSM-U3 SEM, at } that same time, the column would lose vacuum before it could be misaligned } sufficiently to allow x-rays to come through. } } } But there are at least several circumstances in which radiation can escape: } } a) Someone breaks the lead glass viewing screen of a TEM and thinking they } are saving money, replaces the broken glass with ordinary glass. I have } heard of this very thing happening, on occasion over the years. Such a } replacement of the glass would be picked up instantly during any kind of a } radiation survey, however. Without such a survey, it is not possible to } differentiate, by appearance, lead vs. non-lead glass. } } b) Adjustment knobs for the lenses seem to be slightly (x-ray) "leaky" in } some instruments, but the exposure is localized to the area of the knobs, } and therefore also to the fingers. With modern instruments where this is } done by the computer, this risk is much less and might no longer be present. } I have heard of people wrapping lead foil around their several fingers when } adjusting the knobs but I don't know if that is necessarily a "recommended" } procedure. But a general room survey would not necessarily reveal this } exposure risk. } } c) Home made or radically altered commercial instruments which do not have } the "safety engineering" built into them that would otherwise be found in a } commerical instrument. I have always thought that this type of instrument } should be surveyed quite frequently because of the frequency that changes } seem to be made to the basic instrument. } } } Much of the motivation for those who do radiation surveys involving } commercially purchase instruments is to protect the organization from } lawsuits from former or present employees who make the claim that some } medical condition was either caused or aggrivated by their operation of } their TEM or SEM. Considering all the persons who have used both SEMs and } TEMs over the years, if there was some health issue, I would have thought } that it would have shown up clinically by now. When considering all the } risks in an EM loboratory, I have always thought that exposure to the } embedding resin chemicals would represent a far greater health risk than } operating a column instrument. } } This is a very important topic and if there are other perspectives on this, } I would surely like to hear them. } } Chuck } } =================================================== } Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 } President 1-(800)-2424-SPI } SPI SUPPLIES FAX: 1-(610)-436-5755 } PO BOX 656 e-mail: cgarber-at-2spi.com } West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com } } } Look for us! } ############################ } WWW: http://www.2spi.com } ############################ } ================================================== } } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 29 11:52:45 2004
Short Course and Workshop Announcement University of Missouri - Columbia (originally scheduled for August 16-18, 2004 was postponed due to Dr. Russ' illness)
The Electron Microscopy Core Facility is hosting a 3-day Short Course and Workshop on Computer-Assisted Image Analysis and Measurement taught by Dr. John C. Russ on November 9=11, 2004. This popular course is intended to familiarize users of image analysis equipment with the fundamental principles and methods available to obtain meaningful results, and to educate laboratory supervisors or research professionals seeking to learn how to use such methods in their applications. The techniques are applicable to fields ranging from materials, geological and biological/medical research to food technology and manufacturing quality control.
The course relies heavily on tightly coupled lectures and hands-on experience with the various techniques. The laboratory includes a wide variety of image analysis methods designed to cover the range of approaches and tools, and a detailed set of practical instructions to enable their use with a minimal learning curve. No specific background is assumed, although users should already be familiar with microscopy or other imaging technology, and the techniques required to obtain the images to be measured. Many of the examples used in the course involve light or electron microscope images, but students are invited to bring their own most interesting images (TIFF files) for discussion and analysis.
Image analysis and measurement methods are used in a broad range of applications and are usually concerned with extracting a few numerical values, such as the number, size, shape or location of objects from the image. In other cases, global structural parameters such as measures of the volume and surface of structures present are of interest. These measurements may require image processing to correct defects, feature enhancement, comparison of multiple images, object recognition, or other steps. Ultimately, the image is reduced to just the features of interest. Measurements on these individual features, or on the image as a whole, must then be obtained and interpreted in a proper stereological context to obtain useful data about the objects. Statistical interpretation of the data allows comparisons of different populations, understanding of distribution plots, and other inferences about the original objects. Structural modeling and geometric probability can be used to develop models for this interpretation.
The course will have an enrollment limit of 20, and there are ~10 spots still open. More information including a registration form can be found at http://www.emc.missouri.edu, or by contacting Lou Ross at 573.882.4777 or rosslm-at-missouri.edu. -- Senior Electron Microscope Specialist Electron Microscopy Core Facility W136 Veterinary Medicine University of Missouri Columbia, MO 65211-5120 (573) 882-4777, fax 884=5414 email: rosslm-at-missouri.edu http://www.emc.missouri.edu
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 29 11:59:53 2004
Can you help my colleague, Milos Kalab, to solve his problem. His question is posted below. Thank you.
Ann Fook Yang EM Unit/ Unite EM Agriculture and Agri-Food Canada/Agriculture et Agroalimentaire Canada Telephone/Téléphone: 613-759-1638 Facsimile/Télécopieur: 613-759-1701 960 Carling Ave/960 Boul Carling Ottawa,Ontario/Ottawa, Ontario Canada K1A 0C6 yanga-at-agr.gc.ca
My objective is to obtain SEM images of stainless steel and other surfaces used in food processing. For this reason I wish to replicate small parts of the surfaces and examine the negative replicas. Advice on materials and procedures which would reproduce details down to 1 micrometer would be appreciated. (I am aware of Ted Pella and SPI cellulose acetate replication materials for coarser surfaces and I wonder why I have not been successful.)
Thank you.
Milos Kalab
Agriculture and Agri-Food Canada in Ottawa
kalabm-at-agr.gc.ca
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 29 12:23:42 2004
Is any one using or know of a consumer camera with full computer control and live video preview mounted on an optical microscope?
Peggy Miller UTHSCSA Ophthalmology Lions Sight Research Center 7703 Floyd Curl Drive MC6230 San Antonio, Texas 78229-3900 PH: (210)567-8460 FAX:(210)567-8413
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 29 16:11:25 2004
We routinely use acetate replicas from machined metal surfaces that resolve to the micron level.
Can you describe the procedure used by Milos? Perhaps it differs from what we do.
Regards, Woody
-----Original Message----- } From: Yang, Ann-Fook [mailto:YANGA-at-agr.gc.ca] Sent: Wednesday, September 29, 2004 1:01 PM To: microscopy-at-microscopy.com
Hi everyone,
Can you help my colleague, Milos Kalab, to solve his problem. His question is posted below. Thank you.
Ann Fook Yang EM Unit/ Unite EM Agriculture and Agri-Food Canada/Agriculture et Agroalimentaire Canada Telephone/Téléphone: 613-759-1638 Facsimile/Télécopieur: 613-759-1701 960 Carling Ave/960 Boul Carling Ottawa,Ontario/Ottawa, Ontario Canada K1A 0C6 yanga-at-agr.gc.ca
My objective is to obtain SEM images of stainless steel and other surfaces used in food processing. For this reason I wish to replicate small parts of the surfaces and examine the negative replicas. Advice on materials and procedures which would reproduce details down to 1 micrometer would be appreciated. (I am aware of Ted Pella and SPI cellulose acetate replication materials for coarser surfaces and I wonder why I have not been successful.)
Thank you.
Milos Kalab
Agriculture and Agri-Food Canada in Ottawa
kalabm-at-agr.gc.ca
From MicroscopyL-request-at-ns.microscopy.com Wed Sep 29 17:10:08 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mark.evans-at-uvm.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, September 29, 2004 at 15:53:40 ---------------------------------------------------------------------------
Email: mark.evans-at-uvm.edu Name: Mark Evans
Organization: University of Vermont
Title-Subject: [Microscopy] [Filtered] MListserver: Isoamyl acetate and critical point drying
Question: Some protocols for sem indicate that following ethanol series dehydration samples should be soaked in isoamyl acetate and that this media should also be used for critical point drying. Can anyone tell me what the theory is behind using isoamyl acetate and what benefit, if any, there is over leaving samples in 100% ethanol prior to the critical point drying? Thanks, Mark
Milos Kalab asks: ==================================================== My objective is to obtain SEM images of stainless steel and other surfaces used in food processing. For this reason I wish to replicate small parts of the surfaces and examine the negative replicas. Advice on materials and procedures which would reproduce details down to 1 micrometer would be appreciated. (I am aware of Ted Pella and SPI cellulose acetate replication materials for coarser surfaces and I wonder why I have not been successful.) ==================================================== If you are talking about pretty smooth surfaces, there is no reason that I can see why either a) the cellulose acetate system should not "work" or b) you could also try what we at SPI Supplies call the SPI Wet Replica Kit, which is a very rapidly curing silicone system.
You did not say what your basis was for it "not working" but I will assume that you just did not "see" anything when in fact you expected to be seeing something. There could be several possible reasons for this:
a) there is some kind of organic contamination coating the surface, so if you solvent washed the surface (or plasma etched it if it was not too large to get into a plasma etcher) you might "uncover" what you wanted to be seeing and/or
b) work with a more dilute solution of the cellulose acetate or
c) there is structure there but you might do better seeing it by Pt/C shadowing the cellulose acetate (negative) replica, dissolving away the plastic, picking up the replica on a TEM grid and viewing the replica in a TEM instead of an SEM. It is possible that the topographical variation of your structure is insufficient to give the level of contrast needed to be seen by SEM.
The silicone system might be easier to work with, especially if you wanted to return to the same identical area, if you wanted to be following the same area as a function of time (e.g. as a function of use if you are doing wear studies). But the same comments about making sure the surface is "clean" and properly "prepared" would apply, otherwise you will just be replicating the surface of the contamination or residues. The silicone system would work only for SEM examination, not TEM.
Disclaimer: SPI Supplies offers both cellulose acetate replicating tapes and sheets and also the SPI Wet Replica kit as described on the website given below.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 02:28:47 2004
I have not used amyl acetate for CPD since my first CPD run almost 30 years ago. The head of department was incensed by the strong pear/banana smell and banned it as antisocial. In those days nobody seemed to worry about its toxicity. Amyl acetate also damages the rubber seals of the CPD faster than other solvents.
The objective is to fully replace the dehydrating solvent with liquid CO2. Amyl acetate is more soluble in liquid CO2 than the more polar ethanol, but many people nevertheless use ethanol routinely and obtain great results, especially if the specimens are small, thin, and unenclosed by cuticle (cultured cells on coverslips, for example). If the specimens are large chunks of plant material it would be better to work with acetone (propanone) as both dehydrating and intermediate solvent.
Dr. Chris Jeffree University of Edinburgh
----- Original Message ----- } From: "by way of MicroscopyListserver" {mark.evans-at-uvm.edu} To: {microscopy-at-microscopy.com} Sent: Wednesday, September 29, 2004 11:12 PM
Peggy
most of the recent Canon Powershot range allow for control and previewing of the camera from a computer via the USB port. The A80 (recently superseded by the A95) and the A95 also have a screen which can be rotated to a useful angle for viewing when the camera is on the top of a microscope. One disadvantage is that they only produce JPEG images (not RAW or TIFF). You could check some of the specifications at: http://consumer.usa.canon.com/ir/controller?act=ProductCatIndexAct&fcategoryid=113
A more upmarket Canon would be the Powershot G6 (this does JPEG and RAW format I believe) - see website below for a review which includes detail of the computer control facilities: http://www.dcresource.com/reviews/canon/powershot_g6-review/index.shtml
I would assume that some Nikons may have the facilities that you want but I have no personal experience of them - just my trusty little A80.
Certainly there are are microscope and camera adaptors to enable attachment to light microscopes for Nikon and Canon digital cameras - just do a web search.
Malcolm
Malcolm Haswell e.m. unit School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear SR1 3SD UK e-mail: malcolm.haswell-at-sunderland.ac.uk
----- Original Message ----- } From: "Miller, Margaret M" {MILLERMM-at-uthscsa.edu}
Hi, Peggy!
We do quite a lot of LM digital imaging these days. Through Nikon we had purchased the Photometrics Cool Snap cf camera to place atop an Olympus BH-2 light microscope. The software program we're using is called Metavue; it allows you to view live and make changes to lighting and such before saving. The best part is being able to make much smaller files by making duplicate images and saving into TIFF. It also allows you to put in micron bars after calibrating your scope and camera system together. Afterwards, the duplicated images are able to be used in Photoshop or Power Point.
Good luck in your search!
*Disclaimer--I do not work for, or receive any compensation from, Nikon or the makers of Metavue. We are just satisfied with the setup.
Donna R. Clarkson, HT (ASCP) Northrop Grumman Information Technology for U S Army Medical Research Detachment at Brooks City-Base Phone (210) 536-1416 FAX (210) 536-1449 e-mail donna.clarkson-at-brooks.af.mil "Our Army at War--Relevant and Ready"
-----Original Message----- } From: Miller, Margaret M [mailto:MILLERMM-at-uthscsa.edu] Sent: Wednesday, September 29, 2004 12:25 PM To: Microscopy-at-microscopy.com
Dear Listserver,
Is any one using or know of a consumer camera with full computer control and live video preview mounted on an optical microscope?
Peggy Miller UTHSCSA Ophthalmology Lions Sight Research Center 7703 Floyd Curl Drive MC6230 San Antonio, Texas 78229-3900 PH: (210)567-8460 FAX:(210)567-8413
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 09:04:36 2004
Amyl acetate is supposed to be more miscible in lqCO2 than is EtOH. I have also heard that it's supposed to be a suspected carcinogen, but I don't have any references on that. One reason I was given for using Amyl acetate when I was learning EM Back When was that it smells of bananas, and therefore it can be easily known when it's all been flushed out. I've never used A. acetate, just EtOH, and I get fine results. I may have to purge more than I would if I used A. acetate, but not much. And the EtOH has plenty of odor, so it's also easy to tell when the bulk is gone by smell. But it's the Aac or EtOH *in* the samples that has to be gotten rid of, not the bulk fluid, and that's too small a volume to smell anyway. Phil
} Email: mark.evans-at-uvm.edu } Name: Mark Evans } } Organization: University of Vermont } } Title-Subject: [Microscopy] [Filtered] MListserver: Isoamyl acetate } and critical point drying } } Question: Some protocols for sem indicate that following ethanol } series dehydration samples should be soaked in isoamyl acetate and } that this media should also be used for critical point drying. Can } anyone tell me what the theory is behind using isoamyl acetate and } what benefit, if any, there is over leaving samples in 100% ethanol } prior to the critical point drying? } Thanks, } Mark } } ---------------------------------------------------------------------------
-- Philip Oshel Supervisor, BBPIC microscopy facility Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 09:07:27 2004
Greetings Margaret, I don't know of any "consumer grade" cameras (the kind you can get at BestBuy) with real time focusing and robust computer control. We had an older Kodak DC210 "micro-imaging package" and it was, and still is, quite possibly the worst ccd solution marketed for microscopy that I have ever seen. The little Pixellink cameras for microscopy are fine for routine brightfield work but they cost about $2k (the Kodak system was about the same amount several years ago). A favorite among amateur microscopists is the Nikon CoolPix (google nikon coolpix microscopy), but I don't know of a real time computer control interface for it. In order to use such a camera on a microscope you either have to dimantle the camera to expose the bare ccd, then make some sort of c-mount adapter, or you will have to get a hold of a special optical coupler with a lens system to project the image on the ccd correctly through the camera lens. Actually I've heard that you can take a snapshot directly through the eyepiece if you hold the camera just right. I highly recommend a camera designed for microscopy if you don't want to look through the eyepieces or don't like judging image quality from a 2 inch low-res lcd. -Karl
-- Karl Garsha Light Microscopy Specialist Imaging Technology Group Beckman Institute for Advanced Science and Technology University of Illinois at Urbana-Champaign 405 North Mathews Avenue Urbana, IL 61801 Office: B650J Phone: 217.244.6292 Fax: 217.244.6219 Mobile: 217.390.1874 www.itg.uiuc.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 10:08:14 2004
The following is the agenda for the Fall MATCSEM and Midwest Society meeting. For those planning on attending, please email me for parking permits:
wcarmichael-at-matcmadison.edu
Annual Fall MATCEM / MMMS Microscopy Meeting
Friday, October 15 at Madison Area Technical College, Truax campus
8:45 - 9:30 Meeting registration (Coffee, tea, juice, donuts and bagels will be provided)
9:30 - 9:45 Welcome, Introductions and Business information
9:45 - 10:30 Food Microstructure: Industry Perspectives By Randy Brandsma of Schreiber Foods, Inc. Randy is currently a research scientist with Schreiber Foods Inc. of Green Bay WI. After receiving his MS in Dairy Science and working as a QA manager for Davisco International he earned his PhD from Cornell University with focus on membrane filtration for cheese manufacturing. He is currently using Confocal and SEM in his work to understand the microstructure of cheese and its effects on performance.
10:30 - 12:00 Forensic Microscopical Analysis of Food, Beverages and Containers from Top to Bottom. By Richard E. Bisbing and Elaine F. Schumacher of McCrone Associates, Inc.
Richard is the Executive Vice President, managing the technical operations group and overseeing all services at McCrone Associates. Prior to joining McCrone Richard was involved in trace evidence work for the Michigan state police and Dept. of public health. He continues to consult on analytical light microscopy and forensic sciences.
Elaine is a Senior Research Scientist and utilizes TEM techniques such as high resolution imaging, electron diffraction, x-ray microanalysis and EELS to characterize a wide variety of industrial materials and problems. Prior to McCrone Elaine worked at UOP Research Center developing catalysts and process technology for the refining industry.
Lunch - A box lunch will be provided courtesy of the MATC vendors
1:15 - 2:30 How Measurements in the nm to µm Size Range Help in Polymer Product Development by Bodan Ma and Sunil Jayasuriya of SC Johnson Polymer
Sunil is the Technical Manager for Polymer Characterization and Applications Research group. He earned his PhD in Physical Chemistry from the Univ. of Bristol, UK. Sunil has worked on applications of surface chemistry and colloid science for printing, personal care, and package coating industries.
Bodan is the Technical Manager for Coatings and has extensive experience in the structure-property relationship of polymeric materials, such as adhesives and coatings. He holds a PhD in Polymer Sciences from Univ. of Massachusetts, Amherst and holds 8 US patents in formulation of new materials.
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 11:10:47 2004
The very old Sorvall Critical Point Dryer came with a spare parts box that had special O-Rings that needed to be used if amyl acetate was to be used. The directions did not give any reason for using it. I have usually used Acetone for dehydrations and a freshly opened bottle of Mallinckrodt #2440 acetone for the final exchange before the drying.
Many years ago an investigator insisted on using amyl acetate with his sample and he ran the operation himself. He later admitted that similar samples that I ran in acetone looked better.
If one uses amyl acetate in the critical point dryer it is extremely necessary for the amyl acetate to be properly vented into a hood (acetone also). If the amyl acetate of this volume gets into the room it can make one behave strangely. My co-workers thought that I was drunk! I did not make that mistake again. Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu =========================== } I have not used amyl acetate for CPD since my first CPD run almost } 30 years ago. } The head of department was incensed by the strong pear/banana smell } and banned it as } antisocial.In those days nobody seemed to worry about its toxicity. } Amyl acetate also } damages the rubber seals of the CPD faster than other solvents. } } The objective is to fully replace the dehydrating solvent with liquid CO2. } Amyl acetate is more soluble in liquid CO2 than the more polar ethanol, but } many people nevertheless use ethanol routinely and obtain great } results, especially if } the specimens are small, thin, and unenclosed by cuticle (cultured } cells on coverslips, } for example). If the specimens are large chunks of plant material it } would be better to } work with acetone (propanone) as both dehydrating and intermediate solvent. } } Dr. Chris Jeffree {c.jeffree-at-ed.ac.uk} } University of Edinburgh ============================ } } Email: mark.evans-at-uvm.edu } } Name: Mark Evans } } Organization: University of Vermont } } Title-Subject: [Microscopy] Isoamyl acetate and critical point drying } } } } Question: Some protocols for sem indicate that following ethanol } } series dehydration samples should be soaked in isoamyl acetate and } } that this media should also be used for critical point drying. Can } } anyone tell me what the theory is behind using isoamyl acetate and } } what benefit, if any, there is over leaving samples in 100% ethanol } } prior to the critical point drying? } } Thanks, } } Mark } } ---------------------------------------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 12:36:11 2004
We have been doing replicating products since the 1950s, so if you wish to discuss replication please call Mike Bouchard at 1-800-451-3406 and he'll be glad to provide assistance on replicating. Mike has many years of experience in this area.
Thanks, Deb Sicard
Disclaimer: Ladd has been a supplier of EM products, including replicating materials, for more than 50 years
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: ladres-at-att.net
----- Original Message ----- } From: "White, Woody N." {nwwhite-at-bwxt.com} To: "'Yang, Ann-Fook'" {YANGA-at-agr.gc.ca} ; {microscopy-at-microscopy.com} Sent: Wednesday, September 29, 2004 5:06 PM
We routinely use acetate replicas from machined metal surfaces that resolve to the micron level.
Can you describe the procedure used by Milos? Perhaps it differs from what we do.
Regards, Woody
-----Original Message----- } From: Yang, Ann-Fook [mailto:YANGA-at-agr.gc.ca] Sent: Wednesday, September 29, 2004 1:01 PM To: microscopy-at-microscopy.com
Hi everyone,
Can you help my colleague, Milos Kalab, to solve his problem. His question is posted below. Thank you.
Ann Fook Yang EM Unit/ Unite EM Agriculture and Agri-Food Canada/Agriculture et Agroalimentaire Canada Telephone/Téléphone: 613-759-1638 Facsimile/Télécopieur: 613-759-1701 960 Carling Ave/960 Boul Carling Ottawa,Ontario/Ottawa, Ontario Canada K1A 0C6 yanga-at-agr.gc.ca
My objective is to obtain SEM images of stainless steel and other surfaces used in food processing. For this reason I wish to replicate small parts of the surfaces and examine the negative replicas. Advice on materials and procedures which would reproduce details down to 1 micrometer would be appreciated. (I am aware of Ted Pella and SPI cellulose acetate replication materials for coarser surfaces and I wonder why I have not been successful.)
Thank you.
Milos Kalab
Agriculture and Agri-Food Canada in Ottawa
kalabm-at-agr.gc.ca
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 15:02:08 2004
I can offer only the comments of an experimenting amateur, with a Nikon Coolpix 990. I am using GNU/Linux as an operating system, and have had a modicum of success with setting up several programs to manipulate the camera's controls. It's exciting enough to encourage me to do further experiments with a newer model of Coolpix.
I take photos through the eyepiece, using a relatively inexpensive coupler that is merely a clamp to hold the camera onto the eyepiece. Several companies manufacture these, as well as optical couplers (so-called relay lenses) which take the place of an eyepiece. (I don't understand the difference between relay lenses and the projection lenses of the past).
I really like that the coolpix can be monitored in real time through the TV, which works wonders for me, as a teacher, in teaching about microscopical realms without having student microscopes at our school. Using some tips for setting up this camera for microscopy (more than one set of which I found on the Inet) I find it relatively easy to take at least snapshots. With a cable release, and a bright enough light---perhaps I will finally get around to setting up a strobe---the results promise to be able to freeze motion even of cilia.
A promising suggestion was to use a video capture card to move the video image to the computer screen while using software to remotely fire the camera. I don't know whether this is possible, but I hope to try it. I can see why the Coolpix 990 has received so much attention from the microscopical community. It is available relatively cheaply on ebay, too, so perhaps this could serve as the basis for some experiments for you, too.
I am doubly encouraged at how well the camera is supported by a GIMP (Gnu Image Manipulation Program---a photoshop like program) plugin. Better than any other software, so far, I have found it easy to even monitor the image in *pretty much* real time and snap on demand, at which point the image is immediately available for cropping and editing. The program is free (in terms of your being able to share copies---in terms, in other words, of freeDOM, as well as free BEER freedom). This, again, encourages my further experimentation. I am considering the upcoming Nikon Coolpix 8800 which has vibration reduction technology included!
Take my comments with the appropriate grain of salt.
Alan Davis Kagman High School Saipan, Northern Mariana Islands aedavis-at-eccomm.com
On Thu, 30 Sep 2004 08:02:31 -0500 Clarkson Donna R Contr USAMRD/MCMR {donna.clarkson-at-brooks.af.mil} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Hi, Peggy! } } We do quite a lot of LM digital imaging these days. Through Nikon we had } purchased the Photometrics Cool Snap cf camera to place atop an Olympus BH-2 } light microscope. The software program we're using is called Metavue; it } allows you to view live and make changes to lighting and such before saving. } The best part is being able to make much smaller files by making duplicate } images and saving into TIFF. It also allows you to put in micron bars after } calibrating your scope and camera system together. Afterwards, the } duplicated images are able to be used in Photoshop or Power Point. } } Good luck in your search! } } *Disclaimer--I do not work for, or receive any compensation from, Nikon or } the makers of Metavue. We are just satisfied with the setup. } } Donna R. Clarkson, HT (ASCP) } Northrop Grumman Information Technology } for U S Army Medical Research Detachment } at Brooks City-Base } Phone (210) 536-1416 } FAX (210) 536-1449 } e-mail donna.clarkson-at-brooks.af.mil } "Our Army at War--Relevant and Ready" } } } } -----Original Message----- } } From: Miller, Margaret M [mailto:MILLERMM-at-uthscsa.edu] } Sent: Wednesday, September 29, 2004 12:25 PM } To: Microscopy-at-microscopy.com } Subject: [Microscopy] LM digital imaging } } } } } ---------------------------------------------------------------------------- } -- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- } --- } } Dear Listserver, } } Is any one using or know of a consumer camera with full computer control } and live video preview mounted on an optical microscope? } } Peggy Miller } UTHSCSA Ophthalmology } Lions Sight Research Center } 7703 Floyd Curl Drive MC6230 } San Antonio, Texas 78229-3900 } PH: (210)567-8460 } FAX:(210)567-8413 } } }
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 16:51:56 2004
Last year a couple of students from BYU Hawaii were interested in looking for copper in the mitochondria of yeast by EELS and/or ESI, using our LEO 912 EFTEM. The first (but by no means the only) stumbling block was getting decent ultrastructure, or at least good enough that they could identify the mitochondria, and good enough infiltration that 25-30 nm sections could be obtained.
I do not have any experience with yeast and so I am open to any and all hints, tips, and suggestions!
If this works out, I invite you all to come to see the results at M&M 2005 in Honolulu next August.
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 17:57:15 2004
After years of trying desperately to get one of our microtomes working, I'm thinking of changing the company that provides service on our instruments (2 Reichert Ultracut E and 2 Leica UCT/FCS). Does anyone in the NorthEast know of someone capable of providing service in Connecticut? I'm especially interested in someone with good experience with the Leica instruments. Thank you
Marc -- Marc Pypaert Department of Cell Biology Center for Cell and Molecular Imaging Ludwig Institute for Cancer Research Yale University School of Medicine 333 Cedar Street, PO Box 208002 New Haven, CT 06520-8002 TEL 203-785 3681 FAX 203-785 7446
From MicroscopyL-request-at-ns.microscopy.com Thu Sep 30 21:23:11 2004
See http://http://www.math.ualberta.ca/imaging/ There, one might find these remarks:
"... newer cameras (Coolpix 995/775 and beyond) can only be controlled via a serial cable"
There's more to all this than I understand.
Alan
On Thu, 30 Sep 2004 20:58:25 -0400 John Twilley {jtwilley-at-sprynet.com} wrote:
} Dear Alan, } } Do I understand you correctly that the 990 has some input for control by a PC? Is this true of the 995 as well? Are you writing this control routine yourself? } } John Twilley } } Alan E. Davis wrote: } } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } ------------------------------------------------------------------------------- } } } } I can offer only the comments of an experimenting amateur, with a Nikon Coolpix 990. I am using GNU/Linux as an operating system, and have had a modicum of success with setting up several programs to manipulate the camera's controls. It's exciting enough to encourage me to do further experiments with a newer model of Coolpix. } } } } I take photos through the eyepiece, using a relatively inexpensive coupler that is merely a clamp to hold the camera onto the eyepiece. Several companies manufacture these, as well as optical couplers (so-called relay lenses) which take the place of an eyepiece. (I don't understand the difference between relay lenses and the projection lenses of the past). } } } } I really like that the coolpix can be monitored in real time through the TV, which works wonders for me, as a teacher, in teaching about microscopical realms without having student microscopes at our school. Using some tips for setting up this camera for microscopy (more than one set of which I found on the Inet) I find it relatively easy to take at least snapshots. With a cable release, and a bright enough light---perhaps I will finally get around to setting up a strobe---the results promise to be able to freeze motion even of cilia. } } } } A promising suggestion was to use a video capture card to move the video image to the computer screen while using software to remotely fire the camera. I don't know whether this is possible, but I hope to try it. I can see why the Coolpix 990 has received so much attention from the microscopical community. It is available relatively cheaply on ebay, too, so perhaps this could serve as the basis for some experiments for you, too. } } } } I am doubly encouraged at how well the camera is supported by a GIMP (Gnu Image Manipulation Program---a photoshop like program) plugin. Better than any other software, so far, I have found it easy to even monitor the image in *pretty much* real time and snap on demand, at which point the image is immediately available for cropping and editing. The program is free (in terms of your being able to share copies---in terms, in other words, of freeDOM, as well as free BEER freedom). This, again, encourages my further experimentation. I am considering the upcoming Nikon Coolpix 8800 which has vibration reduction technology included! } } } } Take my comments with the appropriate grain of salt. } } } } Alan Davis } } Kagman High School } } Saipan, Northern Mariana Islands } } aedavis-at-eccomm.com } } } } On Thu, 30 Sep 2004 08:02:31 -0500 } } Clarkson Donna R Contr USAMRD/MCMR {donna.clarkson-at-brooks.af.mil} wrote: } } } } } } } } } } } ------------------------------------------------------------------------------ } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } ------------------------------------------------------------------------------- } } } } } } Hi, Peggy! } } } } } } We do quite a lot of LM digital imaging these days. Through Nikon we had } } } purchased the Photometrics Cool Snap cf camera to place atop an Olympus BH-2 } } } light microscope. The software program we're using is called Metavue; it } } } allows you to view live and make changes to lighting and such before saving. } } } The best part is being able to make much smaller files by making duplicate } } } images and saving into TIFF. It also allows you to put in micron bars after } } } calibrating your scope and camera system together. Afterwards, the } } } duplicated images are able to be used in Photoshop or Power Point. } } } } } } Good luck in your search! } } } } } } *Disclaimer--I do not work for, or receive any compensation from, Nikon or } } } the makers of Metavue. We are just satisfied with the setup. } } } } } } Donna R. Clarkson, HT (ASCP) } } } Northrop Grumman Information Technology } } } for U S Army Medical Research Detachment } } } at Brooks City-Base } } } Phone (210) 536-1416 } } } FAX (210) 536-1449 } } } e-mail donna.clarkson-at-brooks.af.mil } } } "Our Army at War--Relevant and Ready" } } } } } } } } } } } } -----Original Message----- } } } } From: Miller, Margaret M [mailto:MILLERMM-at-uthscsa.edu] } } } Sent: Wednesday, September 29, 2004 12:25 PM } } } To: Microscopy-at-microscopy.com } } } Subject: [Microscopy] LM digital imaging } } } } } } } } } } } } } } } ---------------------------------------------------------------------------- } } } -- } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } } To Subscribe/Unsubscribe -- } } } http://www.msa.microscopy.com/MicroscopyListserver } } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } ---------------------------------------------------------------------------- } } } --- } } } } } } Dear Listserver, } } } } } } Is any one using or know of a consumer camera with full computer control } } } and live video preview mounted on an optical microscope? } } } } } } Peggy Miller } } } UTHSCSA Ophthalmology } } } Lions Sight Research Center } } } 7703 Floyd Curl Drive MC6230 } } } San Antonio, Texas 78229-3900 } } } PH: (210)567-8460 } } } FAX:(210)567-8413 } } } } } } } } } } } }
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