You can not use glass for this kind of application and SPI Supplies has offered a full range of quartz slides and cover slips designed to solve this problem on URL http://www.2spi.com/catalog/ltmic/quartz.shtml
They are generally available from stock for overnight shipment worldwide.
One should note that, like mica, if you don't know the grade you are purchasing, there is no way to tell what you actually do have. So from the above page, you can see that we feature GE 124 quartz in the fabrication of our quartz slides and cover slips. And full transmittance data is disclosed. See URL http://www.2spi.com/catalog/ltmic/quartz-slides-coverslips.html
Whoever you do purchase from, if not from SPI, make sure you know what quartz you are getting and its characteristics.
One further comment: For the near infrared, MgO might be a better alternative. See URL http://www.2spi.com/catalog/submat/magnesium-oxide.shtml and to the graph showing the transmittance spectrum for MgO.
The absorption curves are given for both materials so one can match product selection with objectives in terms of what part of the spectrum they need their greatest transparency.
Disclaimer: SPI Supplies has offered a full range of quartz slides and cover slips to those working in the UV and near infrared range for some number of years. We over standard shapes and sizes but can also produce custom shapes and sizes.
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 Sat Nov 1 14:07:49 2003
I would like to know the origin of terms of lacey carbon and holey carbon films. When I was in Japan, most people use microgrid to refer carbon films having many holes. I have not yet investigated who developed such carbon films. However, I think that Fukami, Adachi, Sakata, and Sjostrand.
Papers written by Prof. Akira Fukami are: Fukami and Adachi (1965); J. Electron Microscopy, 14, 112-118. Fukami, Adachi, Katoh (1966) The proceedings of 6th International Congress for Electron Microscopy, Vol. 1, 263-264.
Later, he wrote “Specimen Preparation Techniques for Electron Microscopy” in 1967 (Japan Electron Optics Laboratory Co., LTD.), based on the above two papers.
Prof. Fukami used a term “Micro Plastic Grid” in his 1967 paper. I do not know who changes it to microgrid, but Japanese Electron Microscopy Dictionary (1986) edited by Hashimoto and Ogawa uses “microgrid”. Some Japanese textbooks I used in Japan also used "microgrod" or "micro grid".
I think that the preparation method of holey and lacey carbon films is the same as that of microgrid. Japanese prefer to use "microgrid", but most researchers outside Japan use "holey or lacey carbon films". I would like to know historical reason international researchers use holey or lacey carbon films instead of microgrid.
I use “holey or lacey carbon” in my papers because most people do not know “microgrid” and the venders use such terms. However, if “microgrid" is not strange English, I think that people should use microgrid, assuming that people basically follow Fukami’s method to prepare such films.
Thank you,
Hiromi Konishi, Ph.D. Dept. of Earth and Planetary Sciences The University of New Mexico
From MicroscopyL-request-at-ns.microscopy.com Sun Nov 2 14:41:40 2003
Here is the table of contents for the November/December 2003 issue of Microscopy Today. This issue is mailed with the Call for Papers for the 2004 Microscopy and Microanalysis meeting
New Subscriptions via http://www.microscopy-today.com only, please New subscriptions will close on Tuesday 11 November for this issue.
There has been a major change in subscription policies coming out of the MSA Winter Council meeting. Briefly: Canadians and Mexicans are now offered free subscriptions along with microscopists in the USA. MSA members anywhere have free subscriptions. Non-MSA, non-North American subscriptions have been reduced from $80 or $110US to $35US. Additional details in the magazine or on our web site.
THIS WILL BE THE LAST ISSUE NON-MSA MEMBER, NON-NORTH AMERICAN, UNSUBSCRIBED OR NOT-CURRENTLY-SUBSCRIBED INDIVIDUALS WILL RECEIVE!
Table of Contents:
Zaluzec The Scanning Confocal Electron Microscope Marko, et al. Correlative Electron Tomography And Elemental Microanalysis In Biology: A Preview Sedgewick Creating Pseudocolored Images in Photoshop Steigerwald Ultra Low Voltage BSE Imaging Fasolka, et al. Techniques for Combinatorial and High-Throughput Microscopy Part 2: Automated Optical Microscopy Platform for Thin h Mustoe Microscopy of Silicified Wood Gerrity, et al. Microwave Processing in Diagnostic Electron Microscopy Shribak and Oldenbourg A Polarizing Microscope for Mapping Birefringent Objects in 3D Space Casavan and Gaidoukevitch Colocalization of Fluorescent Probes Using Image-ProR Plus v. 5.0 Munroe Electron Microscopy in Australia Tengowski Converting Right-Left Stereo Pairs Into Colored Pairs For Electronic Presentation Ahlstrand Cellulose Acetate Replication of Plant Surfaces for SEM Ellis Safe Handling of Embedding Media Schooley Microscopes as Gifts MSA Council Microscopy Society of America Position on Ethical Digital Imaging
Ron Anderson, Editor Microscopy Today
From MicroscopyL-request-at-ns.microscopy.com Sun Nov 2 14:45:19 2003
I have 120 square feet for a JXA-840A EPMA, including sample loading and storage of spares, etc and boy, let me tell you, it's not enough!
I wish I had been more assertive.............
cheers
rtch
} From: Garry Burgess {GBurgess-at-exchange.hsc.mb.ca} To: MSA {microscopy-at-msa.microscopy.com}
Listers,
Here is the table of contents for the November/December 2003 issue of Microscopy Today. This issue is mailed with the Call for Papers for the 2004 Microscopy and Microanalysis meeting
New Subscriptions via http://www.microscopy-today.com only, please New subscriptions will close on Tuesday 11 November for this issue.
There has been a major change in subscription policies coming out of the MSA Winter Council meeting.
Briefly: Canadians and Mexicans are now offered free subscriptions along with microscopists in the USA.
MSA members anywhere have free subscriptions.
Non-MSA, non-North American subscriptions have been reduced from $80 or $110US to $35US. Additional details in the magazine or on our web site.
THIS WILL BE THE LAST ISSUE NON-MSA MEMBER, NON-NORTH AMERICAN, UNSUBSCRIBED OR NOT-CURRENTLY-SUBSCRIBED INDIVIDUALS WILL RECEIVE!
Table of Contents: November/December 2003
Zaluzec-The Scanning Confocal Electron Microscope Marko, et al.-Correlative Electron Tomography And Elemental Microanalysis In Biology: A Preview Sedgewick-Creating Pseudocolored Images in Photoshop Steigerwald-Ultra Low Voltage BSE Imaging Fasolka, et al.-Techniques for Combinatorial and High-Throughput Microscopy Part 2: Automated Optical Microscopy Platform for Thin h Mustoe-Microscopy of Silicified Wood Gerrity, et al.-Microwave Processing in Diagnostic Electron Microscopy Shribak and Oldenbourg-A Polarizing Microscope for Mapping Birefringent Objects in 3D Space Casavan and Gaidoukevitch-Colocalization of Fluorescent Probes Using Image-ProR Plus v. 5.0 Munroe-Electron Microscopy in Australia Tengowski-Converting Right-Left Stereo Pairs Into Colored Pairs For Electronic Presentation Ahlstrand-Cellulose Acetate Replication of Plant Surfaces for SEM Ellis-Safe Handling of Embedding Media Schooley-Microscopes as Gifts MSA Council-Microscopy Society of America Position on Ethical Digital Imaging
Ron Anderson, Editor Microscopy Today
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 3 00:59:31 2003
Dear Garry Depends what you are referring to. This question is currently wrecking my sleep. I need more space! We have a multiuser facility in the University of Botswana and we are training students (sadly the academics are not interested in learning) to operate the scopes. There is a possibility of a new building somewhere in the distant future and for the planning phase I had to give some input. I will give real distances rather that square meter since I found the real distances (Shape) of the room is important. For the TEM we currently have 4.63m X 3.25m which seems adequate as long as there is not more that 3 people in the room at the same time. Two people work fine but it does get crowded with 3. Often it is the operator + client sometimes the supervisor as well. We do not store samples in the room and except for loading space the rest is dedicated to the scope (Technai 12). The ESEM room we have is a bit to small. We need more space on the sides to walk to the bac where the pumps and electronics are (XL 30 ESEM). We can not fit a cryo, the room is just to small. 3.25m X 4.02 currently a 4x4m room will be my minimum requirement for an SEM. The confocal room is 2.08 X 2.27 it just does not work! 3 X 3 (Just for the confocal) will work. (Again there will be no spare space for "clients"to be involved. All sample prep must take place outside the microscope rooms. The biological sample prep area is 3.25 X 3m. It work fine for a max of 2 people in the area at the same time. The CPD and Sputter coater fit. All chemicals must be stored at another location. We would like to have a microwave in there as well but the room is to small. Thus my suggestion is 4X5m room for biological sample prep. and the same size for materials science prep. 4x3 for light microscopy. assuming 2 stereo and 1 transmitted light as well as one inverted light microscope. Space for the PC for each scope to store images on and the high end scopes are computer driven. Still you need storage space and a general work area gass bottle storage, UPS storage (we have 3 9kvA units)and most EM units have a small collection of books. To get this amount of space in a country is not an easy task. Then there is the long term planning of adding/expanding? Just hope this is useful. Remember these are my personal views.
-----Original Message----- } From: Garry Burgess [mailto:GBurgess-at-exchange.hsc.mb.ca] Sent: Friday, October 31, 2003 9:50 PM To: MSA
Does anyone know the recommended floor space suitable for an electron microscopy lab? I'm interested in the requirements for both research and also hospital labs space requirements - or at least the recommended square footage requirements.
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 3 07:03:49 2003
We used to buy these from Ernest F. Fullam and SPI.
Fullam's shows a Rh and a Pd 'back coated' Cu grid in the old red catalog.
SPI Inc. also sells them, 2020P.
I am sure other EM suppliers do also.
Paul
-----Original Message----- } From: Philip Oshel [mailto:peoshel-at-wisc.edu] Sent: Thursday, October 30, 2003 12:46 PM To: Microscopy-at-sparc5.microscopy.com
Micromavens,
One of our lab members was looking for bimetallic grids: Cu on one side and Ni on the other. She couldn't remember the supplier, but remembered using them -- within the last couple of years. I've gone on the web and checked all the usual suspects to no avail. Has any one heard of such grids, and knows who supplies them? Thanks.
Phil -- 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 Mon Nov 3 07:35:58 2003
Dear all, Is there a standard reference book on TEM of semiconductors? A grad student here wants to look for info on this topic and a good text would be a very helpful starting place.
Please copy your answers to sigmund-at-hf.tu-darmstadt.de
Thanks
-- Ian MacLaren Technische Universität Darmstadt Material- und Geowissenschaften Petersenstr. 23 64287 Darmstadt Germany http://www.tu-darmstadt.de/fb/ms/fg/sf/projekte/maclaren-Dateien/maclaren.html
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 3 08:53:10 2003
Garry Try to get your hands on a copy of "Practical Methods in Electron Microscopy, Vol.4: Design of the Electron Microcope Laboratory" by Ronald H Alderson. It is part of the series edited by Audrey M. Glauert. Even thought it is over 25 years old, it is full of useful information on this topic. 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 Nov 3 08:53:51 2003
Hi, Anyone out there using a JEOL 6300 and using an inexpensive frame grabber to capture the video output? If so can you provide me with any info on manufacturer, model number, and performance?
TIA Mike -- ******************************************************************** Michael M. Cheatham 312 Heroy Geology Laboratory Phone (315)-443-1261 Syracuse University Fax (315)-443-3363 Syracuse, NY 13244-1070
owner of PLASMACHEM-L: http://listserv.syr.edu/archives/plasmachem-l.html owner of XRF-L: http://listserv.syr.edu/archives/xrf-l.html owner of TIMS-L: http://listserv.syr.edu/archives/tims-l.html owner of SIRIS-L: http://listserv.syr.edu/archives/siris-l.html ********************************************************************
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 3 12:16:30 2003
Background Information: We have a JEOL 6460LV which was installed about 6 months ago. Recently we have noticed a significant degradation in image quality. We have had field service come in and clean the column and the apertures twice. The response they gave us on both occasions was that this is normal for this instrument.
Normal use and operation is HV and low KV, {2. When LV mode is used, the samples are clean and LV is only used to get rid of charging. "Dirty" samples are never placed into the chamber. Primary use of the instrument is polymer imaging.
The Question: What is the expected frequency of final aperture cleaning, given the above stated operating conditions? (This one is seriously starting to go downhill after about 1.5 months of operation). Previous experience with a JEOL 5900 was that after 6 months there was no significant performance loss. The apertures of other microscopes, i.e., AMR, ISI, TOPCON, were changed when LaB6 filaments were changed. That is after ~4 to 6 months of continuous use. Thanks Nancy Simons,
Molecular Interventions Boston Scientific Corp. 1 Boston Scientific Place A4 Natick, MA 01760-1537
tel. 508-650-8603 fax 508-647-2329
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 3 12:27:26 2003
you can pretty much use any frame grabber that understands the video protocol the machine is using (I think it's RS-170, but I'm not sure. Check the manuals) to acquire images from the SEM. There are many drawbacks using this technique, though:
1) Your images will be relatively low resolution (640x480), due to the limitations of video signals. 2) It is possible that you can only acquire the images recorded at video speed. They are typically noisy compared to slow scan images. 3) You will have no calibration information, unless you can also acquire the data bar and calibrate the images that way. 4) Your images will be 8-bit images (256 levels of gray).
Item 2 you can improve through frame averaging, but the frame grabber and software must support that.
If you need more than that, you need to consider SEM interfaces, such as our ADDA II (there are other manufacturers as well). These devices can acquire slow scan signals at a higher resolution (up to 4Kx4K), and they are usually better than 8 bit (12-bit in the case of the ADDA).
If you need more information, please contact me off-line.
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: Michael Cheatham [mailto:mmcheath-at-mailbox.syr.edu] Sent: Monday, November 03, 2003 08:03 To: microscopy-at-ns.microscopy.com
Hi, Anyone out there using a JEOL 6300 and using an inexpensive frame grabber to capture the video output? If so can you provide me with any info on manufacturer, model number, and performance?
TIA Mike -- ******************************************************************** Michael M. Cheatham 312 Heroy Geology Laboratory Phone (315)-443-1261 Syracuse University Fax (315)-443-3363 Syracuse, NY 13244-1070
owner of PLASMACHEM-L: http://listserv.syr.edu/archives/plasmachem-l.html owner of XRF-L: http://listserv.syr.edu/archives/xrf-l.html owner of TIMS-L: http://listserv.syr.edu/archives/tims-l.html owner of SIRIS-L: http://listserv.syr.edu/archives/siris-l.html ********************************************************************
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 3 18:56:54 2003
You may also want to consider utility rooms or corridors for noisy pumps and chillers if space allows. We had an opportunity to setup a lab with the ability to place utility rooms adjacent to the microscope rooms and it makes for a much enjoyable place to work. For our Philips CM200 we also have the power supply in the utility corridor, mechanical pumps are not a problem because they don't run all the time. Our Hitachi S-4700 mechanical pumps do run all the time and I'm glad they are not in the same room. I would prefer smaller rooms with a utility corridor over a large room.
-- David R. Hull NASA Glenn Research Center at Lewis Field Advanced Metallics Branch Mail Stop 49-1 21000 Brookpark Road Cleveland, OH 44135
I couldn't agree more about Ronald Alderson's book - I have found it very useful on several occasions in the past. There have been some changes over the years such as digital imaging so you may or may not need a darkroom for instance.
One final source of information about lab space that is the microscope manufacturer's data at the back of their brochures/literature. This will give you the basic minimum for the instrument and then you would need to factor in people and preparation space. A difficult one is always allowing for occasional groups of visitors - in my case I am limited to groups of about 10 people because there are several small rooms in the lab.
I know it's not high-tech but it's sometimes useful to sit down with a piece of graph paper and some cut-outs of the instruments going into the lab space. This is very useful for highlighting narrow access points (eg access to the backs of instruments for serving).
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: Leona Cohen-Gould {lcgould-at-med.cornell.edu}
Less than 1 year old, Zeiss has agreed to service it on a per call basis. (But not install it) , I have someone to install it, and people who can teach operational in house software maintenance.
We have recently moved into a new building that includes a spacious EM facility. I agree with David about the utility room. Not only are the noises of the pumps and chillers out of the room (The EM users get to sit in a wonderfully quiet space), but we have less problem with temperature fluctuations in the EM room, because all that happens on the other side of the wall. Lastly, this keeps the dirty work--oil changes etc --away from the scope. We also have placed the LN2 tank in this room. If you have the space and flexibility, I fully recommend it.
Our engineers recommendeded at least 3' on each side and the back of the console to allow for access for service. (In our old building, there was precious little space for even the thinnest engineers). The amount of space on the operator side probably depends on how you use the space. We went from 3 feet of space (you could not fully open the door while a user was seated) to 10 ft of space between the console and the wall. This has left plenty of room for door sweeps, a small work table, and room for my class of 12 to sit and watch a demonstration.
Don
On Mon, 3 Nov 2003, David R Hull 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 } ------------------------------------------------------------------------------- } } You may also want to consider utility rooms or corridors for noisy pumps and chillers if space allows. We had an opportunity to setup a lab with the ability to place utility rooms adjacent to the microscope rooms and it makes for a much enjoyable place to work. For our Philips CM200 we also have the power supply in the utility corridor, mechanical pumps are not a problem because they don't run all the time. Our Hitachi S-4700 mechanical pumps do run all the time and I'm glad they are not in the same room. I would prefer smaller rooms with a utility corridor over a large room. } } -- } David R. Hull } NASA Glenn Research Center at Lewis Field } Advanced Metallics Branch } Mail Stop 49-1 } 21000 Brookpark Road } Cleveland, OH 44135 } } (216) 433-3281 } fax (216)977- 7132 } david.r.hull-at-nasa.gov } http://www.lerc.nasa.gov/WWW/AdvMet/ASGWEB2000/asghome.html } } }
______________________________________________________________________ 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 Tue Nov 4 08:15:23 2003
All Some kind soul has presented me with samples of debris which are very small (I would guesstimate {10 microns). These are stuck to magic tape and then onto paper. What I now need to do to prepare for EDX analysis is to get them out of the adhesive of the tape... On ordinary tape, the adhesive, I have found, can be dissolved using chloroform (acetone just makes a sticky goo). The Scotch Magic Tape appears different though and I am left with a sticky goo again. Question: Anyone any idea how I can dissolve this adhesive?
Regards Adrian Jenkinson
__________________________________ Do you Yahoo!? Protect your identity with Yahoo! Mail AddressGuard http://antispam.yahoo.com/whatsnewfree
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 4 08:54:30 2003
All, Let me add couple cents to the discussion. We have moved to a new lab in January. The lab was designed into a old FAB space so we did not worry about vibrations etc. When we were designing the facility we went ahead and used the manufacturers brochures as a minimum requirements. After hearing our opinions, the interior designer forced the rooms to be exactly the dimensions provided by FEI for their scopes. Lesson 1: plan and fight for as much room as possible within your budget.
Our two TEMs were disjoined into separate rooms. JEM2010 fits nicely and there is plenty of space to move around. Our mechanical pumps and chillers were relocated to separate utility rooms (great idea!!!), power supplies needed to stay in microscope rooms due to cabling distance. Works great!
Our CM300 on the other hand was shoehorned into Technai F30 minimum room. The mechanicals of the microscopes are essentially the same, so it looked as a good fit in view of future upgrade. Besides, we could not get the original spec from FEI for the CM.
Lesson 2: What went wrong? The two scopes have different power supply, chiller, and pump requirements, we again were able to re-locate mechanical pump and chillers to utility room (great), but the power supplies need to stay in the main room (boo). Now we need to put up with fan noise coming out of the main PS (we used to live with it in the old lab, but now we actually notice it). Annoying but not detrimental to daily operation. Moreover, we did not realize that FEI made minimum requirement for the height of the room different. In order to disassemble the FEG, the whole assembly needs to be lifted on a crane supplied with the microscope. With Techani room, we are 1" short of the height. Now each time FEI works on the gun, we need to remove soft panels from the ceiling and essentially open up a large AC plenum that is above the whole lab. This dumps a lot of particulate and dust into the room since this bypasses the filters. That's a NO-NO when working on a ultra high vacuum systems ! and something to take into account when designing air flows. I get ribbed for it each time FEI service comes in.
Our SEM, FIB, and SIMS tools were relocated into new rooms doubling up. The used to be in a large ball-room setting in the old lab. This tremendously helps in facilitation of work and nap times. Great idea, stay away from ballrooms even with curtains around tools. One thing to take into consideration is the selection of tools that are put together. We tried to pair a daily workhorse with a tool that was specialized and used only on occasion. This helps to reduce interference between operators or service.
Hope this helps.
Jerzy
****************************************************** Jerzy Gazda, Ph.D. Advanced Micro Devices Supervising Engineer 5204 E. Ben White Blvd. - MS 512 PCAL - AIM Section Austin, TX 78741 TEL: 1-800-538-8450, Ext. 51453 jerzy.gazda-at-amd.com ******************************************************
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 4 15:28:08 2003
During my exploration of adhesives to make some "poor man's" grid glue, I only used chloroform. Bozzola discusses the use of ethylene dichloride as your solvent. I believe you can trial-and-error with most any organic solvent. I found that with enough time, the backing on any of the tapes we had in the lab became a "sticky goo" (is that a scientific term?) in the chloroform.
Good luck and let us know what you come up with!
Chris
----------------------------------------------------------------------- Christopher S. Zurenko Research Assistant II Kresge Hearing Research Institute, Otopathology The University of Michigan Medical School MSRB 3, Room 9303 1150 W. Medical Center Dr. Ann Arbor, MI 48109-0648 Lab Phone: 734.763.9680 Fax: 734.615.8111 czurenko-at-umich.edu http://www.khri.med.umich.edu/research/raphael_lab/index.htm
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 4 16:10:46 2003
This meeting is sponsored by the Midwest Microscopy & Microanalysis Society (MMMS) in association with the ASM Chicago Chapter and will be held at Harper College in Palatine, Illinois on Friday November 14th. Pre-registration is open through Friday November 7th. For a registration form please contact Robb Mierzwa (see below).
For further information including hotel accommodations please contact either:
Robert Mierzwa (MMMS President - Elect): Tel: 920-803-8945, email: mierzwa-at-jeol.com Arvid Casler (MMMS Program Coordinator): Tel: 847-674-7700, email: arvid_casler-at-fmo.com Jim DiOrio (MMMS President): Tel: 847-270-4676, email: jim_diorio-at-baxter.com
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 4 18:10:11 2003
} We also have placed the LN2 tank in this room. If you have } the space and flexibility, I fully recommend it. }
Always dreaming up the worst scenarios and I apologise in advance for that. However:
How big and how well ventilated should a separate "utility room" be if an LN2 tank is being stored in there? How big is the LN2 tank? Also, where are the LN2 transfers from that larger vessel into smaller hand-held dewars being done? If you are storing a large LN2 tank that lets out a "lot" of nitrogen into that room over time, and it is a room that is normally closed up nearly all the time because "nobody goes in there that much", then could there be a slight risk of too much air (oxygen) getting displaced? Maybe this could be a potential problem if there is no formal ventilation in that room, or if the room is only serviced as part of a closed A/C system.
There have been a few postings on this list in past years relating tales of air displacement by evaporating LN2 in confined spaces. Perhaps this is also something that should be considered when thinking about where to put stuff associated with running EMs these days.
Just my paranoid thought for the day!
Arthur Day, Electron Microscope Unit Phone: 61-2-9717-3457 Ansto Materials Division Fax: 61-2-9543-7179 PMB 1, Menai (Sydney), NSW, 2234 Email: ard-at-ansto.gov.au Australia www: http://www.ansto.gov.au/
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 4 18:10:45 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (anjeanette.ormonde-at-unilever.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, November 4, 2003 at 14:44:59 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] MListserver: Dismantling TEM
Question: We have a Philips 400 TEM that we are no longer going to be keeping. Right now it is under vacuum and everything is working well. However, we do not use the system enough to justify the costs of keeping it (we let the service contract expire in Dec. 2002 but even beyond that depreciation costs come out of our budget). Anyhow, I was not able to find anyone who wanted the system (free to a good home!) so I am (sadly) faced with dismantling and discarding the system. However, I have no experience with this. Are there components we can separate out and either keep or give to others? What happens with unwanted pieces? Can most things go out with regular garbage? Can anything be recycled? Thanks in advance for any advice or tips you can offer.
Adrian Jenkinson wrote: ======================================================= Some kind soul has presented me with samples of debris which are very small (I would guesstimate {10 microns). These are stuck to magic tape and then onto paper. What I now need to do to prepare for EDX analysis is to get them out of the adhesive of the tape... On ordinary tape, the adhesive, I have found, can be dissolved using chloroform (acetone just makes a sticky goo). The Scotch Magic Tape appears different though and I am left with a sticky goo again. Question: Anyone any idea how I can dissolve this adhesive? ======================================================= The fact that you don't worry about the chloroform either dissolving or in some other way altering the particles, would suggest that the particles are either metal or ceramic, neither of which will be affected by an oxygen plasma.
Therefore, I would suggest putting the particles, goo and all into a glass petri dish and placing the entire dish with gooey particles into a plasma etcher, such as the SPI Plasma Prep II plasma etcher (see URL http://www.2spi.com/catalog/instruments/etchers1.shtml ).
The oxygen plasma will remove all traces of the organics while leaving the inorganics behind undisturbed and unchanged (from the oxygen plasma).
What is left is a dry residue on the bottom of the glass petri dish.
In our own laboratory, we would take an SEM mount, apply a double sided conductive carbon adhesive disc such as on URL http://www.2spi.com/catalog/spec_prep/cond_adhes-discs.shtml and then either a) sprinkle on the dry ashed residue particles onto the surface of the adhesive or if static or other forces keep the particles from being released from the glass, then use either b) a Zerostat antistatic gun (see URL http://www.zerostat.com ) to cause the particles to release and if that does not work, then c) press the SEM mount and adhesive gently onto the particles on the glass surface which then will be efficiently lifted off and will be attached to the SEM mount, ready for carbon coating and viewing.
If you do not have a plasma etcher, please contact me off line and we can maybe set up a demo for you and at the same time, clean up your particles.
If the collected particles are organic, then of course ,this approach most definitely should not be used.
Disclaimer: SPI Supplies manufactures the Plasma Prep II plasma etcher and supplies the carbon discs and the Zerostat antistatic gun.
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 Nov 5 07:35:49 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (pastasalda-at-aol.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, November 5, 2003 at 05:52:22 ---------------------------------------------------------------------------
Email: pastasalda-at-aol.com Name: john downes
Organization: J.G.M.Downes. consultants.
Title-Subject: [Microscopy] MListserver:
Question: Retired SEM required for good home in the UK. To be used as part of a training project.
Thanks for your word of warning, especially since I was not explicit about the situation in my facility. Due in part from previous discussion from this group, I "designed in" a concern about LN2. My current utility room is 8 x 15 ft in size. It has one door that opens directly to the corridor--so tanks can be delivered without zig-zagging through the lab-- and another door to the TEM room. All users are instructed to never dispense LN2 with the doors shut.
Your paranoia (caution) is well-placed!
Don On Wed, 5 Nov 2003, Arthur Day wrote:
} } We also have placed the LN2 tank in this room. If you have } } the space and flexibility, I fully recommend it. } } } } Always dreaming up the worst scenarios and I apologise in advance for } that. However: } } How big and how well ventilated should a separate "utility room" be } if an LN2 tank is being stored in there? How big is the LN2 tank? } Also, where are the LN2 transfers from that larger vessel into } smaller hand-held dewars being done? If you are storing a large LN2 } tank that lets out a "lot" of nitrogen into that room over time, and } it is a room that is normally closed up nearly all the time because } "nobody goes in there that much", then could there be a slight risk } of too much air (oxygen) getting displaced? Maybe this could be a } potential problem if there is no formal ventilation in that room, or } if the room is only serviced as part of a closed A/C system. } } There have been a few postings on this list in past years relating } tales of air displacement by evaporating LN2 in confined spaces. } Perhaps this is also something that should be considered when } thinking about where to put stuff associated with running EMs these } days. } } Just my paranoid thought for the day! } } } } } } } Arthur Day, Electron Microscope Unit Phone: 61-2-9717-3457 } Ansto Materials Division Fax: 61-2-9543-7179 } PMB 1, Menai (Sydney), NSW, 2234 Email: ard-at-ansto.gov.au } Australia www: http://www.ansto.gov.au/ } }
______________________________________________________________________ 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 Nov 5 08:47:41 2003
Angie, A few years ago we had a similar situation with an old JEOL TEM. Over several weeks, I dismantaled the column and separated out the various metals for recycling. The take-home-message was that I was able to realize about $35.00 for working through about 10 lunch hours plus carrying MANY pounds of metal to a recycling facility on a Saturday morning. Our facilities department took care of the rest of the scope so I do not know what they needed to do for disposal. Hoping someone else has a better solution, Pat
Patricia Stranen Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu ============================ } Below is the result of your feedback form (NJZFM-ultra-55). } It was submitted by (anjeanette.ormonde-at-unilever.com) from } http://microscopy.com/MicroscopyListserver/MLFormMail.html on } Tuesday, November 4, 2003 at 14:44:59 } ------------------------------------------------------------------- } Email: anjeanette.ormonde-at-unilever.com } Name: Anjeanette Ormonde } Organization: Unilever HPC } Title-Subject: [Microscopy] MListserver: Dismantling TEM } } Question: We have a Philips 400 TEM that we are no longer going to } be keeping. Right now it is under vacuum and everything is working } well. However, we do not use the system enough to justify the costs } of keeping it (we let the service contract expire in Dec. 2002 but } even beyond that depreciation costs come out of our budget). } Anyhow, I was not able to find anyone who wanted the system (free to } a good home!) so I am (sadly) faced with dismantling and discarding } the system. However, I have no experience with this. Are there } components we can separate out and either keep or give to others? } What happens with unwanted pieces? Can most things go out with } regular garbage? Can anything be recycled? Thanks in advance for } any advice or tips you can offer. } } Sincerly, } Angie Ormonde
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 5 09:01:01 2003
Adrian, If your chem lab has hexane, you might try that, or just use gasoline which is mostly a mix of alkanes of varying chain length. Of course, all appropriate precautions need to be taken for a very dense, highly flammable vapor.
Ken Converse owner Quality Images third party SEM service Delta, PA
ady jenkinson wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 5 09:20:00 2003
I'm a 20 year veteran of a clinical (and some research) EM laboratory. I believe that I am underpaid and under appreciated and intend to do something about it. However, I need some hard facts to support my theory. I'm one of those people that don't make a fuss about things and just take what I'm given...the problem is that I think that I've been taken advantage of. Up to 5 years ago, I was the 'junior tech'. When the 'senior tech' left, I took over all the responsibilities and haven't been given any compensation (title or pay wise). To make matters worse, I believe that I'm very underpaid in general for being the only person who can do what I do. I just have to make administration aware of this.
My question is this...would anybody be willing to share what a 20 year clinical specialist would be making in your area??? I'd be happy to tell you what I am making but don't want to publicly announce it in this forum...too many people would laugh at how low it is!
Another question is how are clinical EM technologists classified in your hospital/facility? I do it all...the technical preparation and then investigate the case by taking (hopefully!) representative pictures and then presenting this to the pathologist. Investigating a case seems to put a tech into a different category but how does administration in your facility look at this...any comments?
Any help/ideas are sincerely appreciated.
Unappreciated in Nebraska, Karen Bovard
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 5 10:15:17 2003
Why do you want to put the particles away from the tape ?
Pull the tape away from the paper, cut the zone where the particles are, et fix it on a SEM stub, with carbon paint, carbon tape, or what you normally use. Than sputter some carbon on it, to avoid charges on the tape. It works !
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
} } } ------------------------------------------------------------------------------ } 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 } ------------------------------------------------------------------------------- } } All } Some kind soul has presented me with samples of debris } which are very small (I would guesstimate {10 } microns). These are stuck to magic tape and then onto } paper. What I now need to do to prepare for EDX } analysis is to get them out of the adhesive of the } tape... } On ordinary tape, the adhesive, I have found, can be } dissolved using chloroform (acetone just makes a } sticky goo). The Scotch Magic Tape appears different } though and I am left with a sticky goo again. } Question: Anyone any idea how I can dissolve this } adhesive? } } Regards } Adrian Jenkinson } } __________________________________ } Do you Yahoo!? } Protect your identity with Yahoo! Mail AddressGuard } http://antispam.yahoo.com/whatsnewfree }
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 5 12:28:21 2003
Have you looked into donating it to a local college, community college, high school or museum? I've usually found that somewhere can be found, often a college that someone in the company attended in the past. A growing destination (more so for SEMs than TEMs, because of their more intuitive imaging and interface) is a local high school. Also gives you or others a chance to donate a little of your time integrating it into a curriculum, instructing the teachers and perhaps demonstrating to the kids. Especially nice if you happen to have a student there.
Allen R. Sampson Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174
On Tuesday, November 04, 2003 6:20 PM, by way of Nestor J. Zaluzec [SMTP:anjeanette.ormonde-at-unilever.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 } ------------------------------------------------------------------------ ------- } } Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (anjeanette.ormonde-at-unilever.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, November 4, 2003 at 14:44:59 } --------------------------------------------------------------------------- } } Email: anjeanette.ormonde-at-unilever.com } Name: Anjeanette Ormonde } } Organization: Unilever HPC } } Title-Subject: [Microscopy] MListserver: Dismantling TEM } } Question: We have a Philips 400 TEM that we are no longer going to be keeping. Right now it is under vacuum and everything is working well. However, we do not use the system enough to justify the costs of keeping it (we let the service contract expire in Dec. 2002 but even beyond that depreciation costs come out of our budget). Anyhow, I was not able to find anyone who wanted the system (free to a good home!) so I am (sadly) faced with dismantling and discarding the system. However, I have no experience with this. Are there components we can separate out and either keep or give to others? What happens with unwanted pieces? Can most things go out with regular garbage? Can anything be recycled? Thanks in advance for any advice or tips you can offer. } } Sincerly, } Angie Ormonde } } --------------------------------------------------------------------------- } }
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 5 13:23:07 2003
On Wednesday, November 5, 2003, at 07:14 AM, Pat Connelly wrote:
Dear Angie and Pat, Our old TEM in Albany NY found a good home in one of the local high schools. We thought that it was a much better solution than recycling it as scrap. Unilever could even write off the donation, so the company would likely be happy to cooperate with any arrangement you might make. We found a school in the Albany area (population ~ 0.5M), so it is likely you can find one near you. Good luck. Yours, Bill Tivol 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
} Angie, } A few years ago we had a similar situation with an old JEOL TEM. Over } several weeks, I dismantaled the column and separated out the various } metals for recycling. The take-home-message was that I was able to } realize about $35.00 for working through about 10 lunch hours plus } carrying MANY pounds of metal to a recycling facility on a Saturday } morning. Our facilities department took care of the rest of the scope } so I do not know what they needed to do for disposal. } Hoping someone else has a better solution, } Pat } } Patricia Stranen Connelly } The University of Pennsylvania } Department of Biology } Philadelphia, PA 19104-6018 } 215-898-7145 } psconnel-at-sas.upenn.edu } ============================ } } Below is the result of your feedback form (NJZFM-ultra-55). } } It was submitted by (anjeanette.ormonde-at-unilever.com) from } } http://microscopy.com/MicroscopyListserver/MLFormMail.html on } } Tuesday, November 4, 2003 at 14:44:59 } } ------------------------------------------------------------------- } } Email: anjeanette.ormonde-at-unilever.com } } Name: Anjeanette Ormonde } } Organization: Unilever HPC } } Title-Subject: [Microscopy] MListserver: Dismantling TEM } } } } Question: We have a Philips 400 TEM that we are no longer going to be } } keeping. Right now it is under vacuum and everything is working } } well. However, we do not use the system enough to justify the costs } } of keeping it (we let the service contract expire in Dec. 2002 but } } even beyond that depreciation costs come out of our budget). Anyhow, } } I was not able to find anyone who wanted the system (free to a good } } home!) so I am (sadly) faced with dismantling and discarding the } } system. However, I have no experience with this. Are there } } components we can separate out and either keep or give to others? } } What happens with unwanted pieces? Can most things go out with } } regular garbage? Can anything be recycled? Thanks in advance for } } any advice or tips you can offer. } } } } Sincerly, } } Angie Ormonde
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 5 18:11:40 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mccaulak-at-wfu.edu) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, November 5, 2003 at 13:46:45 ---------------------------------------------------------------------------
Email: mccaulak-at-wfu.edu Name: Anita McCauley
Organization: Wake Forest University
Title-Subject: [Microscopy] lymphocyte preparation for the SEM
Question: I am in need of information on how to prepare CD4 and CD8 lymphocytes for viewing on the SEM. I would appreciate any protocols, references, tricks of the trade, etc...
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dwaraka-at-casimir.ece.uic.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, November 5, 2003 at 16:12:42 ---------------------------------------------------------------------------
This question is regarding a Digital Instruments Nanoscope III AFM/STM. I wanted to know if it was ok to leave the piezo scanner plugged in "before" switching on/off the Nanoscope controller. Initially I was of the opinion that the scanner had to be unplugged before switching on/off the controller.
I would like to transfer the license of MacTempas, software, and hardware key with a reasonable price. The transfer does not violate the license agreement. Please contact me at: hkonishi-at-unm.edu
I also handle Japanese university or Japanese NSF funds (KOHI BARAI in Japanese)through third company.
Thank you,
Hiromi Konishi, Ph.D Dept. of Earth and Planetary Sciences The University of New Mexico
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 6 02:10:25 2003
We are using the Bal-Tec CDP 030 critical point dryer. A collegue, who's working on another type of machine just told me that the specimen chamber of the CPD 030 is filling up too slowly. This may lead the the material being exposed to surface tension forces, hence deforming the specimen. Is it normal that the specimen chamber on the CPD 030 is filling up slowly or could it be that the problem lies in the fact that our liquid CO2 cylinder is placed in a safety cabinet and that the tubes leading to the CPD are too narrow (outside diameter 3mm)?
Dear folks liquid nitrogen always scares me a little. You need to be very careful about room size, ventilation and potential displaced air.
Three things you should consider above all else are: 1. you can't smell or in any other way detect oxygen depletion. 2. you will be comatose long before all of the oxygen is depleted and sleepy and less aware long before that. 3. the amount of air displaced by one litre of liquid nitrogen is 830 litres.
Add a few more points to this. If you're unconscious in a room and someone tries to rescue you, they will almost certainly suffer the same fate and so on. Further if cleaners, maintenance or security ever go into that room how will they be protected?
The worst possible scenario should always be considered in a risk assessment over something as insidious as this. How big a volume of liquid nitrogen and how much air will it displace. Remember dewars can lose their insulation (we've had 2 or 3 go over 20 years) so that would release all of the nitrogen in a few hours.
You must have warning signs and should seriously consider an oxygen depletion monitor in any area where there is a serious risk. If your room is not ventilated then should you store liquid nitrogen in there under any circumstances?
There have been several well publicised tragedies in the past.
Malcolm
PS paranoia is only when you think everyone's out to get you - in this case 'paranoids' have a much better long term chance of survival.
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: Donald Lovett {lovett-at-TCNJ.EDU}
LAST CALL FOR REGISTRANTS
31st Scottish Microscpy Symposium Wednesday 12th November 2003
University of Dundee, West Park Centre 319 Perth Rd, Dundee, Scotland
Registration deadline extended to 7/11/03 - see http://www.abdn.ac.uk/emunit/smg2003.htm for contact details.
Scientific Programme and Trade Exhibition
Programme
09.30-10.0 Registration and coffee
Morning Session
Chair William Maxwell
10.00-10.45 Photochemical internalisation: From microscopy towards treatment Kristian Berg, Norwegian Radium Hospital, Oslo
10.45- 11.15 Overview and Recent Advances in SPM for the Life Sciences' Mike Conroy, Applications Scientist, Veeco Instruments Ltd, Cambridge, UK
11.15-12.00 High pressure freezing - the ultimate approach for immunolabelling? Paul Monaghan, Institute of Animal Health
12.00-13.45 Lunch, Trade Exhibition and Posters
Afternoon Session
Chair / Laurence Tetley
13.45-14.30. A comparison of cryo-SEM with other preparation techniques for the study of plant cell walls. Kim Findlay, John Innes Centre, Norwich 14.30-14.45 Tobacco mosaic virus-movement protein function investigated by FRAP Kathryn M Wright, Cell-cell communications programme, Scottish Crop Research Institute, Invergowrie, Dundee, DD2 5DA. 14.45-15.00 In situ hybridisation to study gene expression patterns in nematode infected root tissue. Jane Wishart, Alison Paterson, Hui Liu, Vivian Blok. Scottish Crop Research Institute, Dundee Refreshments/Trade exhibition
Chair / Eric Lachowski
15.30-16.15 Mapping surface properties at micro/nanometre levels - by a multi-function Tribological Probe Microscope (TPM) Xianping Liu, School of Engineering, University of Warwick
16.15-16.30 Prize presentations and End of Meeting announcements
This Symposium is CPD accredited by the IBMS (1 Credit).
Exhibitors :
Veeco Instruments Ltd Microscopy Supplies and Consultants Ltd, Improvisionl Leica Microsystems (UK) Ltd, Oxford Instruments Analytical TAAB Laboratories Equipment Ltd Royal Microscopical Society ISS Group Services Ltd Agar Scientific Ltd Hitachi High-Technologies Corporation Princeton Gamma Tech (UK) Ltd Imaging Associates Ltd Emitech Ltd LEO Electron Microscopy Ltd Gatan Olympus Optical Co. (UK) Ltd Quorum Technologies
Dr Laurence Tetley Division of Infection & Immunity, IBLS, Integrated Microscopy Facility Joseph Black Building University of Glasgow Glasgow G12 8QQ
l.tetley-at-bio.gla.ac.uk tel 0141 330 4431 FAX 0141 330 3516
I've looked at neutrophils and macrophages in the SEM using the following protocol: Have the investigator plate the cells onto(round) coverslips that have been treated with poly-l-lysine or some other ECM-type molecule) Wash then fix with buffered glutaraldehyde (2-4% is usual, you can use 1%). They should be well adhered to the glass. After that, you can use a pretty standard SEM prep (post-fix, dehydrate, CPD, sputter). I use round coverslips because they fit into a holder I have for my CPD. I load them into the holder after the glut. fix and then do the rest of the processing. Its easier than having many wells or dishes with coverslips, and it minimizes the chances of having your samples flip over and you don't know which side is up. 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 Thu Nov 6 09:01:23 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (useitzer-at-fz-borstel.de) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, November 6, 2003 at 08:49:52 ---------------------------------------------------------------------------
Email: useitzer-at-fz-borstel.de Name: Ulrike
Organization: Research Center Borstel, Germany
Title-Subject: [Microscopy] MListserver:
Question: Hello! I don't know if this is a stupid question, but here goes: I am interested in imaging DNA/RNA using acridine orange and fluorescence microscopy. Is there any way of fixing and preserving the staining without changing the DNA/RNA signals and is there a possibility to use anti-fading reagents for AO? I would be most grateful for any help! Thank you and good bye Ulrike
Are these cells spread or in suspension? If the former, leave them on their substrate, if the latter, filter them down onto a 0.2 or 0.45 micron membrane filter. The kind with neat, round holes, not a torturous-path filter. Use a syringe filter holder, and keep the fliters in the holder -- use the syringe to change the solutions. Fix in 1 or 1.25% glutaraldehyde + 1% tannic acid (Mallinckrodt 1764 seems to work best) in whatever buffer you're using for the cells *minus* any serum proteins or other peptides. The tannic acid reduces or prevents holes in membranes. Fix for 1 (maybe 2 hrs) hour at room temp or overnight at 4 deg C. OsO4 is usually not needed, although if desired, you can try 1% OsO4 + 1% tannic acid for 1 hr at room temp. Wash in buffer and dehydrate starting with 30% EtOH through 3 X 100% *dry!* EtOH, with 5 to 10 min. steps. Usually 5 minutes for cells. Critical point dry using 4 or 5 5 min. soaks and 2 or 3 min purges (depending on your CPD's specimen holders, and what nooks and crannies they have for trapping EtOH/lqCO2). After the EtOH series, you can also try drying with HMDS (hexamethyldisilizane). There are various recipes for this on the U Florida Tips & Tricks page: http://www.biotech.ufl.edu/EM/tips/index.html I haven't done lymphocytes with HMDS, though, just CPD, so I can't say how well HMDS works for this purpose.
Phil
} Email: mccaulak-at-wfu.edu } Name: Anita McCauley } } Organization: Wake Forest University } } Title-Subject: [Microscopy] lymphocyte preparation for the SEM } } Question: I am in need of information on how to prepare CD4 and CD8 } lymphocytes for viewing on the SEM. I would appreciate any } protocols, references, tricks of the trade, etc... } } Thanks } } ---------------------------------------------------------------------------
-- 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 Nov 6 09:22:08 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (couryhouse-at-aol.com) from on Thursday, November 6, 2003 at 09:07:36 ---------------------------------------------------------------------------
Email: couryhouse-at-aol.com Name: ed sharpe archivist for smecc
Organization: southwest museum of engineering, communications and computation
Title-Subject: [Microscopy] Museum needs several micro manipulators
Question: For several projects and demonstrations we need several micro manipulators Please respond off list and let us know what you can share....
Ed Sharpe archivist for SMECC
see the museum's web site at www.smecc.org or when in Arizona stop in and visit....
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (couryhouse-at-aol.com) from on Thursday, November 6, 2003 at 09:09:44 ---------------------------------------------------------------------------
Email: couryhouse-at-aol.com Name: ed sharpe archivist for smecc
Organization: southwest museum of engineering, communications and computation
Title-Subject: [Microscopy] Museum needs RCA EMT electron microscope parts and docs....
Question: The museum has an RCA EMT electron microscope we are restoring.... need some parts and more documentation
to see what one of these looks like go to the museum's page at... http://www.smecc.org/rca_emt_tabletop.htm
There are some nice images you can download of the brochure, the large ones are really large so a hi speed connection is preferable!
Identification of Man-Made Textile Fibers using PLM.
December 5, 2003
class=Section2} Step by Step Procedure for the Identification of Man-Made Textile Fibers using PLM.
The day will begin with a brief overview of a personalized approach to modern textile fiber identification using PLM and then move on to a full workshop in which the participants will be given unknown fabric samples throughout the day to identify. As a beginning review, each participant, together with the group, will look at the natural fibers such as cotton,linen, silk (cultivated and tussah), and wool to be sure we can eliminate them from our unknown pool. The workshop will then concentrate on the most common man-made fiber types found in contemporary and historic textiles, such as viscose, cuprammonium rayon, acetate, triacetate, polyester, acrylic, nylon, olefins, and even elastomerics, Beginning with sample taking and mounting techniques, this day will cover the most effective techniques to use in the identification of unknown man-made fibers: refractive index, the 530nm red tint plate, and the Michel Levy chart. Although the day is about how far one can go with fiber identification using PLM alone, a discussion of corroborative solvent and burning tests will also be covered. The day will end with a "final exam" for which each student will be given a mystery fabric of mixed fiber types to identify. The instruction will be supplemented by printed materials to take home.
The Workshop Instructor is Denyse Montegut, the Chair of the Museum Studies Department in the School of Graduate Studies at the Fashion Institute of Technology where she has taught conservation science courses since 1991. She received her B.A. in art history and mathematics from Brooklyn College, and holds an M.A. in art history and a certificate in conservation from the Institute of Fine Arts, NYU. She is currently a Ph.D. candidate in art conservation research at the University of Delaware. Denyse also has a small private conservation and consulting practice.
WHEN: Friday December 5, 2003, from 10 A.M. to 5 P.M.
Because the fumes of the resin are so objectionable, is it permissible to do the actual embedding step out of the cold chamber (-50) in the fume hood or must the specimens remain at the low temperature suggested for the entire infiltration series? We have figured out the logistics of changing our specimens from lower to higher concentrations of resin while maintaining temperature discipline, but the act of teasing out individual specimens, placing them in embedding molds or gel-caps and filling them with resin seems problematical if some warming isn't tolerated. Any advice or guidance would be much appreciated.
As always, thanks to Nestor for keeping up with us and the web world and thanks to those who share their hard won expertise.
Regards, Bill Sharp
William P. Sharp Arizona State University School of Life Sciences, box 4501 Tempe, AZ 85287-4501 Phone - (480)-965-3210 Fax - (480)-965-6899
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 6 13:21:25 2003
Hi all, I would like to purchase a good modern histology text to use as a general reference for this facility. I would like one that is sufficiently up-to-date to include SEM, TEM and ICC (Light & EM).
I would appreciate recommendations.
Thanks, 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
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 6 15:10:29 2003
A workshop on electron micro-diffraction and direct methods for crystalline structure determination using electron diffraction/imaging will be held at the National Center for Electron Microscopy (NCEM), Lawrence Berkeley National Laboratory, University of California, Berkeley from April 18 through April 23, 2004. For further details, which will be posted as they are finalized, see http://ncem.lbl.gov/workshop.htm.
----------------------------------------------- Laurence Marks Department of Materials Science and Engineering Northwestern University Evanston, IL 60201, USA Tel: (847) 491-3996 Fax: (847) 491-7820 mailto:ldm2-at-risc4.numis.nwu.edu http://www.numis.nwu.edu -----------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 6 16:03:03 2003
If you know the room can have an atmosphere that won't support life you pull some one out by holding your breath and working in short sprints. You should first get some one to back you up and tie a line to your leg so he can drag you out without having to go in himself. And your effort should be to get a rope on the person that is down and get out of the room and drag them out.
Spend some time thinking before you go in that room because as soon as you blood oxygen saturation drops you are through thinking. You can only act on plans made before hand.
With all the safety warriors at every turn if there is a room that has a liquid nitrogen container large enough to pose any risk at all without a oxygen monitor you should roast your safety officer over a bed of low coals for ignoring real risks and enforcing petty ones.
Gordon Gordon Couger red-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. : : Dear folks : liquid nitrogen always scares me a little. You need to be very careful about room size, ventilation and potential displaced air. : : Three things you should consider above all else are: : 1. you can't smell or in any other way detect oxygen depletion. : 2. you will be comatose long before all of the oxygen is depleted and sleepy and less aware long before that. : 3. the amount of air displaced by one litre of liquid nitrogen is 830 litres. : : Add a few more points to this. If you're unconscious in a room and someone tries to rescue you, they will almost certainly suffer the same fate and so on. Further if cleaners, maintenance or security ever go into that room how will they be protected? : : The worst possible scenario should always be considered in a risk assessment over something as insidious as this. How big a volume of liquid nitrogen and how much air will it displace. Remember dewars can lose their insulation (we've had 2 or 3 go over 20 years) so that would release all of the nitrogen in a few hours. : : You must have warning signs and should seriously consider an oxygen depletion monitor in any area where there is a serious risk. If your room is not ventilated then should you store liquid nitrogen in there under any circumstances? : : There have been several well publicised tragedies in the past. : : Malcolm : : PS paranoia is only when you think everyone's out to get you - in this case 'paranoids' have a much better long term chance of survival. : : 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 -----
: } : } : } Thanks for your word of warning, especially since I was not : } explicit about : } the situation in my facility. Due in part from previous : } discussion from : } this group, I "designed in" a concern about LN2. My current : } utility room : } is 8 x 15 ft in size. It has one door that opens directly to the : } corridor--so tanks can be delivered without zig-zagging through : } the lab-- : } and another door to the TEM room. All users are instructed to never : } dispense LN2 with the doors shut. : } : } Your paranoia (caution) is well-placed! : } : } Don : } On Wed, 5 Nov 2003, Arthur Day wrote: : } : } } } We also have placed the LN2 tank in this room. If you have : } } } the space and flexibility, I fully recommend it. : } } } : } } : } } Always dreaming up the worst scenarios and I apologise in : } advance for : } } that. However: : } } : } } How big and how well ventilated should a separate "utility room" be : } } if an LN2 tank is being stored in there? How big is the LN2 tank? : } } Also, where are the LN2 transfers from that larger vessel into : } } smaller hand-held dewars being done? If you are storing a large LN2 : } } tank that lets out a "lot" of nitrogen into that room over time, and : } } it is a room that is normally closed up nearly all the time because : } } "nobody goes in there that much", then could there be a slight risk : } } of too much air (oxygen) getting displaced? Maybe this could be a : } } potential problem if there is no formal ventilation in that : } room, or : } } if the room is only serviced as part of a closed A/C system. : } } : } } There have been a few postings on this list in past years relating : } } tales of air displacement by evaporating LN2 in confined spaces. : } } Perhaps this is also something that should be considered when : } } thinking about where to put stuff associated with running EMs these : } } days. : } } : } } Just my paranoid thought for the day! : } } : } } : } } : } } : } } : } } : } } Arthur Day, Electron Microscope Unit Phone: 61-2-9717-3457 : } } Ansto Materials Division Fax: 61-2-9543-7179 : } } PMB 1, Menai (Sydney), NSW, 2234 Email: : } ard-at-ansto.gov.au} Australia : } www: http://www.ansto.gov.au/ : } } : } } : } : } ____________________________________________________________________ __ : } 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 Thu Nov 6 16:41:42 2003
Fawcett's The Cell has the best B&W TEM and SEM photos around but little text. The old Bloom and Fawcett "A Textbood of Histology" (later editions were only with Fawcett as the author but still carried Bloom and Fawcett above the title) has the most authoritative text and excellent B&W images but lack the pizazz of modern histology texts. Most of my students prefer Wheater's "Functional Histology" which is a short but probably sufficient text with lots of excellent LM's and a more limited number of EM's. Berman's Color Atlas of Basic Histology is also quite good. It has minimal text but large LM images. Good luck. Tom
At 02:30 PM 11/6/2003 -0500, you wrote:
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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 (cachuca-at-hotmail.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Thursday, November 6, 2003 at 16:49:34 ---------------------------------------------------------------------------
Organization: UNAM, (National Autonomous University of Mexico)
Education: Undergraduate College
Location: Mexico City, Mexico
Question: I would like to know several questions: 1)Are vacuoles exclusive of plant cells? 2)Are lisosomes exclusive of animal cells? 3)If there exists a plant or/and animal cell model (accepted in the scientific community)which shows the proportionate sizes of the ultrastructures in each prototype and is it based on reconstructions from electron microscope observations. If so, I would like to know where it has been published. Thank you very much
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (ldemp-at-mse.ufl.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, November 7, 2003 at 07:05:34 ---------------------------------------------------------------------------
Organization: Major Analytical Instrumentation Center, University of Florida
Title-Subject: [Microscopy] Florida Society for Microscopy Symposium
Question: Florida Chapter of the AVS Science and Technology Society Florida Society for Microscopy Florida Section of the American Ceramic Society 2004 Annual Joint Symposium
March 8-12, 2004 University of Central Florida Student Union Orlando, Florida
Web: www.flavs.org
You are invited to attend the 2004 Annual Joint Symposium of the Florida Chapter of the AVS Science and Technology Society (FL AVS), the Florida Society for Microscopy (FSM) and the Florida Section of the American Ceramic Society (Fl ACerS). This symposium is sponsored by the Florida Chapter of the AVS Science and Technology Society and will be held jointly with the Florida Microscopy Society and the Florida Section of the American Ceramic Society at the University of Central Florida, Student Union Building in Orlando from March 8-12, 2004. The scientific program consists of a keynote lecture, technical sessions, a tutorial session on MEMS, a poster session and competition for students, an equipment exhibit, an AVS national short course program, a Latin-American school of electron microscopy (LASEM) program, and a science workshop for high and middle school teachers. A reception will be held on Monday evening, March 8, for all symposium participants and attendees. There is no registration fee for either the symposium or equipment exhibit but pre-registration is encouraged. On-line registration will be available by the end of December on the Florida AVS website.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (baskin-at-bio.umass.edu) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, November 7, 2003 at 12:05:59 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] postdocs in microscopy
Question: Microscopists, Please bring this to the attention of qualified candidates...
Postdoctoral positions in Computational Biology
Two postdoctoral positions are available to join a research project in the quantification of deformable motion in biology, with emphasis on cell motility. The positions are supported by an NIH-funded collaboration between Tobias I. Baskin (a biologist at Umass Amherst) and K. Palaniappan (a computer scientist at University of Missouri, Columbia). One position will be at Columbia, the other at Amherst. Baskin and Palaniappan have developed new software for quantifying the spatial distribution of velocity within a growing plant organ (a root). The software is called RootflowRT and the biological application is described by van der Weele et al (2003 Plant Physiology, 32:1138-1148). The software implements a novel algorithm for quantifying deformable motion that combines structure-tensor and robust-matching approaches. The current project is to enhance and validate RootFlowRT, apply software engineering principles to the current code base, explore new computational algorithms, and extend the robust-tensor approach to other kinds of biological objects, in particular motile animal cells and embryos. The position at Missouri is primarily computational (with bio-imaging opportunities possible based on candidate interest). The successful candidate for this position will have a strong background in any area of image processing. The position at Amherst will involve imaging different kinds of biological object as well as modifying the software. The successful candidate will have experience with both imaging in biology as well as computer programming, preferably in the area of image processing.
Those interested in the computational position (Missouri) should contact Dr Palaniappan (email: palani-at-cecs.missouri.edu), and can find further information from his web page: http://www.cs.missouri.edu/facultypages/palani.html and the multimedia communications lab page: http://meru.cecs.missouri.edu/ .
Those interested in the biological computation position (Amherst) should contact Dr Baskin (email: baskin-at-bio.umass.edu), and can find further information from his web page: http://www.bio.umass.edu/biology/baskin/ and the page for RootflowRT http://meru.rnet.missouri.edu/mvl/bio_motion.
We encourage applications from anyone regardless of skin color, religion, sex, sexual orientation, or nationality.
Thank you all very much for the procedures. I was going to wait to report on the quality until the negatives I've got are scanned and formatted for a web page, but I've got the negatives scanned and no time to put the following in a web page yet.
On the 28th (maybe the date even makes a difference with formvar films :wink:), I set up to do a few different methods. And now having done my different trials, I can say what I think works for me.
} From the suggestions here, I set about to do two different methods, 'breath' and 'steam' and of course a 'control.' The common procedure for all: -0.5% Formvar (too thick, commenting further down) in Ethylene Dichloride -Fisher Superfrost Slides, fresh out of the box, and wiped vigorously with dry fisher lens paper. -'Cleaned' slide was dipped into a dip-miser containing the fomvar solution -It was pulled out, letting the excess wash down (this process is all about watching and observing the formation of the film - too slow, too fast - lumpy draining...) -Treatment: ----Breath: As one suggestion said fog it up but DO NOT breath in, and the proximity of the wet ED coated slide to my mouth was disconcerting but I carried on in the name of 'education' (and with great caution NOT to breath in). So I fogged it up... the formvar on the slide immeadiately turned a bit white, foggy. The slide was then allowed to dry completely
----Steam: I put an Erlenmeyer flask on a hot plate and had it boiling prior to starting any of the previous steps. The still wet slide was wafted above the mouth of the flask in the steam... (I made 6 slides for this test). The slides were affected very differently with different times in the steam, and what I found later was that different areas of the same slide had different results. After steaming the slides were allowed to dry.
----Control: Dip and dry.
-Collection of the film was standard: cut sides of film with a razor blade, float off in a 9" straight sided bowl in clean water at a 45 degree angle, breath on it before floating and after each cut on the sides, but only after eating a taco salad and spinning around three times on the chair chanting "formvar is my friend..." (sorry, couldn't help myself).
-Grids were placed dull side down on the film -Grids and film were picked up with parafilm covered slides (my preferred method - but after the next step and reading Andrew's procedure I might chance it for making lacey films). -Grids were removed and placed on filter paper in a 110 degree C oven for 3 minutes (or not) -Imaged on the TEM
Results: The control grids were not heated and yielded a thick film (with one or two small holes-it was an older batch of formvar). But in general a standard film. The Breath grids were covered with lacey patterns but very few true holes, most of the lace was just thicker areas of formvar. The Steam grids, both heated and un-heated, were lacey. Hole size was a bit variable, but they were true holes, and from what I can tell very suitably for our purposes here. Many of the holes had stragler filaments of formvar, and heating them for the full 12 minutes at 110 didn't appreciably change the holes or the filaments... The next batch I will add the ED dish vapor variable as outlined by Andrew Blackwood, to see if that changes anything. (images will be posted)
To be honest I was rather surprised how well they turned out. And as luck might have it they might not be the same the next time. But as specific as this is and as much sample as can be loaded on a single grid, the time spent checking the grids at each stage is well spent. One observation I have is that the film didn't look like the clean 'control' film. I might find that as I move on to the sample loading phase that the holes in the lace are too big.
Another observation noted was the thickness of the film and the supports. They should be acceptable, esp since they are for looking at samples suspended over the holes, but the film is thick, and the next batch I try (to verify and refine my technique) I will do with both a 0.5% and a 0.25% formvar/ED solution, and evaluate that variable. The un-coated film is, as to be expected, increasingly unstable as the intensity of the beam goes up.
After carbon coating even the thinnest filaments are stable. Even at crossover. And the films are clean and free from evaporated debris as reported in one or two papers.
Finally (11/7/03) I managed to get a sample of the carbon "nanotube" (no nanotubes - just an amorphous carbon sample unfortunately) and was very successful in imaging the sample.
I also had the great pleasure to switch from the old formula 4489 film to the new stuff in process of collecting before carbon and after carbon coating. This film is aweful - simply horrible. Even constant hand agitation yielded poor results (better than just agitating ever 30 seconds as we had with the old formula). This is incredibly frustrating (the new film that is).
And of course as with a few things in the EM world, explaining and duplicating the lacey carbon procedure in a different place under a different phase of the moon in a different drainage basin might be impossible... or more seriously as Larry Allard said "there is an art associated with each step."
(and I will get the procedure and images on the cmich facility web page soon)
Geoff Williams Microscopy Facility Supervisor
Central Michigan University Biology Department Microscopy Facility http://www.cst.cmich.edu/centers/microscopy/
} -----Original Message----- } From: Blackwood, Andrew [mailto:ablackwood-at-2spi.com] } Sent: Thursday, October 30, 2003 7:43 AM } To: microscopy-at-sparc5.microscopy.com } Subject: [Microscopy] Making Lacey Carbon Grids }
} 29 October 2003 } } Geoff Williams asked how to make lacey carbon coated grids. There may be } many methods; a number of methods may be found in textbooks. Here's how we } do it. The problem with instructions like these is that the devil is in } the } details, and times, temperatures and concentrations all vary with the } season.
snip
} And it helps to have been doing the process for several decades. This is a } skilled art, and the only substitute for going through the learning curve } yourself is to ask someone who has "been there" before you to make the } grids } you need. } } Andy } } Andrew W. Blackwood, Ph.D. } Vice President, Technical } SPI Supplies } P.O. Box 656 } West Chester, PA 19381-0656 } Ph: 1 610 436 5400 X108 } FAX: 1 610 436 5755 } e-mail: ablackwood-at-2spi.com } WWW: http://www.2spi.com }
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 7 14:35:21 2003
I don't think there is one book that covers everything you need but there are several good texts available. "Histology, a text and atlas", 4th edition, by Ross, Kaye and Pawlina is just out, looks very nice. I liked the third edition muchly. One of the few books with a good balance of text and atlas. "Wheater's Functional Histology", 4th edition, by Young and Heath, is good but is more atlas than text. Earlier editions are also good. "Color Textbook of Histology", 2nd edition, by Gartner and Hiatt is good but is lacking in the atlas department. There is a seperate atlas by the same authors. For EM only (mostly TEM) "Cell and Tissue Ultrastructure" by Cross and Mercer is excellent.
Geoff
Debby Sherman 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 Fri Nov 7 15:45:49 2003
Dear Listers, We would like to set up reflected-light Kohler illumination on an optical bench using a half-silvered mirror to show our students how an epi-fluorescence microscope works. We've managed the transmitted-light set-up and it was very time-consuming.
If any of you have been down this path before and can help us save time from re-inventing the wheel, by giving us an idea of distances, which lens focal lengths and what diameter lenses work, I'd really appreciate hearing from you.
Cheers, Jeremy
Jeremy Sanderson.
________________________________________________________________________ Want to chat instantly with your online friends? Get the FREE Yahoo! Messenger http://mail.messenger.yahoo.co.uk
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 7 16:00:56 2003
I've looked at a lot of TB & BCG bacteria. Could you tell me more about this:
"Rife was the first worker known to have isolated and photographed the tuberculosis virus."
-mc
} Hi Bruce: } If you want a "sane discussion" about this man's work how about } providing some real evidence of what he has accomplished, that is, where } his work has been published. Claims made on a website are just that, } claims made on a website. When I went on to the homepage and read about } how the FDA and the AMA are conspiring to keep his work out of the } medical manistream I got a bit skeptical, but that's just me. } Geoff } } Bruce Grosso wrote: } } } Check this out. } } } } http://www.rife.de/mscope/mscope2.htm } } } } Let's have a sane discussion about why all you geniuses are not following in } } his footsteps.
____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 7 16:28:12 2003
Geoff, You left out a critical detail. Do you spin clockwise or counter-clockwise? Thanks for the advice. Kim
On 7-Nov-03, at 11:44 AM, Geoff Williams wrote:
} -Collection of the film was standard: cut sides of film with a razor } blade, float off in a 9" straight sided bowl in clean water at a 45 } degree angle, breath on it before floating and after each cut on the } sides, but only after eating a taco salad and spinning around three } times on the chair chanting "formvar is my friend..." }
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 7 18:36:52 2003
Of all the replies I received you are the first person who understands that Rife's work has less to do with Microbiology or cancer research as it has to do with the general search for the "Mortal Oscillatory Rate" of virtually ALL pathogens.
He didn't set out to "cure cancer" that was never his goal. His goal was to see what if any effects his wavelength theory applied to cell structure. Not unlike breaking a crystal glass with sound wave, in fact exactly the same principle. That was it in a nut shell. BUT, in order to do that he had to first invent the device to View the living specimen.
Then; POOF! The universal microscope was invented and then continually upgraded for several generations. Now everyone focuses on the biological properties of Rife's work. Everyone is barking up the wrong tree. Everyone tears Rife's work apart based on his work as a biologist, which he was not, the biologists came to him after the fact.
I believe he shattered the shattered the glass and now everyone is looking at his flawed biological science when that isn't even the subject and never has been.
You are right! Someone needs to explore Rife's work from Rife's perspective. Someone needs to prove you can shatter the glass, not the biology, the biology will come all by itself and without any help. It will just happen in cadence with the real science.
Thank you for being so insightful. That makes you one of those geniuses I am always talking about on the list.
Thanks you also helped me encapsulate Rife into less than a paragraph for future discussions should the opportunity ever arise again. Who knows maybe one day Rife will be right up there where he belongs with Tesla and others.
From MicroscopyL-request-at-ns.microscopy.com Sat Nov 8 01:17:34 2003
I know a lot of people in this forum are biting their cheeks, or simply deleting any messages in this particular thread. But, in my usual fashion, I'm more than happy to jump into an area that I confess to have little knowledge of.
I would just like to add my two cents worth, since this captured my attention - and my interest comes down to one simple fact. If you (Bruce) want to argue some miracle cure, you can find a more suitable soapbox than the microscopy listserver.
There are plenty of venues out there for this avenue of thought. Whether or not Rife managed to produce an optical instrument capable of resolutions exceeding current instruments is a valid topic here. I, personally, haven't seen any recent serious work proposing that, but would love to see a peer reviewed research paper on it. I suspect that there are more than a few individuals on this listserver that would love to find this all true, as the commercial aspects alone would be nirvana, not to mention the accolades to the researcher. However, this stinks of an early 1900's Pons and Fleischmann - bad and incomplete science brought to media attention to aggrandize the researcher and attract money to an endeavor that was never really viable.
I'm sure that you will want to respond to this, so while you're at it, you might want to provide us with the particular credentials that entitle you to rant to such a wide variety of professionals who have spent their careers in microscopy. This isn't a particularly large field, and the real experts in optical microscopy are getting fewer, but we all share an interest in the sub-atomic particle interactions at the heart of this problem.
Your desire to see Rife elevated to the heights of scientific minds such as Tesla tends to put your rantings in perspective. Aside from his influence in Westinghouse's decision to promote the use of AC currents in electrical power distribution, I really can't think of a single contribution of Tesla that has resulted, after all of the ensuing years, in any practical application. Of course, I'm a Troglodyte, and don't have idea of the vision of the future that you have. But please don't clog the microscopy listserver to enlighten me - email me directly. I'll be more than happy to engage you for a while.
Allen R. Sampson Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174
On Friday, November 07, 2003 8:45 PM, Bruce Grosso [SMTP:bgrosso-at-AssetRecovery.Net] 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 } ------------------------------------------------------------------------ ------- } } Michael: } } Of all the replies I received you are the first person who understands that } Rife's work has less to do with Microbiology or cancer research as it has to } do with the general search for the "Mortal Oscillatory Rate" of virtually } ALL pathogens. } } He didn't set out to "cure cancer" that was never his goal. His goal was to } see what if any effects his wavelength theory applied to cell structure. Not } unlike breaking a crystal glass with sound wave, in fact exactly the same } principle. That was it in a nut shell. BUT, in order to do that he had to } first invent the device to View the living specimen. } } Then; POOF! The universal microscope was invented and then continually } upgraded for several generations. Now everyone focuses on the biological } properties of Rife's work. Everyone is barking up the wrong tree. Everyone } tears Rife's work apart based on his work as a biologist, which he was not, } the biologists came to him after the fact. } } I believe he shattered the shattered the glass and now everyone is looking } at his flawed biological science when that isn't even the subject and never } has been. } } You are right! Someone needs to explore Rife's work from Rife's perspective. } Someone needs to prove you can shatter the glass, not the biology, the } biology will come all by itself and without any help. It will just happen } in cadence } with the real science. } } Thank you for being so insightful. That makes you one of those geniuses I } am always talking about on the list. } } Thanks you also helped me encapsulate Rife into less than a paragraph for } future discussions should the opportunity ever arise again. Who knows maybe } one day Rife will be right up there where he belongs with Tesla and others. } } }
From MicroscopyL-request-at-ns.microscopy.com Sat Nov 8 08:16:53 2003
Allen, Are you going to let the rest of us know if there is some REAL information out there? Given that resolution is given as magnification and TB is a virus, I have my doubts that there will be anything to report. It's been a long time since I was a microbiology major, but I've heard of E. coli, can anyone confirm the existence of B. coli?
If it's on a web site, it must be true........
Ken Converse owner Quality Images third party SEM service Delta, PA
Allen Sampson wrote:
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From MicroscopyL-request-at-ns.microscopy.com Sat Nov 8 09:22:37 2003
I've lately been seeing an increase in REJECTED mail questions from subscribers. As you know I have tightened the noose on the Junk/Spam mail filters significantly in the last few months.
If you get a rejection message obviously the first thing to do is:
1.) READ the instructions and try to correct the problem yourself or 2.) follow the instructions exactly and contact me
One of the more significant changes is that the subscription database is compared to the senders Email address. If the FROM address does not match EXACTLY your subscription address then a REJECTION message is sent to the senders FROM address. This in principle, allows the sender to fix the problem or contact me.
The most common problem I am seeing now is that many subscribers have over the years changed their Email addresses subtly.
For example, if you originally subscribed as
Nestor-at-Microscopy.Com
and have changed/modified your sending address in your Email program to
Nestor.Zaluzec-at-Microscopy.com
and tried to post, it will be flagged as suspect spam. The logic here is that SPAMMERS get hold of addresses, spoof them by sometimes making small changes and then use them. In order to control this I check both the subscription and senders address, if there is a discrepancy it is flagged and a REJECTION message sent. Remember, the software is literal and if there is the slightest difference, a rejection message will be generated.
If you are one of those subscribers that encounters this problem you can recognize it immediately by simply looking at the detailed text which is provided by the REJECTION software.
The message which the listserver sends in this case clearly says"
"Suspect or Possibly an Unregistered User"
if you get one of these check your subscription address and compare it to your sending (FROM) address.
You might ask, but I still get mail using my original address, what is the difference. This can happen if your MAIL Administrator has setup a forwarding/redirection service to allow mail addressed to your "original" Email address to be sent to your new address. You have 2 options to cure this problem.
1.) change your FROM address back to the original subscription address or
2.) unsubcribe your old address and subscribe your new one.
Option #2. will create less work for me in the long run. If you cannot change your sending address contact me off-line and we can work out a solution on a case by case basis. There is an exceptions list, but it has to be manually edited, and I obviously don't want to do this for the thousands of users on this list. Most of you don't have this problem so this issue is a mute point.
A second variation of this problem is that your posting and subscription addresses don't match, but you never receive a rejection message.
You can be confident that a rejection message is being sent out , but the problem is that your ISP may be rejecting the warning message.
This is a classic CATCH 22 situation. I can't contact you because your SERVER is rejecting the error messages and you can't post because there is a problem but your not seeing the reason.
If you attempt to post and after ~ 24 hours you don't see your posting, try using the WWW based form at:
http://www.microscopy.com/MicroscopyListserver
This form allows anyone to post to the server, however, each message is first read and approved. This is to allow anyone to post (even non-subscribers) and provides a mechanism (albeit an organic computer) to verify the validity of the post. Obviously there is a longer delay in this process, because of the necessary intervention.
I will also take this opportunity to remind you all that all EMail with attachments is rejected. There are no exceptions.
The most common problem here is with MS Outlook type Email programs adding text formatting to your message (color, bold, italics, fonts) which is encoded as HTML and added as a hidden attachment. You must send your message as PLAIN/ASCII text. Most Email programs will allow you to set this option in the Settings/Preferences area. If you cannot then you can also use the WWW based form if your Email program is unintentionally adding attachments to your message. The WWW form does not create any attachments, nor will it permit you to add any.
Finally, as long as I have your attention,
I'll also remind/ask you all to use the WWW based form to subscribe and unsubscribe at http://www.microscopy.com/MicroscopyListerserver
This form minimizes the work I have to do, however, I do personally monitor and execute the update scripts daily just to make sure no spammers sneak into the list.
Cheers
Nestor Your Friendly Neighborhood SysOp
BTW, if your curious there are over 100+ junk mail messages/day now being rejected by the SPAM filters. So although all of this is a hassel, it is IMHO worth the effort, otherwise this list would have stopped being useful long ago.
From MicroscopyL-request-at-ns.microscopy.com Sat Nov 8 09:43:43 2003
I sent this already to Allen but in light of Ken's comment I feel I am obligated to share these few thoughts with everyone.
First off I am not on a Rife soapbox. Second, I had already dropped the discussion idea as out of the dozens of mostly derisive replies only a few, not always agreeable people, kept their heads and dissected what they knew about Rife's' work in intelligent and respectful tones, Thirdly, I am not an educated person, I have always meant "You geniuses" in the most flattering tones, or at least tried to on account of the fact that to me you all "ARE" geniuses compared to me.
What I know about Rife's work began after reading, and then on several occasions, rereading the book by Barry Lynes.
Several lengthily phone conversations with Lynes and two personal meetings with one of Rife's living coworkers in his home in San Diego in the late 1980's with him at that time in his 80's., Touring his garage with him, seeing what I saw in the way of documents and equipment and listening to what he had to say intensely. It wasn't until those two meetings that it dawned on me that Rife actually did produce the microscope, I viewed a few of the remaining original 35mm through the lens photos with him, and bits and pieces of what could only be described as detailed documentation of something I could not make heads nor tails of, for want of the academic background to comprehend, but could not imagine that this man, who stood before me, so late in years was clinging to a lie so close to the end of his life.
It also occurred to me begged the question: What is so unfathomable about the theory of cellular wall destruction by waveform? It is already a widely accepted fact that sound wave can kill you given the right frequencies.
Rife never contended that, what he set out to do was to isolate each individual cells particular, individual "Mortal Oscillatory Rate" and for each individual pathogen, be it bacterial or viral.
What is so hard to accept? That he concocted this microscope and lied relentlessly about the results he was reporting, given the man's credentials and well documented personal achievements?
I am more prone to believe his lab was vandalized and parts of the microscope were pilfered than to think someone of his already renowned stature fabricated his claim of magnifications for which he was well qualified to create according to academic records as outlined in the book, duly footnoted, cross referenced and included in the bibliography.
If you haven't read the book there is no point in defending or being offensive towards something you haven't read. I lost my copies (I had six altogether) years and years ago.
And yes! Now after all this, I do plan on acquiring another copy so I can brush up on my rebuttals to repudiations.
Lastly is Barry Lynes himself who I have had the pleasure of several lengthily telephone conversations with and he in no way appears to be the type who would idealize something or someone and instead seems to be a thorough and competent, objective reporter, who has been caught up in a shit storm of his own making but quite unintentionally.
He has no illusions of Rife. Does not idolize Rife and reported the facts as he acquired them with no slant or favoritism in most depictions of his dissertation on Rife's life and work. He respects Rife but does not idolize him.
So what am I then therefore to think? Uneducated by and large except in the most cursory way towards each of these sciences, yet tantalized by the sheer scope and magnitude of the drug companies strangle hold on an industry who sheer might and force of power is nothing less than awesome, combined with the BILLIONS of dollars in cancer research monies and yet the plague still grows exponentially with no end in sight.
For myself I chose to believe in the adage. "Keep It Simple Stupid" which is what I think Rife did. He never set out to cure cancer. That was never his intent. He set out to shatter the crystal glass and I am convinced to whatever degree; he did it, he actually did it. Even with the possibility that his achievements could possibly have been over stated in the broadest sense by others I am convinced he succeeded to some extent and THAT is what needs to be recreated.
But alas, it is so much easier to criticize than scrutinize, or deride than to investigate. It is so much simpler to hold the party line and shout "Impossible" instead of conceding that vandalism and arson destroyed the man's work in an age where backup was not as easy as the click of a mouse. Hell there weren't even copy machines back then.
Given Rife's penchant for detailed reporting of the simplest and most benign movement of the scopes adjustments a (few surviving pages of which I saw myself but made no sense to me) just the simple replication of his work by a clerk would have been nothing less than a monumental undertaking in it's own right.
You say you have some knowledge of this man's work, But do you have any knowledge of the man? Have you ever tried to get into his head? Read about his life from an objective reported viewpoint and try to see into his mind and what he was thinking all those long 10, 15, 20 or 30 hour sessions sitting at the scope recording his every turn of the dial, pencil in hand every time he touched a screw or adjusted a component, this way and that. And then documenting that failure after another and another until he found what he was looking for, all or in part.
Like a movie is an encapsulation of a book, it doesn't all fit into 1 hour and 10 minutes, a book is an encapsulation of a life, it doesn't all fit into a few hundred pages.
No I think you geniuses are the ones who need to get real, not me, not Rife. If you don't think he did it. Prove it. Recreate his every footstep and then tell me he was wrong. Especially when the underlying theory is just all too simple in it's concept and all too complicated in it's implementation.
I can say, yes here is how we can get to the moon and back, but what it took to actually do that was a huge undertaking. I believe so it is too with Rife's work, his theory and I believe he did the work and took on the monumental task and left his foot print in the sand. Now someone needs to go back and with today's technology retrace his footsteps and prove he was wrong or right.
That is what gets me. No one is willing to prove him wrong, yet everyone wants to accuse him of something I just don't think the man was capable of.
I got into his head and I wish I had your education and an extra 20 years left because I would be willing then to undertake the task. I am just a salesman. You guys are the geniuses.
I am convinced someone in your fields of expertise will take up the mantle one day. It is just a matter of time.
Cheers and God Bless
Bruce
Bruce Grosso www.AssetRecovery.Net, Inc. 67 County Rd. 274 Iuka, MS 38852 662-423-1757 Ph. 775-213-9028 FAX bgrosso-at-AssetRecovery.Net Equipment Recovery Specialists ----- Original Message ----- } From: "qualityimages" {qualityimages-at-netrax.net} To: {ars-at-sem.com} ; "Microscopy" {Microscopy-at-Sparc5.Microscopy.Com} Sent: Saturday, November 08, 2003 6:24 AM
Colleagues.....
I see no discussion of microscopy in this thread and note that it is beginning to breakdown into off-topic issues unrelated to the subject of the Listserver.
Further postings should be taken off-line, between those that are interested in continuing.
Nestor Your Friendly Neighborhood SysOp
From MicroscopyL-request-at-ns.microscopy.com Sun Nov 9 09:24:10 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dougbaldwin-at-mindspring.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Saturday, November 8, 2003 at 22:54:42 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] Macro-Nikkor Lenses and Nikon Multiphot Macroscope
Question: Dear Listers,
I'm shooting the circuitry of microchips such as a Pentium-type processor. The chip faces range in size from 1mm to 20mm. My 6 Mb Fuji S2 digital camera with 60mm Micro-Nikkor lens don't resolve enough detail and resolution on the larger chip faces (for one important client) and the large chips are too big for a microscope to capture in one shot.
I'm thinking of shooting them with a MACRO-Nikkor, either the 65mm f4.5 or 120mm f6.3 very high resolution lens on a 4x5" camera or a Nikon Multiphot MACROscope which was specifically designed for macro photography in the 1x-40x range.
The Mulitphot system was last made in the mid 1980's by Nikon. They were not standard camera store items so they don't show up in current used camera inventories. I can't find either the lenses or the Multiphot for sale anywhere.
I'm hoping one of you microscopists out there might have one or both of the lenses or even the whole Microphot system (including the 19mm f2.8 and 35mm f4.5 Macro lenses) sitting in a closet unused and lonely, just waiting to be put back into service shooting hi-tech photos of microchips in the 21st century.
There are other macro lenses that might work as well. Those would be the Zeiss Luminars or the Leitz Photars. Does anyone have any of these lenses?
I'm thinking of purchasing an older Nikon Optiphot microscope with coaxial lighting (necessary for reflective microchip faces) to shoot the small 1-5mm chips. My budget is limited and thought this might be an appropriate microscope for this part of the project. Any feedback?
I would appreciate any thoughts on this subject and especially a Nikon Multiphot system that might be for sale.
You may well have already rejected this on some other grounds, but why don't you get someone with a SEM to image them for you?
Or use a lower-magnification optical microscope.
cheers
rtch
Date sent: Sun, 9 Nov 2003 09:35:52 -0600 To: microscopy-at-ns.microscopy.com } From: dougbaldwin-at-mindspring.com (by way of MicroscopyListserver)
This is done either of two ways, depending on the final resolution desired. The simplest and the method with least overall resolution is a stereo zoom.
The highest quality method is to take multiple shots of small portions of the die and stitch them together. This method can produce up to 60"x60" full color 300dpi prints. Or, they can be PDF for on-line viewing. This method uses a metallurgical microscope with motorized stage.
I don't think your main problem is resolution, per se. Rather, it is the field of view that you can obtain with a corresponding ability to resolve whatever degree you need.
gary g.
} ------------------------------------------------------------------------------ } 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 (dougbaldwin-at-mindspring.com) from } http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on } Saturday, November 8, 2003 at 22:54:42 } --------------------------------------------------------------------------- } Email: dougbaldwin-at-mindspring.com Name: Doug Baldwin Title-Subject: [Microscopy] } Macro-Nikkor Lenses and Nikon Multiphot Macroscope Question: } Dear Listers, I'm shooting the circuitry of microchips such as a } Pentium-type processor. The chip faces range in size from 1mm to 20mm. My } 6 Mb Fuji S2 digital camera with 60mm Micro-Nikkor lens don't resolve } enough detail and resolution on the larger chip faces (for one important } client) and the large chips are too big for a microscope to capture in one } shot. I'm thinking of shooting them with a MACRO-Nikkor, either the 65mm } f4.5 or 120mm f6.3 very high resolution lens on a 4x5" camera or a Nikon } Multiphot MACROscope which was specifically designed for macro photography } in the 1x-40x range. The Mulitphot system was last made in the mid 1980's } by Nikon. They were not standard camera store items so they don't show up } in current used camera inventories. I can't find either the lenses or the } Multiphot for sale anywhere. I'm hoping one of you microscopists out there } might have one or both of the lenses or even the whole Microphot system } (including the 19mm f2.8 and 35mm f4.5 Macro lenses) sitting in a closet } unused and lonely, just waiting to be put back into service shooting } hi-tech photos of microchips in the 21st century. There are other macro } lenses that might work as well. Those would be the Zeiss Luminars or the } Leitz Photars. Does anyone have any of these lenses? I'm thinking of } purchasing an older Nikon Optiphot microscope with coaxial lighting } (necessary for reflective microchip faces) to shoot the small 1-5mm chips. } My budget is limited and thought this might be an appropriate microscope } for this part of the project. Any feedback? I would appreciate any } thoughts on this subject and especially a Nikon Multiphot system that } might be for sale. Thanks in advance. Doug Baldwin Baldwin Photography } Scottsdale, Arizona dougbaldwin-at-mindspring.com } ---------------------------------------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 10 17:34:59 2003
This is kind of a fishing expedition. We are thinking of adding a 120KV cryoTEM to our lab and would like to get any helpful hints from more experienced users.
I know practically nothing about this area so anything would be helpful to bring me up to speed. Not only instrument selection questions, but things like service, parts, etc.
One issue that has come up, in addition to the choice of instrument, is the question of cameras. I have read lots about the problems with the new formulations of TEM film and can't avoid noticing the rise in popularity of digital cameras. Anyone have experience that would help guide us in the right direction? Film or digital, 1K, 2K, or 4K? Same questions about service, availability, parts, etc.
Any comments would be welcome.
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 Mon Nov 10 22:19:50 2003
Hi all, Does anyone have a method to jazz up the uranyl acetate followed by lead citrate post-staining procedure? I am working with some tissue that just doesn't seem to want to stain...it looks very dull in the scope. I'm using a saturated aqueous UA then Reynold's lead citrate. Should I try UA in 50% ethanol rather than aqueous? Any suggestions would be greatly appreciated, Beth -- ********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
Phone - (706) 542-1790 & FAX - (706) 542-1805
"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 need to modify or replace our standard fixation protocol to handle pedicels of legume pods for preparation of wax sections for light microscopy. These are about 1 mm in diameter and combine woody (phloem fibers and xylem) and soft tissues (cortex, phloem, cambium, pith) and the tissues readily distort or tear during sectioning. Currently we use an age old protocol consisting of standard fixation in FAA or parafomaldehyde, then ethanol series dehydration through TBA and into Paraplast. Clearly this is not good enough. Any suggestions for an improved protocol that would produce undamaged sections would be greatly appreciated.
Greg
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 04:13:00 2003
On Mon, 10 Nov 2003 23:30:09 -0500 Beth Richardson {beth-at-plantbio.uga.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 } ------------------------------------------------------------------------------- } } Hi all, } Does anyone have a method to jazz up the uranyl acetate followed by } lead citrate post-staining procedure? } I am working with some tissue that just doesn't seem to want to } stain...it looks very dull in the scope. } I'm using a saturated aqueous UA then Reynold's lead citrate. } Should I try UA in 50% ethanol rather than aqueous? } Any suggestions would be greatly appreciated, } Beth } -- } ********************************************************************** } Beth Richardson } EM Lab Coordinator } Plant Biology Department } University of Georgia } Athens, GA 30602-7271 } } Phone - (706) 542-1790 & FAX - (706) 542-1805 } } "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) } } *************************************************************************** } } } } 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"
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 04:53:09 2003
Dear Beth! You will find very good results with UA 2% in methanol (100º) , then the usual lead citrate. (8 minutes for UA, whash in 70º,50º and 30º methanol then double destilated water and them , prior wash dd water, the lead citrate(10 minutes), sometime there are some variation related with the hardness and the tipe of the resin used for inclusion. We get very good results in plant tissues with the above technique. Francisco
Prof. Francisco Freire FRMS Head Service Electron Microscopy Service University of Las Palmas de Gran Canaria Fac. Cs. Salud P.O. Box 550 Las Palmas Canary Islands Spain
Beth Richardson 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, } Does anyone have a method to jazz up the uranyl acetate followed by } lead citrate post-staining procedure? } I am working with some tissue that just doesn't seem to want to } stain...it looks very dull in the scope. } I'm using a saturated aqueous UA then Reynold's lead citrate. } Should I try UA in 50% ethanol rather than aqueous? } Any suggestions would be greatly appreciated, } Beth
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 06:40:08 2003
Years ago, we used uranyl acetate dissolved in anhydrous methanol. As I remember, you can dissolve something like 25% of the UA. We would dip the grids in the the methanol solution for a few seconds and then rinse in methanol. The only drawback was that it seemed to etch the plastic and make the sections weak. The stain was a bit grainy as well. However, in emergencies, it might be useful.
Date sent: Mon, 10 Nov 2003 23:30:09 -0500 To: microscopy-at-msa.microscopy.com } From: Beth Richardson {beth-at-plantbio.uga.edu}
Beth, I use 70% ethanol with UA and it appears to yield better contrast than the aqueous solution. Good luck, Mary Gail Engle
------------------------------------------------------------------------------ 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, Does anyone have a method to jazz up the uranyl acetate followed by lead citrate post-staining procedure? I am working with some tissue that just doesn't seem to want to stain...it looks very dull in the scope. I'm using a saturated aqueous UA then Reynold's lead citrate. Should I try UA in 50% ethanol rather than aqueous? Any suggestions would be greatly appreciated, Beth -- ********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
Phone - (706) 542-1790 & FAX - (706) 542-1805
"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)
Looking at getting a color laser printer for general quick and dirty printing. Particularly looking for feed back from anyone one either the Minolta QMS Magicolor 3100 or the Lexmark C752n. We have excellent experience with our Lexmark B&W printers. And I like the example prints from the Miniolta I pick up at M&M (Thank you Vitally). Now I'm look for more opinions Good the bad and the ugly.
Yes, we have a number of ink jet printers, excellent quality but slow, looking for some thing faster, basically to serve for walk away prints as we do more and more digital.
Thank you!
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 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 Tue Nov 11 08:00:10 2003
Jon It is perfectly understandable that you should want a digital camera on your new TEM. Anyone purchasing today would want one for a variety of reasons, but I am alarmed that you should be driven to this choice by scare stories about film quality. In my experience these are completely unjustified, and I worry that this sort of of talk will unnecessarily accelerate the demise of EM film. Despite enormous advances in digital camera performance the film image still sets the standard to beat for sheer image quality. Kodak electron image films are every bit as good as they have ever been. They are a complete piece of cake to process, and it mystifies me that anyone with darkroom experience could have serious difficulty with this.The process is quick, easy and in our hands completely reliable. The most difficult bit is to find a rack that holds the film securely. After that the film practically processes itself, no voodoo required*. Best wishes with your purchase Chris
p.s. I have to concede that the results are improved if the D19 is matured six months buried in the gizzard of a ritually-slaughtered chicken. Anyway it's a good excuse for a lab party.
On 10 Nov 03, at 15:41, Jon Krupp 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 } -------------------------------------------------------------------- } -- --------- } } Greetings: } } This is kind of a fishing expedition. We are thinking of adding a } 120KV cryoTEM to our lab and would like to get any helpful hints } from more experienced users. } } I know practically nothing about this area so anything would be } helpful to bring me up to speed. Not only instrument selection } questions, but things like service, parts, etc. } } One issue that has come up, in addition to the choice of instrument, } is the question of cameras. I have read lots about the problems with } the new formulations of TEM film and can't avoid noticing the rise } in popularity of digital cameras. Anyone have experience that would } help guide us in the right direction? Film or digital, 1K, 2K, or } 4K? Same questions about service, availability, parts, etc. } } Any comments would be welcome. } } Thanks } } Jonathan Krupp } Microscopy & Imaging Lab } University of California } Santa Cruz, CA 95064 } (831) 459-2477 } jmkrupp-at-cats.ucsc.edu } } }
========================================== Dr. Chris Jeffree University of Edinburgh BIOSEM - Biological Sciences Electron Microscope Facility Institute of Cell and Molecular Biology Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 (0) 131 650 5554 FAX. #44 (0) 131 650 5392 Mobile 07710 585 401 email c.jeffree-at-ed.ac.uk =========================================
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 08:02:16 2003
I have a user with gold-coated samples after doing ion probe analyses, and now he needs to do more electron microprobe work, both imaging and quantitative analyses. The gold coats (I'm not certain about thickness) need to be removed. We cannot, however, polish the gold off because he needs to preserve the pits left from the ion probe. As a result, we are looking for chemical means to remove the gold coats. Any suggestions are welcome (particularly those known to work and with details about concentrations, temperatures, etc.).
Thanks, Ellery
--------------- Ellery E. Frahm Research Scientist/Manager Electron Microprobe Laboratory Department of Geology & Geophysics University of Minnesota - Twin Cities Lab Website: http://probelab.geo.umn.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 09:12:02 2003
I have been using the Minolta QMS Magicolor 3100 for about 1 year, and am extremely pleased with it. Good image quality (both resolution and color fidelity), fast results, low cost. The dye-sub printer is now rarely used and the ink-jet has been completely retired.
John Russ
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 09:58:20 2003
You didn't mention the tissue type nor the resin, but if its Spurr's epoxy, that can make staining of embedded tissues more dificult, especially with Reynold's lead, I think. Or maybe the tissue type just doesn't pick up much stain.
I don't usually use Reynold's stain here, but prefer to use the Sato triple lead stain (lead citrate, lead nitrate, lead acetate), which I've been using for many years. The way I make it up, its VERY stable and gives excellent staining, usually with no lead ppt contamination, tho if its older than 2 months, I may use a 0.2 micron filter as a precautionary measure. A 100 ml batch typically lasts 3-4 months. If you want the protocol for making Sato lead, lemme know off-line.
Before the Sato lead, I stain with 3% aqueous UA for about 20 minutes, room temp. But you may need to cut the UA with methanol and/or elevate staining temperature as others have suggested on this thread for your special case.
Good luck, of course!
Gib
} Hi all, } Does anyone have a method to jazz up the uranyl acetate followed by } lead citrate post-staining procedure? } I am working with some tissue that just doesn't seem to want to } stain...it looks very dull in the scope. } I'm using a saturated aqueous UA then Reynold's lead citrate. } Should I try UA in 50% ethanol rather than aqueous? } Any suggestions would be greatly appreciated, } Beth
-- 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
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 10:42:55 2003
Richard, We are getting very satisfactory results from our recently purchased HP 5500 color laser. It was purchased with image printing in mind but also has to provide color printing for the rest of the facility. Turned out to be a very good compromise and is reasonably quick. Sorry I cannot compare it to the others you mentioned, but thought you like another data point.
No $ connection or interest, just another user.
Kevin Battjes Impact Analytical Voice 989-832-5555, ext 556 Michigan Molecular Institute Fax 989-832-5560 1910 W. St Andrews Road e-mail: battjes-at-mmi.org Midland MI 48640 battjes-at-impactanalytical.com
Visit us at PittCon, March 8 - 11, 2004, booth 2306
-----Original Message----- } From: Richard Edelmann [mailto:edelmare-at-MUOHIO.EDU] Sent: Tuesday, November 11, 2003 8:56 AM To: microscopy-at-MSA.Microscopy.com
Looking at getting a color laser printer for general quick and dirty printing. Particularly looking for feed back from anyone one either the Minolta QMS Magicolor 3100 or the Lexmark C752n. We have excellent experience with our Lexmark B&W printers. And I like the example prints from the Miniolta I
pick up at M&M (Thank you Vitally). Now I'm look for more opinions Good the
bad and the ugly.
Yes, we have a number of ink jet printers, excellent quality but slow, looking for some thing faster, basically to serve for walk away prints as we do more and more digital.
Thank you!
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 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 Tue Nov 11 10:55:41 2003
Beth, If you are using an epon type embedding, then yes the saturated UA in 50% ethanol would be a great help. Some cautions in working with it (besides the radioactive chemical safety ones) are: 1. to really keep it in the dark, even while in the staining dish 2. make it in small batches (5 ml in a small vial works well) because it looses its "umph" after just a few days 3. no need to filter the stain if you let it settle for about 15 minutes after shaking and the stain is taken gently from near the top of the vial so as not to disturb the bottom crystals.
A second place for lack of contrast in staining is to leave the lead stain on too long. The metal will be leached back into the solution.
The samples I work up are usually block stained in 0.5-1.0% aqueous UA in the refrigerator overnight after fixation. My section staining proceedure is to place a dry grid onto a drop of UA stain in a petri dish on parafilm for 8 to 10 min. The lid of the dish is covered with cardboard and opaque tape. Water rinse several times and put the grid into a large drop of water also on parafilm in another dish and leave the grid there until all grids are washed. Transfer right out of the water drop into a drop of lead stain in a third covered dish for about 1-2 min. then wash well before drying it down.
Although I have used Reynold's Lead stain in the past, the lead stain that I am using now is Lead Citrate as described by Aly Famy in Proceedings of the 25th Annual EMSA Meeting - 1967 - 50ml cooled, boiled, distilled water + 1 dry pellet of sodium hydroxide, after it is disolved add 0.2g (reference states 0.25g) lead citrate. I use a plastic (PP) 50ml centrifuge tube so it can be discarded later and store the staining solution in several syringes stuck into a large stopper to keep out the air and keep it in a cabinet to limit light exposure.
Patricia Stranen Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu } ---------------------------------------------------------------------- } The Microscopy ListServer --Sponsor:The Microscopy Society of America } } Does anyone have a method to jazz up the uranyl acetate followed by } lead citrate post-staining procedure? } I am working with some tissue that just doesn't seem to want to stain... } it looks very dull in the scope. } I'm using a saturated aqueous UA then Reynold's lead citrate. } Should I try UA in 50% ethanol rather than aqueous? } Any suggestions would be greatly appreciated, } } Beth Richardson } EM Lab Coordinator } Plant Biology Department } University of Georgia } Athens, GA 30602-7271
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 11:01:36 2003
i've used ethanolic UA for almost 35 years, with very good results. the procedure calls for saturated UA in 50% ethanol, stir for 5 minutes, stain for 10 minutes in reduced light. depending on how much stain you need at a time, you can make it up in as little as 5ml preps. takes just under 200mg in 5mls to get a saturated preparation. i make it up fresh and filter it through a 0.2micropore filter just before use. keeps ok in glass for several days, i've never tried to keep it longer. it keeps very poorly in disposable syringes, only 2-4 hrs - the rubber on the end of the plunger reacts with the solution.
the reference for the procedure is: Stempak,JG; Ward,RT (1964): An improved staining method for electron microscopy. J. Cell Biol. 22, 697-701.
paul hazelton
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 11:14:24 2003
Well, first off the usual sort of advice would be to extend some of the processing times involved, especially the embedding series of dilute and pure Paraplast. I think we are talking times on the order of days for some of those steps, right? - you are probably doing that already. Then, of course, make sure you have a good sharp knife, and have convinced yourself that the microtome is not the problem.
But if you are going to be doing wax embedding of plant tissues for LM on an ongoing basis, you might want to consider throwing a completely different approach at the problem, which is microwave oven processing. In the MW oven, you can do all the steps from fixation of plant tissues to final infiltration in 100% Paraplast in about 5 hours. A book on microwave processing techniques for both LM and EM is available from Ted Pella,Inc.:
Microwave Techniques and Protocols, A New Book on Microwave Processing Edited by Richard T. Giberson and Richard S. Demaree, Humana Press
Chapter 15 is entitled: Microwave Paraffin Techniques for Botanical Tissues.
See also: Schichnes D, Nemson J, Rusin, SE (1998) Microwave protocols for paraffin microtechnique and in situ localization in plants. Microscopy & Microanalysis, 4:491-496.
Here at the CBS Imaging Center, we have taken these two sources, modified the protocols a bit for various reasons and to fit our MW processor and its accessories. We have processed alfalfa nodules for paraffin sectioning, and barley leaf infected with fungus, leaf with viral infections, other fungal specimens in epoxy resin for TEM. Again, in about 5 hours time, including cured blocks in the eposy resin for TEM case.
That approach won't solve your present problems of the moment, but its an approach well worth looking into for many kinds of biological samlple prep for LM and EM.
Laboratory microwave ovens specially designed for this kind of processing are available from vendors such as Ted Pella., Inc., and Electron Microscopy Sciences, maybe others. By the way, I have no financial interst in these two companies. If you want any more info on MW processing of plant materials, contact me off line.
Good luck!
Gib -- 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
} I need to modify or replace our standard fixation protocol to handle pedicels } of legume pods for preparation of wax sections } for light microscopy. These are about 1 mm in diameter and combine woody } (phloem fibers and xylem) and soft tissues } (cortex, phloem, cambium, pith) and the tissues readily distort or tear during } sectioning. Currently we use an age old protocol } consisting of standard fixation in FAA or parafomaldehyde, then ethanol series } dehydration through TBA and into Paraplast. } Clearly this is not good enough. Any suggestions for an improved protocol that } would produce undamaged sections would } be greatly appreciated. } } Greg
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 12:12:27 2003
Try this article from the Sept.-Oct. 2002 Microscopy Today: Peter Tomic Method for Metallization Stripping of Gold Interconnected Semiconductors Using an Aqueous KI Solution The issue is available on-line as a pdf file at http://www.microscopy-today.com/TableofContentsPDF.html
Phil
} Hello all, } } I have a user with gold-coated samples after doing ion probe analyses, and } now he needs to do more electron microprobe work, both imaging and } quantitative analyses. The gold coats (I'm not certain about thickness) } need to be removed. We cannot, however, polish the gold off because he } needs to preserve the pits left from the ion probe. As a result, we are } looking for chemical means to remove the gold coats. Any suggestions are } welcome (particularly those known to work and with details about } concentrations, temperatures, etc.). } } Thanks, } Ellery } } --------------- } Ellery E. Frahm } Research Scientist/Manager } Electron Microprobe Laboratory } Department of Geology & Geophysics } University of Minnesota - Twin Cities } Lab Website: http://probelab.geo.umn.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 Tue Nov 11 12:35:42 2003
I just wanted to thank everyone who replied - there are too many of you to thank individually. The most popular method for improved staining was: UA in 50% ethanol or methanol. Various times (5-20 min) Add 1 drop of acetic acid/per 10mls of solution. Store in an amber bottle. Stain should last a year or longer. Follow UA staining with Reynolds lead citrate.
I appreciate the help! Thanks again, Beth -- ********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
Phone - (706) 542-1790 & FAX - (706) 542-1805
"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)
The first way is to use a sputter coater in the Argon ETCH mode. Use low power and a vacuum of 50mT. The key to using this method is to not have enough energy to zap the devices yet have enough ionic bombardment to clean off the coating. Gradual etching can be examined for effectiveness by SEM imaging at moderate (10KV) energy for charging. Once you get charging, odds are that the coating is gone. I typically use 50A-80A of Au/Pd or Pt.
The second method is to use a Gatan ion mill with Argon. This too is at low power and 50mT or default. Use 2KEV, 40-50uA and 20 minutes, 10RPM, 10 degree rock. You might try building up to 20 minutes in 5 minute increments. This method will not bash metal but it will affect oxide if done too long or too "hot."
gary g.
At 06:13 AM 11/11/2003, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 18:02:23 2003
I am in the process of surplusing our old TN5500. It is available, either as a complete unit or individual components. We have all original monitors, cables, boards, documentation, software, printer,etc., and even the Tracor Northern Training slide show (a collectors item!). The TN5500 was last used in 1996 and was retired in a completely healthy condition. It has since been used as a table top for our new automation system.
The property is offered for sale "As-Is, Where-Is". The University of Washington makes no guaranty, warranty, or representation expressed or implied as to the condition of the property or its fitness for any use or purpose. Seller confirms that it has clear title to the property. The buyer will be responsible for the cost of packing and transporting the equipment from the UW campus.
Cost is negotiable, but will be real cheap. You must pay for shipping but I can do the packaging if the items are small.
************************************************ ....amphiboles do violence to history... T. Feininger, 2001. (taken out of context) ****************************
Dr. Scott Kuehner kuehner-at-u.washington.edu Dept. of Earth and Space Sciences ph.206-543-8393 Mail Stop 351310 Fax 206-616-6873 The University of Washington Seattle, Washington 98195-1310 ************************************************
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 20:52:21 2003
} Dear list friends: } } I am trying to apply AC current to AFM to do some measurement, if teh AFM we have is DI nanoscope III. We do nothave any conductive module yet. But is that possible we can apply AC to AFM? } } Also how can I obtain more information about biological samples preparation for AFM measurement? } } I do appreciate your kindness. } } Chris } } } Life is like a box of hand grenades, you don't know what will blow you to kingdom come.
===== Life is like a box of hand grenades, you don't know what will blow you to kingdom come.
Mr.Yung-fou Chen E-mail: yung_fou-at-yahoo.com
__________________________________ Do you Yahoo!? Protect your identity with Yahoo! Mail AddressGuard http://antispam.yahoo.com/whatsnewfree
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 11 21:57:25 2003
I have been looking for a printer for publication-quality color figures and high-resolution grayscale images. I have been considering the Magicolor 3100, but was put off by some of the reviews I read -- for example, http://www.pcmag.com/article2/0,4149,908304,00.asp
Michael A. O'Keefe Materials Sciences Division LBNL
Richard Edelmann 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 } ------------------------------------------------------------------------------- } } Looking at getting a color laser printer for general quick and dirty printing. } Particularly looking for feed back from anyone one either the Minolta QMS } Magicolor 3100 or the Lexmark C752n. We have excellent experience with } our Lexmark B&W printers. And I like the example prints from the Miniolta I } pick up at M&M (Thank you Vitally). Now I'm look for more opinions Good the } bad and the ugly. } } Yes, we have a number of ink jet printers, excellent quality but slow, } looking for some thing faster, basically to serve for walk away prints as we do } more and more digital. } } Thank you! } } Richard E. Edelmann, Ph.D. } Electron Microscopy Facility Supervisor } 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 Tue Nov 11 23:29:53 2003
Dear all: An investigator of my facility is interested in looking at the cross-section of a peptide nanotubes on a TEM. These tubes have a diameter of 50-60 nm, and the wall of tube is about 4 nm. We would like to start with conventional aldehyde fixation and epoxy resin embedding, however, the sample is very sensitive to pH, and anything above pH 4 would cause disassembling of the peptide tubes. Can anyone recommend an alternative EM fixative that works well at low pH, and explain how this fixative works? Thank you very much.
Hong
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 02:14:03 2003
There was a thread on this in January 2002 The following are representative of the non-polishing methods suggested. Chris ================================================= We routinely use a potassium iodide mixture off the shelf made by Acton Industries, Pennsylvania, USA, on GaAs semiconductors. It's called "GE-6". I imagine for "gold etch." The pure Au metal is ~ 3.5 uM and is removed cleanly by this material. The only caveat is that it attacks exposed GaAs very quickly.
If anyone needs details, please contact me directly.
Peter Tomic Group Leader Failure Analysis & Analytical Services Anadigics, Inc. Warren, New Jersey ============================== Chris,
The solution is
65g iodine 113g potassium iodide dissolved in 100ml cold water.
Dissolve the KI first for a saturated solution, followed by the iodine!
Happy stripping!!
Best Wishes Barry ************************ Barry Lamb Manager, Materials Evaluation Centre C-MAC Engineering Ltd web: {http://www.cmacengineering.com/} e-mail: blamb-at-har.cmac.com tel: +44 (0)1279 403671 (ESN 742-3671) ============================== Chris,
I've used 1:1 Nitric(70%):HCl mixture for gold coat and gold wire removal from semiconductor samples. Although I don't know the etch rate, it will etch off a moderate gold coat in about 20-30 seconds.
Frank Martini ZiLOG Nampa, Idaho ============================= Chris:
A similar question arose on another listserver, although it was removing a 2 mciron gold metallizton from GaAs. The responses from there were as follows:
Response 1 "Greetings!
It is possible that a cut or buffered aqua regia might do the trick. I have had some success with the recipe below on Au metallization at M2 and above on GaAs dice. The interlayer dielectric was silicon dioxide, so there was not a lot of seepage into the die substrate layer - this, more than the recipe, might have kept the damage down. Assuming SiO2 as the ILD on all layers of your part, this might still work:
50 ml deionized water
30 ml HCl
20 ml HNO3
Swirl or agitate the part in the solution for five minutes. Inspect and repeat once if needed.
One minute rinse in deionized water, followed by 30 seconds rinse in acetone. Dry under heat lamp to minimize surface staining.
As always, try this on a practice part first in case this does not buffer the reaction with GaAs enough.
Good Luck !
Regards,
Carl Nail National Semiconductor Carl.Nail-at-nsc.com"
Response 2 "A solution of potassium iodide and iodine in water is a decent gold etch. I have no idea how it would affect GaAs, but I suspect it would be less destructive than Aqua Regia.
Alan Street alan-at-irsi.com"
I hope this helps!
David ================================ Chris; I have had very good luck using a solution of potassium ferrocyanide. Hope this helps.
Jon McGovern Manager, Microscopy and Imaging Facility University of Calgary =================================== We use sodium cyanide solution to remove gold.
Robert Champaign Raytheon Failure Analysis Lab ===================================
========================================== Dr. Chris Jeffree University of Edinburgh BIOSEM - Biological Sciences Electron Microscope Facility Institute of Cell and Molecular Biology Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 (0) 131 650 5554 FAX. #44 (0) 131 650 5392 Mobile 07710 585 401 email c.jeffree-at-ed.ac.uk =========================================
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 02:52:30 2003
A short paper outlining some chemical methods for dissolving gold can be accessed at http://fbe.erciyes.edu.tr/Turkce/eufbedergisi/ DER99/119-127.pdf =================================================== Gold will dissolve into a solution if the solution has an oxidizer (like nitric acid, H2O2, ozone, sodium bromate, etc.) and a chlorine, bromine, iodine, cyanide, thiourea, thiocyanate, or thiosulfate source. This gives a large number of combinations of substances that will dissolve gold. The most famous is aqua regia (muratic acid and nitric acid). Joshua Gulick http://www.ormus.ws/~pyramid/dissolve_gold.htm =================================================== MacArthur was not in fact the first person to realize that cyanide could dissolve gold. That discovery had been made over one hundred years before, in 1783, by the Swedish chemist Scheele who was the first person to prepare the gas hydrogen cyanide. (The companion story of the history of cyanides will be the subject of next week's article.). By the mid)nineteenth century, solutions of gold in cyanide solutions were being used to electroplate gold on to other metals. This process also is used almost unchanged today. Thus we see that MacArthur was not the first person to observe that gold dissolves in cyanide solutions. His contribution to technology, which was recognized in his patent, was to realize that the chemical reaction could be applied economically to the recovery of gold from low)grade ores. As is often the case with technology, cyanide was being used to dissolve gold long before the scientific details of the process were understood. As already mentioned, electroplating of gold was a commercial process by 1850, and gold extraction from ores using MacArthur's patent began in 1889, yet the chemistry of the process was not completely understood until the 1950's. Michael Faraday (1857) was the first person to recognize that oxygen is needed in order for gold to dissolve in a cyanide solution. Neither oxygen alone nor cyanide alone does the job; both substances are needed. An especially puzzling phenomenon is that extremely low concentrations of cyanide (e.g. 0.01 per cent) are sufficient to dissolve gold, and that the rate of dissolution does not seem to be affected by the concentration of the cyanide solution. Eventually it was shown that the limiting factor under typical conditions is how fast the oxygen of the air dissolves in the cyanide solution. A modern view point of the process is that the gold is corroded in a manner not unlike that of iron rusting. extract from THE SCIENCE CORNER by Nigel Bunce and Jim Hunt College of Physical Science University of Guelph http://helios.physics.uoguelph.ca/summer/scor/articles/scor174.htm =================================================== In 1856, M. Faraday prepared the first colloidal gold dispersion in water. To repeat the experiment the reader might reduce gold chloride in water with sodium citrate (which he did with phosphorous). After a short time, a blue coloration will appear and then a ruby-red gold dispersion. =================================================== finally, if all else fails..... Collect a good amount of boys' urine, let it putrefy for some time, and distill the first and subtle spirit over like brandy. Set the filtrate in digestion for 8 days and distill again as before. Keep the spirit but boil the left-overs of both distillations quite dry in a kettle. Calcine it in a potter's furnace, extract from it its fixed salt with rain water, knead it under potter's clay, and distill it like common spirit of salt. You will obtain a yellow, sharp spirit, rather heavy in weight. Rectify it to remove all phlegma, then pour on it by drops the first- prepared volatile spirit. It will effervesce strongly, so that you will be surprised to find so many opposites together in one subject. A white substance will precipitate. Let it settle, pour the phlegma off from it, dry the rest, put it in a curcurbit and sublimate it with a strong fire. A beautiful bright sublimate will rise into the alembic. Remove it and keep it, as it is good for many things. Take one part of it, add to it 3 parts of spirit of salt, digest this together and distill it. Now you will have a wonderful menstruum for dissolving not only gold but all the other metals and minerals. http://www.levity.com/alchemy/agric_06.html ========================================== Dr. Chris Jeffree University of Edinburgh BIOSEM - Biological Sciences Electron Microscope Facility Institute of Cell and Molecular Biology Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 (0) 131 650 5554 FAX. #44 (0) 131 650 5392 Mobile 07710 585 401 email c.jeffree-at-ed.ac.uk =========================================
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 06:21:30 2003
} I have been using the Minolta QMS Magicolor 3100 for about } 1 year, and am extremely pleased with it. Good image quality } (both resolution and color fidelity), fast results, low cost. } The dye-sub printer is now rarely used and the ink-jet } has been completely retired.
Because I'll soon be looking into a high throughput printer as well, this is good info to have. My only question, in the context of grayscale SEM micrographs, is how well the color lasers print good detail in neutral gray(?) That is, I imagine most of them, if not all, achieve better detail when allowed to dither all colors for achieving gray. Is the result neutral? ... and, does the neutrality hold up over time?
Why do one-step immuno-EM? We routinely do GFP immuno-localization using anti-GFP antibody followed by protein A-gold. We get great results. Directly coupling an antibody to gold is never a good idea as it usually decreases the affinity and the complex is not stable very long. There might be some sources of GFP antibody-gold conjugates out there, but I wouldn't really trust them. We have obtained good results using anti-GFP antibody from Molecular Probes. We purchase our protein A-gold from a lab at the University of Utrecht in the Netherlands, but there are many other sources for this reagent.
Marc
On Tuesday, November 11, 2003, at 04:35 PM, Richard Edelmann 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 } ----------------------------------------------------------------------- } -------- } } } ------- Forwarded message follows ------- } Subject: [Microscopy] gold anti-GFP } Send reply to: kissjz-at-muohio.edu } } From: "JOHN KISS" {kissjz-at-muohio.edu} } Date sent: Fri, 31 Oct 2003 14:23:42 -0500 } } Question for listserv; } } There are lots of papers where people use indirect immunofluroescence } to } localize a GFP fusion protein. Does anyone know of a company that } sells an } anti-GFP antibody directly coupled to collodial gold so one can do a } 1-step } labelling for EM level localization? } } } } John Z. Kiss, Ph.D. } Professor } Dept. Botany, Miami Univ. } Oxford OH 45056, USA } tel./Fax: 513-529-5428/-4243 } http://www.cas.muohio.edu/botany/bot/jzk.html } } } ------- End of forwarded message ------- } }
-- 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 Wed Nov 12 08:35:17 2003
Can you make the fixative (glutaraldehyde) in the same buffer the nanotubes are in now? I think a bigger problem might be getting the nanotubes orientated so you can cut them in cross section.
Geoff
Hong Yi 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 all: } An investigator of my facility is interested in looking at the } cross-section of a peptide nanotubes on a TEM. These tubes have a } diameter of 50-60 nm, and the wall of tube is about 4 nm. We would } like to start with conventional aldehyde fixation and epoxy resin } embedding, however, the sample is very sensitive to pH, and anything } above pH 4 would cause disassembling of the peptide tubes. Can anyone } recommend an alternative EM fixative that works well at low pH, and } explain how this fixative works? Thank you very much. } } Hong } }
-- -- ********************************************** 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 Nov 12 08:44:44 2003
We are running a mostly automated Olympus IX81 with IPLab Spectrum 3.6.1. Fortunately, Scanalytics does provide control plugins for the IX81 (for instance, "Set filter to Dapi"), however, we cannot query the status of the microscope from IPLab. For instance, the software cannot ask, "Which filter is the microscope set at?" and make the answer available to a script. Has anybody solved this problem yet? Thanks.
____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 08:53:11 2003
In a message dated 11/12/03 8:40:16 AM, michael-at-shaffer.net writes:
} Because I'll soon be looking into a high throughput printer as well, this is } good info to have. My only question, in the context of grayscale SEM } micrographs, is how well the color lasers print good detail in neutral } gray(?) That is, I imagine most of them, if not all, achieve better detail } when allowed to dither all colors for achieving gray. Is the result } neutral? ... and, does the neutrality hold up over time?
That's a good question and raises several points worth considering: 1. As far as the neutrality of the laser print, when given a monochrome image the Minolta (and in my experience, all other) laser printers use only the black toner so there is no consumption of the color toners (in the Minolta, the CMYK toners are all separate and you can replace K individually as needed) and of course there is no color whatever in the print, and they are smart enough to print with a monochrome halftone pattern rather than leaving space for the color rosettes that are used for full color printing. I've just printed an 8x10 test image consisting of a grey scale ramp, and it shows no banding or "steps" in the grey scale. I would judge that the lightest 3-4% is pure white (no visible toner on the paper) and the darkest 5% or so is "plugged" (i.e., no white showing at all), which is typical of halftone printing. The default halftone pattern used in the Minolta is apparently about 140 cells per inch (by inspection of the print), but the printer is Postscript controlled so this can be set as desired (frankly, I've just left it with default settings and have no complaints). 2. The laser prints have a very long life (i.e., I haven't seen any evidence yet of any deterioration, regardless of some pretty sloppy storage conditions, exposure to light, etc. The same cannot be said for most ink jet prints. The dye based inks fade badly in strong light or humidity, and the "archival" pigment based inks don't have as good tonal resolution and exhibit strong m etamerism (different colors under different lighting conditions). 3. Neither a color laser printer nor a color ink jet printer is the optimum solution for printing only monochrome images. By using an ink jet mechanism with custom inks having multiple shades of grey, Jon Cone (check out piezography.com) has come up with a monochrome printing system that has more tonal range and control than even traditional darkroom printing. But it is slow, and costly per copy. And there are other monochrome-only solutions that use essentially photographic methods (using laser light to expose the "print" which is then developed). These are also slow and costly per copy. For making display images that would rival Ansel Adams prints when hung on the wall, I would not hesitate to go with Piezography. But for routine output I find the halftone printing from the Minolta to rival the quality of a decent offset press, the kind used to print quality books. It is fast, cheap per copy (after an initial investment of nearly $2k), and runs happily on a network with a mixture of Mac, Unix and Windows computers. 4. I have a set of sample (color, but probably now I will include a few monochrome examples as well) images that I carry around printed on dye sub, ink jet and laser printers. Viewed side by side from a normal viewing difference, there are very few differences. I would judge that the dye sub shows slightly more saturation and somewhat richer colors, but not by much. Unless you hold them up very close to see the individual dots in the ink jet, the halftone pattern in the laser output, and the inherent softness in the dye sub, you would probably not be able to guess which was which. And for some of us older folks, it takes a hand lens to really see those details.
John Russ
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 09:00:20 2003
Dear Listers I've found two ex-Leo/Phillips people founded a company to provide SEM & TEM, LM service and support centre at Queens- New York City, they've started by the freelancing services earlier. They've fixed our facility SEM S-435 and dismantled and moved the TEM-900 to another facility with Gemini filter in a very efficient and cost-effective way. We've always faced problem with the companies in their after sales support and Maintenance contract. Hope their service would be very beneficial for many of our fellows who can get many EM related problems solved by them in a inexpensive way. You can contact them in the e-mail id dc-at-infosoftusa.net Regards } From Dr. John Kurtz's Lab Dave NYU ----- Original Message ----- } From: "Greg Barclay" {gbarclay-at-trinidad.net} To: {} Sent: Tuesday, November 11, 2003 2:24 PM
Michael brought up a good point about printing gray images. Sometimes I need to print both a color image and a gray SEM image on the same page. Gray is the hardest color to reproduce. Nearly every color printer I've used uses color ink to reproduce a gray image on a "mixed image" page, resulting in a particualr hue to the gray image. One exception is the Lexmark Optra SC 1275. It can distinguish between a color image and a gray image on the same page, using color ink for the color image and black ink for the gray image and text, even though all are on the same page.
I can't recommend this particular printer because it has a bad reputation of breaking down a lot, plus color saturation was so-so. But I cherished this abovementioned peculiar feature of the printer, and recommend anybody to look for this feature on their future printer purchase.
Stu Smalinskas Metallurgist SKF USA Plymouth, Michigan (734) 414-6862
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
michael-at-schaffer.net wrote:
Because I'll soon be looking into a high throughput printer as well, this is good info to have. My only question, in the context of grayscale SEM micrographs, is how well the color lasers print good detail in neutral gray(?) That is, I imagine most of them, if not all, achieve better detail when allowed to dither all colors for achieving gray. Is the result neutral? .. and, does the neutrality hold up over time?
} I have been looking for a printer for publication-quality color } figures and high-resolution grayscale images. } I have been considering the Magicolor 3100, but } was put off by some of the reviews I read -- for example, } http://www.pcmag.com/article2/0,4149,908304,00.asp
Interesting reading, and this review certainy contrasts with a similar review of a less expensive Magicolor printer, the Minolta-QMS 2350 EN ... http://www.pcmag.com/article2/0,4149,1217172,00.asp ... which gets kudos instead of kriticism for its "photographic" quality and color saturation. Logical questions not answered in the review would be with respect to the expense of the toner cartridges.
} Richard Edelmann wrote: } } } } Looking at getting a color laser printer for general quick and dirty } printing. } } Particularly looking for feed back from anyone one either the } Minolta QMS } } Magicolor 3100 or the Lexmark C752n. We have excellent experience with } } our Lexmark B&W printers. And I like the example prints from } the Miniolta } I } } pick up at M&M (Thank you Vitally). Now I'm look for more opinions Good } the } } bad and the ugly. } } } } Yes, we have a number of ink jet printers, excellent quality but } slow, } } looking for some thing faster, basically to serve for walk away } prints as } we do } } more and more digital. } } } } Thank you! } } } } Richard E. Edelmann, Ph.D. } } Electron Microscopy Facility Supervisor } } 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 Nov 12 10:21:27 2003
Hi Jonathan We have a Hitachi H-7600 with a Gatan cryo stage and an AMT camera.
We really like the H-7600 and have had very good service from Hitachi. We have adapted the transfer technique for putting grids into the Gatan cryo holder and it is now pretty slick. So we are happy with the cryo holder. My suggestion is from experience that you should try and get a second transfer rod straight away. What I like about the Hitachi-Gatan system is that the liquid nitrogen does not pour out when you put the transfer rod in. This happens with the FEI system but I have no experience with the Joel system.
We have a 1985 Zeiss 10C without digital, just film. Since we got the digital camera, no-one has used the Zeiss for taking pictures. It seems to be used mostly for checking grids and such which makes it extremely useful in that the Hitachi is very often booked when you just want to check your grid! For those odd occasions when someone wants higher resolution, the Hitachi has a film camera too.
We have a 1Kx1K digital camera and it is not enough for an 8x10. A 2Kx2K would now be the basic camera I would want but if I could afford it, I would want the 4Kx4K. One of the nice things about having a digital camera is that you can turn the beam down so much that you cannot see anything on the fluorescent screen, but the camera can pick up the signal and give a good image. This is important if you have a section which blows up just as the beam hits it.
Spare parts to specifically ask for when you are buying are spare Wehnelt assembly, spare objective aperture strip, spare condenser aperture strip, and a box of 10 filaments. These may not seem much but when the filament blows, having a spare system ready to go means the TEM is down only a short time. Same with particulary the objective aperture. It can take a week or more to get a spare but if you have one ready to go, you can replace it in a very short time.
I can't think of anything else at the moment Elaine
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From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 10:39:46 2003
} In a message dated 11/12/03 8:40:16 AM, michael-shaffer.net writes: } } } ..., in the context of grayscale SEM } } micrographs, is how well the color lasers print good detail } } in neutralgray(?) ... } } That's a good question and raises several points worth considering: } 1. As far as the neutrality of the laser print, when } given a monochrome image the Minolta (and in my experience, } all other) laser printers use only the black toner ...
That's certainly different from the way inkjets work. Leastwise, if you send the printer neutral RGB data, they'll use all colors. It's my experience with Epsons, this use of all inks is impossible to control, whereas other printers can be told to print "don't use color". For clarification, can you confirm the QMS printer drivers will discriminate neutrality even if RGB data is sent?
} ... I've just printed an 8x10 test image consisting } of a grey scale ramp, and it shows no banding or } "steps" in the grey scale. I would judge that the } lightest 3-4% is pure white (no visible toner on the paper) } and the darkest 5% or so is "plugged" (i.e., no white } showing at all)
Although Photoshop "printer transfer functions" are implied to work with CMYk EPS files only, I find they will work with RGB data ... and could possibly remedy the loss of detail for either end of the histogram (depending on the printer).
This question is regarding a Digital Instruments Nanoscope III AFM/STM. I wanted to know if it was ok to leave the piezo scanner plugged in "before" switching on/off the Nanoscope controller. Initially I was of the opinion that the scanner had to be unplugged before switching on/off the controller.
Does it really matter
thank you dwarakanath
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 11:10:56 2003
I'm looking for information on any procedure using Davidson's Fixative for eyes for both Light and Electron Microscopy
Any information would be greatly appreciated
Thanks,
Pedro
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From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 11:33:28 2003
Hong, It sounds like this is a sample that would benefit from high pressure or plunge freezing followed by freeze substitution. That way you would minimize collapse and pH would not be a critical factor. Even adding fixative to the freeze substitution solution wouldn't matter since you would not be buffering it.
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
On 11/12/03 12:41 AM, "Hong Yi" {hyi-at-emory.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 } ---------------------------------------------------------------------------- --} - } } Dear all: } An investigator of my facility is interested in looking at the } cross-section of a peptide nanotubes on a TEM. These tubes have a } diameter of 50-60 nm, and the wall of tube is about 4 nm. We would } like to start with conventional aldehyde fixation and epoxy resin } embedding, however, the sample is very sensitive to pH, and anything } above pH 4 would cause disassembling of the peptide tubes. Can anyone } recommend an alternative EM fixative that works well at low pH, and } explain how this fixative works? Thank you very much. } } Hong } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 12:30:47 2003
In a message dated 11/12/03 1:19:42 PM, michael-at-shaffer.net writes:
} That's certainly different from the way inkjets work. Leastwise, if you } send the printer neutral RGB data, they'll use all colors. It's my } experience with Epsons, this use of all inks is impossible to control, } whereas other printers can be told to print "don't use color". For } clarification, can you confirm the QMS printer drivers will discriminate } neutrality even if RGB data is sent?
Yes. Even if the document has black text with both color and monochrome image insets, the printer correctly uses no color toner except in the color image region. This is indeed a much better solution than ink jets, which must use colored inks as well as black in order to get enough paper coverage to generate a "black" appearance. Laser printers are capable of complete area coverage with any of the toner colors, although for printing color images they normally lay down the colors using a halftone grid pattern.
} Although Photoshop "printer transfer functions" are implied to work with } CMYk EPS files only, I find they will work with RGB data ... and could } possibly remedy the loss of detail for either end of the histogram } (depending on the printer).
Setting up a printer curve that skips the very ends is easy, and solves the problem as you note. I printed my test sheet intentionally without that correction. By the way, the printer drivers for the Minolta (and all other laser printers I've worked with) expect an RGB image not CMYK. They perform the conversion themselves. If you do print a CMYK image, it first gets converted back to RGB! The Photoshop CMYK conversion is appropriate for making color separations for offset web press, but is not useful for laser (or ink jet) printers.
Also, to respond to a point raised in a private email, I have no financial interest in Minolta, or in Piezographics. Just a very satisfied user.
John Russ
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 18:26:49 2003
I've been using the HP Color LaserJet 4550N for several years. Excellent results and long life for the toner. It prints with CMYK so greyscale is done with just the black toner. Grey, black and color on one page all render perfectly. Image quality is excellent. The printer is 600dpi and produces outstanding color prints. The newer version is the 5500. With the network option, any computer on the LAN can get to the printer. Using DAVE, a Mac can also print to the color laser using TCP/IP.
toner cartridges cost about $175 each. But they last a long time and only need to be replaced individually when empty.
gary g.
At 08:05 PM 11/11/2003, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 20:36:54 2003
I have used these old protocols with success on woody materials. I am not sure how long you processed your tissues, but plant material always requires greatly extended times. Also we softened the embedded material in Gifford's solution. The difficulty is always trying to get the harder tissue soft enough so it doesn't tear the softer tissue during sectioning.
The details we used are as follows: Fixation for 12 to 48 hours (you might also try Navashin's fixative which contains chromic acid) Infiltrate through TBA series (Johansen series) for 2 hours each step 3 changes of pure TBA over 24 hours 3 changes of Paraplast over 24 hours Embed
Soften the blocks by soaking overnight to 7 days in water at up to 38°C If this doesn't work try 2 days to 2 weeks in Gifford's (room temp in fume hood): 20 ml glacial acetic acid 80 ml 60% ethanol 5 ml glycerine You will have to try the times with your own tissue. If you leave it too long the soft tissue will become macerated. If this fails then you may have to go with a resin embedding.
Good luck,
Kim On 11-Nov-03, at 12:54 AM, Greg Barclay wrote: } } I need to modify or replace our standard fixation protocol to handle } pedicels of legume pods for preparation of wax sections } for light microscopy. These are about 1 mm in diameter and combine } woody (phloem fibers and xylem) and soft tissues } (cortex, phloem, cambium, pith) and the tissues readily distort or } tear during sectioning. Currently we use an age old protocol } consisting of standard fixation in FAA or parafomaldehyde, then } ethanol series dehydration through TBA and into Paraplast. } Clearly this is not good enough. Any suggestions for an improved } protocol that would produce undamaged sections would } be greatly appreciated. } } Greg } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 12 22:13:45 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (steven.young-at-sri.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, November 12, 2003 at 18:39:57 ---------------------------------------------------------------------------
Email: steven.young-at-sri.com Name: Steve Young
Organization: SRI International
Title-Subject: [Microscopy] MListserver:Vintage Cambridge SEM available
Question: We have a Cambridge Mark IIA SEM that was purchased new in 1968 that has been up and running until retired recently. This instrument is 90% vacuum tube electronics and was completely overhauled 10 years ago. It is clean and reasonably reliable. We need to find a good home for it, preferably a museum or a private collector. I can't stand to see it end up in a dumpster. It includes a spare stage and many options including all manuals and documentation. The only cost would be packing and shipping from Menlo Park, CA. 94025
Any other ideas? I've tried several local museums.
The point is that we need to connect Link AN 10000 computer with IBM PC. We tried to use XMODEM program. But it was useless in spite of the program works well with other applications. May be the reason is not up-to-date programs (1986-87). So we need your advice. Thanks. Best regards, Sadova
http://www.ipm.sci-nnov.ru/
mailto:sadova-at-ipm.sci-nnov.ru
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 13 08:23:23 2003
Hi, has anybody longer experience with diamond knives from Drukker (The Netherlands / Europe)? Can You recommend them? Have you ever had re-sharpened a knive? Thanks for a short informal and offline reply to
peter.heimann-at-uni-bielefeld.de
Peter Heimann
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 13 10:37:05 2003
I currently using a Gatan GIF for EELS on a Tecani thermal field emitter. I was told that extraction voltage can affect energy resolution to the spectrometer. If so how may I go about optimizing extraction voltage for optimizing Gif for high resolution PEELS?
Thanks
Thurston Herricks
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 13 10:39:49 2003
This may be a broad inquiry, but at this point my knowledge on the topic is limited and thus my need for help from the group:
Our lab is considering expanding some capabilities, and we are investigating the possibility of establishing a hot-lab for various analyses on low level radioactive samples. Theoretically, this could potentially include a SEM, which leads me to contact everyone.
Since all of my experience is with strictly non-radioactive materials, what are some of the key considerations for such an undertaking? Beyond shielding and safe handling (which other people here will help assure), with specific regards to a SEM, imaging and analytical capabilities, what (if any) are some things to become aware of, investigate, and learn about as we get into this issue?
TIA!
If desired, I can be contacted off list.
Chris Holp FirstEnergy Corp. HolpC-at-firstenergycorp.com 440-604-9704
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From MicroscopyL-request-at-ns.microscopy.com Thu Nov 13 12:14:10 2003
Microscopy Society of America 2004 Undergraduate Research Scholarship Program
With this year's call for applications, the MSA Undergraduate Research Scholarship Program begins its 16th year providing funding for undergraduate research. To date over 81 projects covering a wide range of topics in the physical and biological sciences have received support through this program. Over the years, nearly all of the scholarship recipients have maintained a strong interest in imaging sciences and have gone on to graduate school, professional school, teaching, or industry positions.
The program, which is funded by MSA and by matching funds from MSA Sustaining Members, is able to support over 50% of applicants. The maximal award for the Undergraduate Scholarships is $3,000 and helps to provide student stipends, supply costs, and limited travel expenses associated with the research. Additional support in the form of instrument use time, equipment purchases, etc., is generally provided by the student's supervisor and/or through the sponsoring institution. Abstracts reporting the research results, are prepared by scholarship awardees, and published in "Microscopy and Microanalysis".
The program actively seeks sources of matching funds in order to maintain the favorable levels of support both in terms of the number of projects supported and the level of support for each. We are extremely grateful for the matching support provided by MSA sustaining members and individuals. Their support over the years has enabled the program to increase both the number of awards and the maximum amount of each award. For 2003 we are particularly grateful for support provided by Gatan and JEOL.
The MSA Undergraduate Research Scholarship Program is currently soliciting applications from students interested in conducting a research project which involves the use of any microscopy technique. Students should be sponsored by a member of MSA. The maximal award is $3000. The application deadline is Dec 31, 2003. Application forms and instructions are available on the MSA web site, www.msa.microscopy.com. Applications can also be obtained from the MSA Business Office, businessoffice-at-msa.microscopy.com or call toll free at (800) 538-3672. If you have any questions or require additional information regarding the program please contact:
Dr. Ralph Albrecht, University of Wisconsin 1675 Observatory Drive, Madison, WI 53706 (608) 263-3952 or 263-4162; (608) 262-5157 FAX; albrecht-at-ansci.wisc.edu
2003 awardees Renee Lopez-Smith. Microscopic study of fertilization in the fern Ceratopteris: the role of actin in gamete fusion. Southern Illinois University, Advisor: Karen Renzaglia Brigit O'Donnell. Effect of vein pattern alteration on movement of turnip vein clearing virus. New Mexico State University, Advisor: Dr. Souuumitra Ghoshroy Julie Takacs. Inhibition of bacterial invasion mechanisms using anti gC1q-R monoclonal antibody carbon nanotube probes. Franklin and Marshall College, Advisor: Barbara Panessa-Warren Aurelie Thuaire. Characterization of nanocomposites using electron microscopy. McGill University, Advisor: Raynald Gauvin
-- 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 Nov 13 13:18:32 2003
Dear Chris, I suggest you contact Woody White of GE (I think). He has a lot of experience running SEMs analysing radioactive materials. His web site is: home.att.net/~woody.white/ and his e-mail address is on the site. Regards, Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 e-mail: mager-at-interchange.ubc.ca ----- Original Message ----- } From: {HolpC-at-firstenergycorp.com} To: {Microscopy-at-msa.microscopy.com} Sent: Thursday, November 13, 2003 8:44 AM
Thurston,
Using a GIF, we measured the zero-loss energy spread from our CM300FEG as a function of extraction voltage from 4kV down to 1.5kV. The measured energy spread dropped from 0.93eV FWHH at 4kV to 0.65eV at 1.5kV extraction voltage. Backing out the contribution from the GIF gave us a true beam spread of 0.85eV FWHH at 4kV and 0.53eV FWHH at 1.5kV -- see figure 2a of “Estimation of the Electron Beam Energy Spread for TEM Information Limit”, Michael A. O’Keefe, Peter C. Tiemeijer and Maxim V. Sidorov, Microscopy & Microanalysis 8 (2002) suppl 2: 480-481.
We wanted to reduce the energy spread to improve our HRTEM information limit, but found that imaging below 3kV extraction voltage was difficult due to the low beam current. For PEELS I guess you also want a low spread, but I don't know how low a beam current you can tolerate.
Mike O'Keefe
thurston e herricks 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 } ------------------------------------------------------------------------------- } } Hello, } } I currently using a Gatan GIF for EELS on a Tecani thermal field emitter. } I was told that extraction voltage can affect energy resolution to the } spectrometer. If so how may I go about optimizing extraction voltage for } optimizing Gif for high resolution PEELS? } } Thanks } } Thurston Herricks
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 13 17:43:53 2003
We routinely analyse filtrate samples in SEM, specifically for morphology of particles and composition. Unfortunately due to the continuous changes in manufacturers and suppliers merging etc, we no longer have an appropriate source of filter for our needs.
This is the type of filter we used to use. { {...OLE_Obj...} } a Support 0.8um gelman filter
since the merger gelman/pall now produce the following
{ {...OLE_Obj...} } which is unsuitable for our needs. the small round nodules can be easily confused with particles - sends our engineers quite a panic!
we are currently using this one below: { {...OLE_Obj...} } a GN-4 0.8um filter
but this is not ideal as there still are small nodules on the filter surface.
The filter needs to be caustic resistant to high pH, 0.8-0.2um pore size, low background for EDS/XRD analysis.
Can anyone recommend a brand or supplier who could help?
thanks for your assistance.
Samantha Taylor TDG Experimental Officer Alcoa of Australia XRD, Microscopy and Thermal Analysis * Phone:(08) 9410 3588 * Fax: (08) 9410 3167 * Email: Samantha.Taylor-at-Alcoa.com.au
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 01:51:13 2003
The Institute for Transuranium Elements in Karlsruhe, Germany, has experience through many years with electron microscopy (both TEM and SEM) of radioactive materials. Their SEM webpage has the address
Joergen Bilde-Soerensen Materials Research Dept. Risoe National Laboratory Roskilde, Denmark.
-----Original Message----- } From: HolpC-at-firstenergycorp.com [mailto:HolpC-at-firstenergycorp.com] Sent: 13. november 2003 17:45 To: Microscopy-at-msa.microscopy.com
Probably a very basic question, but can anyone tell me whether araldite resin actually infiltrates cells in tissue blocks? I've small (1 mm2) para fixed tissue blocks, which are processed into araldite resin. Does the resin cross cell (and intracellular) membranes?
Thanks.
G McGovern
Veterinary Laboratories Agency (VLA)
This email and any attachments is intended for the named recipient only. Its unauthorised use, disclosure, storage or copying is not permitted. If you have received it in error, please destroy all copies and inform the sender. Whilst this email and associated attachments will have been checked for known viruses whilst within VLA systems we can accept no responsibility once it has left our systems. Communications on VLA's computer systems may be monitored and/or recorded to secure the effective operation of the system and for other lawful purposes.
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 07:52:50 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mikko.uittamo-at-m-real.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, November 14, 2003 at 02:50:05 ---------------------------------------------------------------------------
Email: mikko.uittamo-at-m-real.com Name: Mikko Uittamo
Title-Subject: [Microscopy] Link Isis Autobeam problem
Question: Hi
I dont know if this is the right place for this kind of a question, but bare with me. If you know any better place to post this please inform me.
We have a Oxford link isis software in our microscope and the problem is that when usin autobeam software and opening jobs from network drive, software stops responding for couple of minutes and then it works ok. This happens almost on every job. We just updated the system to w2k and that was when problems started.
Could the problem be with the software or with the new operating system.
Any embedding material must infiltrate all of the cells otherwise you will have a hole where the cell was. Araldite is very viscous and may not best choice for the tissue+fixative combo you are using. On the other hand, it is easier to cut and stain than some other resins. Are you using osmium post-fixation? If not, tissues will be harder to infiltrate (osmium makes membranes more permeable to subsequent processing). If you are not using osmium are you using one or more Araldite:clearing agent steps?
Geoff
McGovern, Gillian 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 Fri Nov 14 10:17:43 2003
Yes, the resin will penetrate right into the cells. In fact, if it doesn't adequately infiltrate right into the cells, then you will be unable to section the tissue ultra-thin, and it will turn into garbage right off your knife.
For this reason, it is important to allow some time for the plastic to penetrate the tissue, usually as a mixture of resin/propylene oxide, until the tissue is adequately infiltrated.
-----Original Message----- } From: McGovern, Gillian [mailto:g.mcgovern-at-vla.defra.gsi.gov.uk] Sent: Friday, November 14, 2003 2:15 AM To: 'Microscopy-at-MSA.Microscopy.Com'
Probably a very basic question, but can anyone tell me whether araldite resin actually infiltrates cells in tissue blocks? I've small (1 mm2) para fixed tissue blocks, which are processed into araldite resin. Does the resin cross cell (and intracellular) membranes?
Thanks.
G McGovern
Veterinary Laboratories Agency (VLA)
This email and any attachments is intended for the named recipient only. Its unauthorised use, disclosure, storage or copying is not permitted. If you have received it in error, please destroy all copies and inform the sender. Whilst this email and associated attachments will have been checked for known viruses whilst within VLA systems we can accept no responsibility once it has left our systems. Communications on VLA's computer systems may be monitored and/or recorded to secure the effective operation of the system and for other lawful purposes.
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 11:02:25 2003
Have you tried the Mykrolis Company (http://www.mykrolis.com/)? They make a polyethylene membrane filter in various pore sizes. I used to use these to do airborne particles in the wafer fab. The filter media has a smooth surface with a support grid on the backside and particles are easily imaged on it.
"Taylor, Samantha" 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 All } } We routinely analyse filtrate samples in SEM, specifically for morphology of } particles and composition. } Unfortunately due to the continuous changes in manufacturers and suppliers } merging etc, we no longer have an appropriate source of filter for our } needs. } } This is the type of filter we used to use. } { {...OLE_Obj...} } a Support 0.8um gelman filter } } since the merger gelman/pall now produce the following } } { {...OLE_Obj...} } } which is unsuitable for our needs. the small round nodules can be easily } confused with particles - sends our engineers quite a panic! } } we are currently using this one below: } { {...OLE_Obj...} } a GN-4 0.8um filter } } but this is not ideal as there still are small nodules on the filter } surface. } } The filter needs to be caustic resistant to high pH, 0.8-0.2um pore size, } low background for EDS/XRD analysis. } } Can anyone recommend a brand or supplier who could help? } } thanks for your assistance. } } Samantha Taylor } TDG Experimental Officer } Alcoa of Australia } XRD, Microscopy and Thermal Analysis } * Phone:(08) 9410 3588 } * Fax: (08) 9410 3167 } * Email: Samantha.Taylor-at-Alcoa.com.au
-- ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ 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 Fri Nov 14 14:32:40 2003
We are using JEM-2010F microscope. The smallest field limiting aperture which JEOL commercially provides is 5 micron. But it is still a little large when we want to get electron diffraction patterns for some special materials. However, we do not want to use NBD mode because of the serious radiation damage. So we are trying to get the information about companies who can make smaller field limiting aperture (~ 1 micron). If you know who can make it, please give me their contact information. Thanks.
Best regards, Qi Zhang ***************************************************** Qi Zhang, Ph.D 164 Phillips Hall, CB#3255 Department of Physics and Astronomy the University of North Carolina at Chapel Hill Chapel Hill, NC 27599 Tel: 919-843-2407 Fax: 919-962-0480 E-mail: qizhang-at-physics.unc.edu qizhang-at-email.unc.edu
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From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 14:39:58 2003
I have a project from a faculty member who is studying a protein, the monomers of which form a ring. I'm using 1% uranyl acetate as the negative stain. This was the method used in the background information they brought to me. Does there exist an even finer grained negative stain? I'm looking for a way to improve the visual quality of the material. Also the operating conditions of my TEM (a Philips 410LS with tungsten filament) are 80Kv and 100Kv (the limit of my TEM), aperture sizes are condensor 200micron, objective 20micron, spotsize 2, zero degree tilt. I'm trying to photograph in the range of 240,000X to 500,000X (the upper limit of my TEM). I would appreciate any suggestions to improve the image of the samples either by way of specimen preparation or operating condition of the instrument. Would metal shadowing be worth trying? Thanks in advance for your help.
Tom Bargar Core Electron Microscopy Laboratory UNMC 986395 Nebraska Medical Center Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 15:15:39 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (wwiggins-at-carolinas.org) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, November 14, 2003 at 10:29:12 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] MListserver: Ilford Service
Question: Mary, I was told that Serco Equipment Services, 800-922-0192, handles maintenance and service contracts in the States for Ilford (an Italian company). We've contracted with them for our 2150. Alternatively, you can try Hunt's Photo & Video, 800-924-8682, in Melrose, MA to see if they can help. Winston
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Winston W. Wiggins, Supervisor Cannon Electron Microscopy Lab Carolinas Medical Center P.O. Box 32861; Charlotte, NC 28232-2861 Ship to: 1000 Blythe Blvd; Charlotte, NC 28203 Ofc:704-355-1267 ; Lab:704-355-7220 ; Fax 704-355-7648 E-mail: Winston.Wiggins-at-CarolinasHealthCare.org ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Hi, everyone,
Does anyone have the name of a service person for a tabletop Ilford 2150 RC paper printer in the Boston area? Thanks in advance,
Mary McKee MGH Renal Unit Charlestown, MA 02129 (617)726-3696
We have had problems with Cy5 and Alexa 647 where the dye fluoresces in the green channel. Basically, we were told that the molecules cleave and one of the products fluoresces in the green channel.
We have been using Bodipy 650/665 phalloidin with even a worse problem. [One of our users wanted to use this as a FRET acceptor; more on this later.] The staining looked really bright through a narrow band Cy3 filter set, which, according to the published spectra, it shouldn't emit at all. After bleaching with 637 nm light, the signal in the Cy3 channel looked even brighter, the result we'd like to get if there were Cy3 to Bodipy FRET. However, this led me to think that maybe exposure of the Bodipy to 637 light was not so much bleaching it as modifying the molecule such that it absorbed and emitted at shorted wavelengths. Perhaps the Bodipy 650 behaves similar to the shorter Bodipy 581 which "shifts to green fluorescence upon peroxidation, a feature that has been exploited for ratiometric measurements of lipid oxidation in live cells"?
Anybody know if this is correct? We'd like to find the most stable dyes in the infra-red channel in addition to Alexa 633.
And as for the FRET, I recommended not using it regardless because "These reactive dyes contain an additional seven-atom aminohexanoyl spacer ("X") between the fluorophore and the succinimidyl ester group. This spacer helps to separate the fluorophore from its point of attachment, potentially reducing the interaction of the fluorophore with the biomolecule to which it is conjugated and making it more accessible to secondary detection reagents such as anti-dye antibodies."
Thanks.
____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 17:40:05 2003
At 12:38 PM 11/14/2003 -0800, you wrote: } We are using JEM-2010F microscope. The smallest field } limiting aperture which JEOL commercially provides is } 5 micron. But it is still a little large when we want } to get electron diffraction patterns for some special } materials. However, we do not want to use NBD mode } because of the serious radiation damage.
You can almost always _throw away_ beam current in probe mode if that's what you want to do. Use the smallest condenser aperture, select the smallest possible probe size, and select the smallest possible convergence angle. If that doesn't get you a small enough current, turn down the electrostatic gun lenses starting with A2. I have produced probes ~5 nm in diameter with relatively small convergence angles ( {0.2 mrad) and ~ 1 pA of beam current on a 2010F. And of course, if you are trying to illuminate a larger area, you can reduce the current density by defocusing the focused probe with C2, the brightness control.
There is also some danger with trying to obtain selected area diffraction patterns from very small regions of a sample - spherical aberration of the objective lens means that the higher order reflections in the pattern don't come from the same part of the sample as the lower order reflections (see Williams and Carter, p. 186-187 for an explanation).
Best wishes, Paul Voyles
Paul Voyles Assistant Professor Materials Science and Engineering Department University of Wisconsin - Madison 1509 University Ave. Madison, WI 53706-1595 Voice: (608) 265-6740 Fax: (608) 262-8353 voyles-at-engr.wisc.edu www.engr.wisc.edu/mse/faculty/voyles_paul.html
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 20:35:28 2003
Well not really - some of the new inkjets have three or four black cartridges - and if you are printing mostly black and white. i.e. grey scale images, the ink sets fro Cone or Quad Black provide a very wide gamut of true grey scale printing by having a range of black to grey inks.
Bill
At 01:40 PM 11/12/2003, DrJohnRuss-at-aol.com wrote:
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From MicroscopyL-request-at-ns.microscopy.com Fri Nov 14 21:51:06 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (rpowell-at-nanoprobes.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, November 14, 2003 at 21:18:28 ---------------------------------------------------------------------------
Email: rpowell-at-nanoprobes.com Name: Rick Powell at Nanoprobes
Organization: Nanoprobes, Incorporated
Title-Subject: [Microscopy] Re: Achieving a fine grain negative stain - Vendor reply
Question: Hello Tom:
I can't advise on the microscope, but at Nanoprobes we do offer a vanadate-based negative stain, NanoVan, which several users find gives a very fine grain structure - check the paper by Gregori et al ( J. Biol. Chem., 272, 58-62 (1997)) for an example.
Reprint: http://www.jbc.org/cgi/reprint/272/1/58.pdf Reprint of our 1994 MSA abstract: http://www.nanoprobes.com/MSANV.html
Since it is based on a lower Z element, the staining is lighter. We also offer a tungstate-based analog, Nano-W, for denser staining. Details are on our web site:
http://www.nanoprobes.com/Nstain.html
Hope this helps,
Rick Powell Nanoprobes, Incorporated
***************************************************************************************** Richard D. Powell * rpowell-at-nanoprobes.com * www.nanoprobes.com NANOPROBES, Incorporated 95 Horse Block Road, Yaphank, NY 11980-9710, USA
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
If so, even with a 5 um SA aperture, you potentially have significant errors in your diffraction pattern. The spherical aberation of the objective lens means that in an SA pattern, different reflections come from slightly different areas. This causes displacement errors from ~0.1 nm for low order reflections up to several um for higher order reflections.
In addition, the focus of the objective lens and the vertical position of the SA aperture are critical. Errors in either can result in additional displacement errors, to as much as 20 um or more if the objective is badly out of focus and/or the SA aperture is in the wrong plane.
With a 5 um SA aperture, the projected diameter on your specimen is likely to be ~250 nm to 500 nm. So, even if you have been very careful in setting up the microscope, it is probable that any but the lowest order reflections actually arise from points outside the area selected!
To be confident that you are getting diffraction from the area you want, you must use a technique which limits the illuminated area using the TEM illumunation optics. In the case where you have a very beam-sensitive specimen, I would suggest that the Riecke method for nanodiffraction is most appropriate. In this case, you use a small (~5 um to 10 um) condenser aperture and set up a parallel or near-parallel illumination condition at the specimen (not a focused probe, so beam damage is minimised). This results in diffraction patterns which look very similar to SAD patterns but do not suffer from the displacement errors of SAD. The technique typically allows the selection of areas down to ~100 nm, depending on the demagnification between the C2 aperture and the specimen.
See, for example, David Williams "Practical Analytical Electron Microscopy in Materials Science" for a more detailed discussion of this whole subject.
-- Larry Stoter JEOL UK
From MicroscopyL-request-at-ns.microscopy.com Sat Nov 15 14:41:32 2003
Ellery E. Frahm wrote: =============================================================== I have a user with gold-coated samples after doing ion probe analyses, and now he needs to do more electron microprobe work, both imaging and quantitative analyses. The gold coats (I'm not certain about thickness) need to be removed. We cannot, however, polish the gold off because he needs to preserve the pits left from the ion probe. As a result, we are looking for chemical means to remove the gold coats. Any suggestions are welcome (particularly those known to work and with details about concentrations, temperatures, etc.). =============================================================== Something we learned quite by accident some number of years ago: Approximately ten to fifteen minutes in an (isotropically created) oxygen plasma (for example, in an SPI Plasma Prep™ II plasma etcher) would remove a 10-15 nm sputtered gold coating (at 100 watts power). We never understood the chemistry of how this happens but it does work. It is essentially a "room temperature" process.
Of course your sample has to be something that won't be etched by an oxygen plasma so if it is a metal, or ceramic, or anything else that is inorganic, you are OK, but not if it is organic. We made our original observations on a clay coated photocopy paper, and the oxygen did not seem to be changing the clay particles (but it did of course remove the acrylic binder holding the clay particles to the paper substrate).
There are advantages to using a "dry process" as opposed to a "wet chemical" approach, at least in some instances.
Chuck
Disclaimer: SPI Supplies is the manufacturer of the Plasma Prep II plasma etcher which is described on URL http://www.2spi.com/catalog/instruments/etchers1.shtml
============================================
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 Sun Nov 16 19:17:12 2003
On Friday, November 14, 2003, at 12:45 PM, tbargar-at-unmc.edu wrote:
} Also the } operating conditions of my TEM (a Philips 410LS with tungsten } filament) are } 80Kv and 100Kv (the limit of my TEM), aperture sizes are condensor } 200micron, objective 20micron, spotsize 2, zero degree tilt. I'm } trying to } photograph in the range of 240,000X to 500,000X (the upper limit of my } TEM). I would appreciate any suggestions to improve the image of the } samples either by way of specimen preparation or operating condition } of the } instrument.
Dear Tom, Since Rick Powell covered the stain half of your question, I thought I'd take a stab at the other half. The negative stain consists of clumps of material typically ~1 - 2 nm in diameter, so the magnification you're using results in blobs of stain ~1/4 -1 mm on your detector. If you lower the mag, you will have smaller, but still visible clumps of stain, and spreading the beam can reduce effects from heating and radiation damage; additionally, you will get a larger field of view. Even with a W filament, you are concentrating a large number of electrons on a very small sample area and few electrons outside this area, so you can cause motion even of very heavy atoms. Yours, Bill Tivol 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 Mon Nov 17 07:39:58 2003
I am looking for a good TEM photo of a Canal of Hering, or cholangiole from the liver. Also, I would like some information on how to maximize one's chances of finding this structure under the beam.
Many thanks, Peggy Harger-Allen EM Lab, VAMC, Indianapolis
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 17 08:17:04 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Donnarae48-at-AOL.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Sunday, November 16, 2003 at 14:43:02 ---------------------------------------------------------------------------
Email: Donnarae48-at-AOL.com Name: Donna Morrow
Organization: Moreau Catholic High School
Education: 9-12th Grade High School
Location: Hayward, California
Question: My biology class did a lab using the microscopes. We are able to magnify up to 400 magnification. We used a scraping from our cheeks and dye to make a slide. We were asked to draw what we saw and lable at least 2 things. I am not sure what I saw. At 400 magnification how clear would an epithelial cell be and would I have been able to see a nucleus. All I saw were blue circles and light blue background and very small round things scattered around. I am wondering if the larger dark blue circles were a nucleus and the smaller specks the cell membranes even though they did not seem to be uniform. The teacher said my slide looked good but I am not sure what I was viewing. Also do you know of any other sight that is available to help me with my lab work with the microscope. We have been trying to find a sight that would show me what an epithelial cell would look like at 400 magnification and I keep running in to brick walls. Thank you for your help
I have a Philips CM-12 TEM. Your operationg conditions seem OK to me for the mag range you indicate, except for the spot size. Drop it down to spot size 4-6 and see what you get for particle resolution. Take a photo at your spot 2 setting, then shoot same particle at lower spot sizes for comparison. I have photographed carbon nanoparticles at those mags and the smaller spot size definitely improved the resolution of the details. Your exposure times will increase, so must have stable samlple. You can also kick up the emission level of the electron gun, or even set tungsten filament tip a little closer to Wehnelt cap (but not so clsoe that you get arcing) to increase the brightness at those smaller spot sizes.
Also, just a reminder to use your cold trap. After a half-hour cooldown, you will reduce any beam induced contamination of the particles you are looking at.
Let me know if you get any improvement - or not - with smaller spot sizes. Good luck!
Gib -- 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
..snip!...
} Also the } operating conditions of my TEM (a Philips 410LS with tungsten filament) are } 80Kv and 100Kv (the limit of my TEM), aperture sizes are condensor } 200micron, objective 20micron, spotsize 2, zero degree tilt. I'm trying to } photograph in the range of 240,000X to 500,000X (the upper limit of my } TEM). I would appreciate any suggestions to improve the image of the } samples either by way of specimen preparation or operating condition of the } instrument.
} Tom Bargar } Core Electron Microscopy Laboratory } UNMC } 986395 Nebraska Medical Center } Omaha, NE 68198-6395 } 402-559-7347 } tbargar-at-unmc.edu
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 17 11:04:17 2003
We can supply gold foil apertures down to 2 micron. We have other size limitations though, so please contact me directly with the OD and thickness of the blank.
Michael R. Nesta General Manager Energy Beam Sciences, Inc. Tel: 413 786-9322 Fax: 413 789-2786 mnesta-at-ebsciences.com www.ebsciences.com "Adding Brilliance to Your Vision"
-----Original Message----- } From: Paul Voyles [mailto:voyles-at-engr.wisc.edu] Sent: Friday, November 14, 2003 6:46 PM To: microscopy-at-msa.microscopy.com
At 12:38 PM 11/14/2003 -0800, you wrote: } We are using JEM-2010F microscope. The smallest field } limiting aperture which JEOL commercially provides is } 5 micron. But it is still a little large when we want } to get electron diffraction patterns for some special } materials. However, we do not want to use NBD mode } because of the serious radiation damage.
You can almost always _throw away_ beam current in probe mode if that's what you want to do. Use the smallest condenser aperture, select the smallest possible probe size, and select the smallest possible convergence angle. If that doesn't get you a small enough current, turn down the electrostatic gun lenses starting with A2. I have produced probes ~5 nm in diameter with relatively small convergence angles ( {0.2 mrad) and ~ 1 pA of beam current on a 2010F. And of course, if you are trying to illuminate a larger area, you can reduce the current density by defocusing the focused probe with C2, the brightness control.
There is also some danger with trying to obtain selected area diffraction patterns from very small regions of a sample - spherical aberration of the objective lens means that the higher order reflections in the pattern don't come from the same part of the sample as the lower order reflections (see Williams and Carter, p. 186-187 for an explanation).
Best wishes, Paul Voyles
Paul Voyles Assistant Professor Materials Science and Engineering Department University of Wisconsin - Madison 1509 University Ave. Madison, WI 53706-1595 Voice: (608) 265-6740 Fax: (608) 262-8353 voyles-at-engr.wisc.edu www.engr.wisc.edu/mse/faculty/voyles_paul.html
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 18 00:11:58 2003
Hi all, I just realised that I've been replying to myself so I'll post my emails here in a new post. Therefore, my initial email is at the bottom. Any advice gratefully accepted.
John Brealey __________________________________________________________________
Hello again,
Still no electron beam. We believe the HV Stabilzer board (new capacitors) and the cable are OK. Gun vacuum is 10 (-5) Torr. Column and camera are 10 (-2) Torr. Gun and camera green lights are on. Can anyone advise on interlocks within the system? We think there are at least four... the gun valve, specimen airlock, camera shutter and camera valve. Something in the system is preventing the filament from switching on. Is there anything else that will prevent the filament turning on? We have tried several filament setups.
Thanks again,
John Brealey
-----Original Message----- } From: John Brealey [mailto:john.brealey-at-imvs.sa.gov.au] Sent: Tuesday, 11 November 2003 01:15 PM To: John Brealey
Hi all,
Our Hitachi H-600 TEM is failing to provide an HV reading at any kV (ie 25, 50, 75, 100). As the microscope is 20 years old I believe the HT cable is the problem. Do you agree? What's the best plan of attack from here? Apart from the HT cable and a dirty gun chamber, are there any other potential causes of this failure? I have never attempted to isolate the HT cable. The vacuum system is fine. I've turned the microscope completely on and off a few times and have vented all compartments to air and back to high vacuum but this has had no effect.
Two days ago I reassembled the rear cosmetic guard that encases the upper vacuum system and the HT cable. Could a slight knock to the cable be enough to finish it off? The microscope was working yesterday, though. I've tried gently wiggling the cable but that gave no response.
The microscope has not been hissing and the HV has appeared quite stable. The only sign that something was not quite right was that some mornings the HV would fail to read at 25kV (but would always read at 50, 75 and 100kV).
In the light of problems encountered by Sarah Ellis (refer to archives) a few months ago I am tempted to clean the gun chamber first (however I doubt if this will fix anything).
Any advice would be gratefully accepted before I contact our institution's electrical engineers.
John Brealey EM Unit The Queen Elizabeth Hospital Institute of Medical & Veterinary Science Adelaide Australia (08) 8222 6612 john.brealey-at-imvs.sa.gov.au
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 18 05:06:17 2003
you mentioned in your e-mail that several interlocks may affect the operation of the filament. But you didn't say whether you had any way of monitoring them.
I use an H7000 so it may not have the same interlocks but camera shutter and camera valve have no effect on the filament. The camera can be closed or at air and the filament will still run and the shutter should not affect the filament (I assume you mean the exposure shutter). If the electron gun was closed or interlock had failed you wouldn't get the high voltage operating either. So this means that if it is one of these interlocks it is most likely to be the specimen airlock.
In the H7000 you must have a proper specimen rod actually in the column to be able to turn the filament on normally. I wonder if there is some simple logic attached to the airlock interlock operation which has gone out of sequence and so maybe the specimen rod should be in a different position from normal. It's worth a try if you haven't already checked it.
Also we have a couple of plug in leads just beside the specimen airlock. The important one is HK-CN4 on our machine - if it is loose or detached then the filament will not come on because presumably this carries the signal from the airlock mechanism that there is a specimen in the microscope. I know this because it happened to us once and it's so easily knocked when you're working near the airlock in the dark.
I am assuming in all this that you mean you've got a high voltage but no beam from the filament.
Good luck,
Malcolm
Malcolm Haswell e.m. unit Chemispec School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear SR1 3SD UK tel no: 0191 515 2872 / 3468 e-mail: malcolm.haswell-at-sunderland.ac.uk
----- Original Message ----- } From: John Brealey {john.brealey-at-imvs.sa.gov.au}
Dear all, I asked about this once before but got no replies. Is there any introductory book or nice review paper on the TEM of semiconductors? A student needs to learn about basic features such as 60° threading dislocations and such things but at least in recent publications everything is taken as if one already understands the basics, which we don't. Please give us a hint about where to start.
Thanks
-- Ian MacLaren Technische Universität Darmstadt Material- und Geowissenschaften Petersenstr. 23 64287 Darmstadt Germany http://www.tu-darmstadt.de/fb/ms/fg/sf/projekte/maclaren-Dateien/maclaren.html
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 18 12:03:53 2003
With the H600 as with many other instruments you may find that you cannot obtain a beam until the specimen rod has tripped the microswitch within the specimen exchange unit? Try putting the empty rod into the microscope? Also a green light means that the system has received an instruction - not that it has completed an action. Another common problem is that the gun airlock may jam closed - you will have an emission current in this case but no beam. The figures that you give for column and camera suggest backing pump pressure which is not enough to allow the system to operate correctly - when you run from air do you hear valves switching as the system moves from backing pump to diffusion pump? To check a valve opens place a stethoscope against the valve or rest a screw driver blade against the valve with your ear on the handle end! You should hear the action if its working? Anther reason for no filament is a failure of the diodes in the filament circuit within the HT tank but this is a rare problem.
Steve Chapman Senior Consultant Protrain Electron Microscopy Training and Consultancy World Wide Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Mob 07711 606967 www.emcourses.com
----- Original Message ----- } From: "John Brealey" {john.brealey-at-imvs.sa.gov.au} To: "Listserver" {Microscopy-at-MSA.Microscopy.Com} Sent: Tuesday, November 18, 2003 6:21 AM
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (weazzi-at-unity.ncsu.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 17, 2003 at 11:45:51 ---------------------------------------------------------------------------
Email: weazzi-at-unity.ncsu.edu Name: Wassim Azzi
Organization: NCSU
Title-Subject: [Microscopy] MListserver:
Question: I would please like to get information about E-Beam damage of samples in the SEM. I am primarily looking for recent publications and advancements. Any inofrmation provided will be greatly appreciaed. Regards. Wassim Azzi
Tom: Another negative stain to consider is uranyl formate. It is a bit fussy about staying in solution so you need to make it up just prior to use.
Mr. Donald Gantz Dept. Physiology and Biophysics Center for Advanced Biomedical Research Boston University School of Medicine 715 Albany Street Boston, MA 02118 email: {mailto:gantz-at-bu.edu} gantz-at-bu.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 18 14:57:17 2003
The 37th Annual Fall Symposium of the New England Society for Microscopy (NESM) will be held on Tuesday, December 9th at Gordon College in Wenham, MA. This is the second year that Gordon College has hosted this meeting.
The symposium will begin at 12Noon with registration and at 1 pm, the Technical Sessions (with talks from both the biological and materials sciences) will begin with a short coffee break between the sessions. Following the talks, the Annual Business meeting will convene at 5:00 pm with the election of new officers as one of the main agenda items. A dinner will follow with Prof. Paul Goldberg of the Archaeology Department of Boston University as the after-dinner speaker.
For details re: registration, fees, program (speakers and titles), directions to Gordon College etc., please visit NESM's website: http://prism.mit.edu:8083. Click on "Current Newsletter".
We hope to see many of you there!
Peggy Sherwood Corresponding Secretary, NESM
From MicroscopyL-request-at-ns.microscopy.com Tue Nov 18 18:50:43 2003
Thankyou to all respondents for your replies. The microscope is in pieces at the moment so I can't check your suggestions.
To clarify... yes, we have fixed the HV but now there is absolutely no response from the filament. Our technicians have checked the circuitry of the HV Stabilzer board and believe all is well there. Voltages change as we turn the filament knob and the bias knob however there is no movement in the HV / beam current meter. I have run the microscope for 30 minutes at 100kV with filament and bias on maximum, then, leaking the gun chamber to air, have checked the filament shield and it was stone cold. The filament assembly can only be inserted one way due to the presence of a set screw in the gun assembly. We have turned the filament clockwise (as stated in the manual) until we feel the resistance of the filament pins against the gun contacts. We have tried several clean filament assemblies. All moveable apertures are out. Specimen holder in or out doesn't make any difference. We have removed the gun cable twice from the HV tank. Our technicians are happy with the circuitry of the cable and believe it is making good contact in the HV tank.
I may have been incorrect with the vacuum units on the vacuum gauge. The meter is at 10(-5) for the gun chamber (Penning gauge) and 10 (-2) for the column and camera chambers (Pirani gauge). These readings are the same as they always have been.
Currently we are checking the gun valve. Could this be stuck shut even though the vacuum system thinks it is open? When the camera valve opens I can feel its movement, so that appears to be operating. The camera system appears to be working normally when I take a dummy exposure, ie, negative plates are feeding normally.
I will try your suggestions when the microscope is reassembled.
Thanks again, John Brealey
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 05:50:47 2003
-----Original Message----- } From: Ian MacLaren [mailto:maclaren-at-tu-darmstadt.de] Sent: 18. november 2003 16:40 To: Microscopy Listserver
Dear All,
We are interested in purchasing a new ESEM. The ESEM will mostly be used for biological and some material sciences. I would like to hear your suggestions regarding which ESEM I should choose (manufacturer and model of the ESEM) and any particular reasons. Thank you in advance.
Ping Li
-- Ping Li, Ph.D. Director, Scientific Imaging Suite Department of Biology Dalhousie University Halifax, NS B3H 4J1 Canada
We are considering using a cheaper, Au target instead of a Au/Pd target, for our sputter coater. With our FEI XL30 SEM we should not notice any significant difference, is that correct?
I still wanted to know if the Au coating may be of lesser quality, and that this may show up in the future, examining the same samples with a better SEM or FESEM. In the long term, can we obtain a good quality preparation by re-coating with Au/Pd a sample that was previously coated with Au?
I will very much appreciate any hint on this.
Best regards,
Martín J. Ramírez División Aracnología Museo Argentino de Ciencias Naturales Av. Angel Gallardo 470 C1405DJR Buenos Aires Argentina tel +54 11 4982-8370 fax +54 11 4982-4494
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 07:41:24 2003
Listers I have a file in JPEG format. I need to reduce the file size. Currently the file size is 22 Meg and too large to import into my application. Is there a function in Paintshop that will perform a file size reduction? Thank you
Robert Fowler Quality Assurance Failure Analysis Technician TDK Components USA, Inc. Multilayer Ceramic Capacitor Division 1 TDK Boulevard Peachtree City GA 30269-2051 Telephone: (770) 631-0410 Ext.249 Fax: (770) 487-1460 email: robert.fowler-at-tdktca.com www.component.tdk.com
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From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 08:34:43 2003
Hello all, I've noticed that ASTM method D:3015-95 "Standard Practice for Microscopical Examination of Pigment Dispersion in Plastic Compounds" has been withdrawn (2001). Can anyone provide an EM method for the qualitative or quantitative assessment of pigment dispersion in polymeric materials.
Thanks, Paul
Paul J. Gerroir Microscopy Materials Characterization Xerox Research Centre of Canada 2660 Speakman Drive Mississauga, Ontario L5K 2L1
} Hello Microscopist, Ladies and gents, } } Let me set the stage, no pun intended. We have a Jeol 2010. I } managed and trained students on this instrument from 1993 to some time } } in 2000. During that time I had to tongue lash, in a nice way, users } for } not activating the ACD cycle at the end of the day. This happened ~ 20 } } times with the only failing being a bellows in the gun valve after ~ } 10,000 cycles. This will make since to you if you have a 2010 with the } } old gun valve design. } I still on occasion use the instrument. Please understand, I need } to } refer to specifics of day. This in not intended to be a assault on } Jeol. I am the culprit. } I logged onto the instrument a few days after an new filament was } installed. I found the image of the filament to be moving back and } forth, ~1 Hz, ~1mm at x100K.... This I interpreted as an instrument } charging } problem. I continued to survey samples, encountering 2 more technical } annoyances followed by a 4th that brought the session to an end. We } all } know these things happen. } Perhaps more frustrated than I realize, I left the instrument } without activating the ACD and with the HT at 100 kV. My mistake was } found the following morning. } It's been implied that my mistake caused a contamination of the } gun } so severe that 2 additional filaments were contaminated by it as fast } as they were installed. This I } find hard to believe but I suppose stranger things have happened and } what do I know. } I'll not share other thoughts or the opinions of others for now } least I } bias your response. } } Well what do you think? } 1. has anyone had this happen to their instrument? } 2. Has anyone ever heard of a gun contaminating a filament to this } extent?. } 3. other comments? } } } thanks, } Bruce Brinson } Rice U.
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 10:01:32 2003
In a message dated 11/19/03 10:56:36 AM, Paul.Gerroir-at-crt.xerox.com writes:
} I've noticed that ASTM method D:3015-95 "Standard Practice for Microscopical } Examination of Pigment Dispersion in Plastic Compounds" has been withdrawn } (2001). Can anyone provide an EM method for the qualitative or quantitative } assessment of pigment dispersion in polymeric materials.
I'm unfortunately not familiar with the ASTM method and don't have it here in my library, but let me take a chance and jump in with something that may be helpful. Dispersion of pigment is probably ideally random but with some tendency toward clumping or clustering. On the other hand, for appearance purposes, it would be nice if the pigment could be self-avoiding, which simply means the particles would be as uniformly spaced as possible (not sure how you would produce that in this case, but in some products like Dupont Corian countertops you can see that they manage to do it, and nature does it easily - usually by chemical depletion of the region around a particle - but that's another long story).
Anyway, the measurement procedure is to determine the mean nearest neighbor distance from suitable images. Some software packages will do this directly (e.g. Fovea Pro), otherwise you would need to record the coordinates and calculate the distance to neighbors to find the minimum or nearest neighbor for each particle. A subtle refinement, which may be unimportant if you have a lot of small features and large images, is to skip any feature that is nearer to the border than to another particle, because you may not have found its actual nearest neighbor. Once you have the mean nearest neighbor distance, you can characterize the degree of clustering or self-avoidance. A true random distribution would have a mean nearest neighbor distance equal to 0.5 divided by the square root of the number of features per unit area. Notice that the units of that come out in distance, as they should. If the ratio of actual nearest neighbor distance to the "ideal" value calculated from that equation is less than one it indicates clumping or clustering (and the ratio measures the extent). And vice versa if the ratio is greater than one it indicates and measures self-avoidance.
Hope that helps
John Russ
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 10:12:08 2003
It is my understanding that Au/Pd does a better job at coating rougher surfaces because of its smaller grain size. It may depend on what you coat.
Ron L
-----Original Message----- } From: Martin Ramirez [mailto:ramirez-at-amnh.org] Sent: Wednesday, November 19, 2003 8:40 AM To: Microscopy-at-sparc5.microscopy.com
Hi,
We are considering using a cheaper, Au target instead of a Au/Pd target, for our sputter coater. With our FEI XL30 SEM we should not notice any significant difference, is that correct?
I still wanted to know if the Au coating may be of lesser quality, and that this may show up in the future, examining the same samples with a better SEM or FESEM. In the long term, can we obtain a good quality preparation by re-coating with Au/Pd a sample that was previously coated with Au?
I will very much appreciate any hint on this.
Best regards,
Martín J. Ramírez División Aracnología Museo Argentino de Ciencias Naturales Av. Angel Gallardo 470 C1405DJR Buenos Aires Argentina tel +54 11 4982-8370 fax +54 11 4982-4494
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 10:40:35 2003
I had mentioned in my Lacey grid review that the new batch of 4489 film had made its way into rotation here. And after trying a few suggestions, I did finally call Kodak. From some reports (from vendors and users on the list) Kodak wasn't acknowledging a problem, and that assumption contributed to a reluctance to prioritize calling and complaining about the new film.
I finally called them and was surpised. Sure enough! They've got a new procedure for hand developing that is different than the printed package. I tested the method this morning, and had another user test it after me.
And first looks seem to say: Back in business! (phew - no need to design/construct a nitrogen bubbler system)
I've quoted the pertinent parts of the procedure below. I might be behind the ball on this but looking back at the list traffic it seems this particular solution hasn't been announced here. -quote- PROCESSING Develop: KODAK Developer D-19 diluted 1 part developer and 2 parts water can be used for 4 minutes at 20°C (68°F). Hand agitate by quickly inserting the film rack into the developer tank, tapping the rack on the tank bottom to dislodge any air bubbles that may cling to the film surface. Every 10 seconds, rapidly remove the film rack completely from the developer tank and tip the film rack 45° to one side of the tank and then tip the film rack 45° to the other side of the tank, allowing the developer to drain into the tank (complete tipping procedure is only a few seconds.) Rapidly return the film rack into the tank. Repeat this process throughout the development cycle. Development times longer than 4 minutes will result in a rapid increase in background fog.
Rinse: For 1 1/2 minutes in running water at 18 to 21°C (65 to 70°F) with continuous agitation. Do not use conventional stop bath solutions because they may produce a mottled appearance on films used for electron exposures. -quote- Fix as normal....
I hope that helps, and I apologize to those who find this old news.
Geoff Williams Microscopy Facility Supervisor
Central Michigan University Biology Department Microscopy Facility http://www.cst.cmich.edu/centers/microscopy/
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 11:14:40 2003
Hello Everyone: I am looking for a something that will selectively stain aromatic or amide functional groups. Sawyer and Grubb list ruthenium tetroxide, silver sulfide, and mercuric trifluoracetate as possible stains for aromatics, but RuO4 and Ag2S are not selective and mercuric trifluoracetate is very toxic (we are trying to limit our usage of Hg anyway).
Does anyone know of anything else that will select an aromatic group (vs. alcohol, ester, ether, and acid groups)?
Another option is to stain the amide groups in my sample. Sawyer and Grubb list Tin chloride as a possible stain, but don't give a method for using it. Does anyone have experience with this stain?
Thank you,
Jessica Cervantes Bend Research, Inc 64550 Research Rd Bend, OR 97701 (541) 382-4100 page (541) 382-0212 x240
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 11:25:14 2003
} } } Listers I have a file in JPEG format. I need to reduce the file size. } Currently the file size is 22 Meg and too large to import into my } application. Is there a function in Paintshop that will perform a file size } reduction? Thank you }
If the image does not import because the number of pixels is too great, you can easily resize. With Jasc Paintshop Pro, from the image menu, choose resize. You will have the choice of choosing a specific pixel size or a percentage change in size. You should make sure that the "Maintain aspect ratio" box is checked so that the aspect ratio of the image is not changed.
A similar function is available with Irfanview - choose Resize/Resample from the Image menu.
Resizing will change the resolution of the image, so keep this in mind.
Hope this helps.
-- Larry D. Hanke, P.E. Materials Evaluation and Engineering, Inc. Practical Solutions Through Technology and Innovation http://www.mee-inc.com (763) 449-8870
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 12:12:42 2003
} Hi, } } We are considering using a cheaper, Au target instead of a } Au/Pd target, } for our sputter coater. With our FEI XL30 SEM we should not } notice any } significant difference, is that correct?
I think you will see difference starting with x20k-50k magnifications. Au/Pd target is definitely better for high magnification work.
} I still wanted to know if the Au coating may be of lesser } quality, and that } this may show up in the future, examining the same samples } with a better } SEM or FESEM. In the long term, can we obtain a good quality } preparation } by re-coating with Au/Pd a sample that was previously coated with Au?
No. Au is forming "grains" which is bigger then Au/Pd "grains". Recoating will not eliminate them.
Vladimir
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 12:26:57 2003
my understanding is that gold coating produces a coarser surface coating because Au alone nucleates on the surface as it is coated and islands of Au limit the image quality in high resolution SEM. The Au/Pd alloy forms smaller more numerous islands and so provides some improvement in resolution over gold. Simply coating a previously gold coated sample with Au/Pd will not improve the situation but should make it worse because the sample will be coated with a thicker layer.
I am surprised that you find much difference in price between Au and Au/Pd targets. In the UK they are either the same price or I am sure that I have seen Au/Pd a bit cheaper.
I think the problem is defining what you mean by high resolution in SEM but normally this should be achieved by a good traditional SEM or certainly will be by a field emission SEM. I'm sorry I have no figures - the information I have just says Au/Pd is noticeably better than Au at high resolution. Instinctively I would guess the difference would be noticeable somewhere between 20k and 40k but I would be interested to know if anyone could quote any real hard figures.
There are now lots of alternatives to Au and Au/Pd which give even better resolution and I have seen some remarkable images taken with very low voltages on a high pressure field emission SEM - but I can only dream ...
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: Martin Ramirez {ramirez-at-amnh.org}
Dear Friends,
A user of the lab has asked me to separate urban myth from reality regarding HBO bulbs used for fluorescence microscopy.
His questions are:
What is the difference between a 50W bulb and a 100W bulb? He says he set up a demo, identical except one scope had a 50W bulb, the other a 100W bulb, and he didn't think he could tell much difference.
Do bulbs have a finite life, ie., should they be changed at some maximum hours even if still lighting up and appearing OK? He has heard various advice, like change after 200 hours or 250 hours. Change when the glass gets dark, how dark? Is it true that bulbs with many hours may explode?
He has only used Osram bulbs, are there other brands available and what criteria would one use to select one brand over another?
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 Wed Nov 19 14:49:57 2003
The method Geoff outlines below is the only one we have managed to get to work with any consistency with the new formulation. It is similar to the technique we had used previously, the critical difference being do not ever, repeat ever, let the film stand still. Our method for the old formulation processed up to 25 plates in two batches - we would alternately agitate half the films at the time leaving the other standing in developer (for 20 or so seconds before its turn). When we tested this with the new formulation around a third of the sheets were very unevenly developed (even though developer would be moving around the still batch). If we develop in smaller batches using more or less constant agitation we have no serious problems. I an
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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Dear list,
I had mentioned in my Lacey grid review that the new batch of 4489 film had made its way into rotation here. And after trying a few suggestions, I did finally call Kodak. From some reports (from vendors and users on the list) Kodak wasn't acknowledging a problem, and that assumption contributed to a reluctance to prioritize calling and complaining about the new film.
I finally called them and was surpised. Sure enough! They've got a new procedure for hand developing that is different than the printed package. I tested the method this morning, and had another user test it after me.
And first looks seem to say: Back in business! (phew - no need to design/construct a nitrogen bubbler system)
I've quoted the pertinent parts of the procedure below. I might be behind the ball on this but looking back at the list traffic it seems this particular solution hasn't been announced here. -quote- PROCESSING Develop: KODAK Developer D-19 diluted 1 part developer and 2 parts water can be used for 4 minutes at 20°C (68°F). Hand agitate by quickly inserting the film rack into the developer tank, tapping the rack on the tank bottom to dislodge any air bubbles that may cling to the film surface. Every 10 seconds, rapidly remove the film rack completely from the developer tank and tip the film rack 45° to one side of the tank and then tip the film rack 45° to the other side of the tank, allowing the developer to drain into the tank (complete tipping procedure is only a few seconds.) Rapidly return the film rack into the tank. Repeat this process throughout the development cycle. Development times longer than 4 minutes will result in a rapid increase in background fog.
Rinse: For 1 1/2 minutes in running water at 18 to 21°C (65 to 70°F) with continuous agitation. Do not use conventional stop bath solutions because they may produce a mottled appearance on films used for electron exposures. -quote- Fix as normal....
I hope that helps, and I apologize to those who find this old news.
Geoff Williams Microscopy Facility Supervisor
Central Michigan University Biology Department Microscopy Facility http://www.cst.cmich.edu/centers/microscopy/
______________________________________________________ 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 Nov 19 14:54:14 2003
Hi I'm looking for a supplier of carbon string (fiber) for making carbon films. Any recommendation/suggestion is greatly appreciated.
Gang (Greg) Ning, Ph.D. Director, Electron Microscopy Facility Huck Institute for Life Sciences The Pennsylvania State University 1 South Frear Lab University Park, PA 16802 Phone: 814-863-0994 Fax: 814-863-1357 Email: gxn7-at-psu.edu http://www.lsc.psu.edu/stf/em/home.html
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 19 16:25:17 2003
There are often contradicting explanations in the Hitachi manuals in relation to filament position in its holder. Some say in line with the two slots some show 90 degs. If in doubt insert a filament, remove the HT cable from the tank and check with a multimeter for continuity across two of the contacts; never done it but it should work?
If the gun valve does not open, even when instructed to do so, most Hitachi TEM will still allow HT and filament ON, its just impossible to find the beam (ask some of my customers!!$$??) Under these conditions you have indications on the emission meter that you have HT and an emission current that responds to filament control change. This is not a dirt problem forget cleaning, it appears not to be a supply problem to the tank, it may still be a vacuum problem in relation to the column from your figures, my bet is the filament supply diodes in the tank; if I remember correctly they are rated at 6 amp?
Good luck
Steve Chapman Senior Consultant Protrain Electron Microscopy Training and Consultancy World Wide Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Mob 07711 606967 www.emcourses.com
----- Original Message ----- } From: "John Brealey" {john.brealey-at-imvs.sa.gov.au} To: "Listserver" {Microscopy-at-MSA.Microscopy.Com} Sent: Wednesday, November 19, 2003 1:00 AM
-- [ From: Garber, Charles A. * EMC.Ver #3.1 ] --
Martín J. Ramírez wrote: ====================================================== We are considering using a cheaper, Au target instead of a Au/Pd target, for our sputter coater. With our FEI XL30 SEM we should not notice any significant difference, is that correct?
I still wanted to know if the Au coating may be of lesser quality, and that this may show up in the future, examining the same samples with a better SEM or FESEM. In the long term, can we obtain a good quality preparation by re-coating with Au/Pd a sample that was previously coated with Au?
I will very much appreciate any hint on this. ========================================================== You should use caution here.
When you say "cheaper", just why is it cheaper? If it is because you found someone who will sell you the same identical item at a lower profit margin, then you are getting a good deal and you should take it. But you sound worried that this might not be the case, that it might be cheaper because it might be lacking in some way in quality and that would be purity. The higher the purity the more expensive is the precious metal, not because you are getting more of the precious metal, but because of the economics associated with the production step, when comparing for example, 0.999 vs. 0.9999 vs. 0.99999.
So what if you are getting something of inferior quality, how does it affect the final result? The main problem is that while the gold sputters easily (because of its low work function), the impurities stay behind on the cathode, forming a barrier, preventing further sputtering from occurring. We have heard stories that some of these impurities at some point start to come down on the sample producing an indeterminable artifact effect. True the stories are anecdotal, but it does make sense that that could happen.
You can always decide at some time to put on another layer of gold. However , each time you make the total layer thicker, you start covering up the smallest features you are seeing, so you lose detail. If impurity particles are showing down on the same, you are then going to be covering them up and "seeing" them in your recoated sample. That, in our opinion, does not make for good laboratory practice.
So while I admit that is will sound like a very self-serving kind of statement, since SPI Supplies supplies high purity cathodes, the best advice is to stick to suppliers that you know well and who are selling to you cathodes of known high purity, at least 0.9999 and perhaps even 0.99995. If you can find a jeweler who has gold foil of those levels of purity, by all means, if you can save some money, why not? But make sure you know your vendor and that you trust what you are being told about purity.
RE: Au vs Au/Pd and difference in grain size. Can some one cite a reference or two where that has actually been studied and confirmed for sputtered coatings? I know that this was done years ago on vacuum evaporated coatings , but I can't put my finger on a reference where it was done with sputtered coatings.
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 Nov 19 16:40:51 2003
We tried constant agitation, but in the bath and just vertical (the holder and solution bath don't allow for anything but up and down agitation). The results were somewhat satisfactory but there seems to be something to the 45º and the tapping, because the film suddenly looked more like the old film when I was finished with it.
I did the 10 second interval as outlined with success. So shaking for 2-3 seconds and resting for 5 or so seconds.
Geoff Williams Microscopy Facility Supervisor
Central Michigan University Biology Department Microscopy Facility http://www.cst.cmich.edu/centers/microscopy/
} -----Original Message----- } From: Ian Hallett [mailto:IHallett-at-hortresearch.co.nz] } Sent: Wednesday, November 19, 2003 3:51 PM } To: willi1gl-at-cmich.edu; microscopy-at-sparc5.microscopy.com } Subject: [Microscopy] Re: Kodak 4489 (working solution for low budgetlabs) } } } } ------------------------------------------------------------------------ -- } ---- } 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 } ------------------------------------------------------------------------ -- } ----- } } Geoff and listers } } The method Geoff outlines below is the only one we have managed to get to } work with any consistency with the new formulation. It is similar to the } technique we had used previously, the critical difference being do not } ever, repeat ever, let the film stand still. Our method for the old } formulation processed up to 25 plates in two batches - we would } alternately agitate half the films at the time leaving the other standing } in developer (for 20 or so seconds before its turn). When we tested this } with the new formulation around a third of the sheets were very unevenly } developed (even though developer would be moving around the still batch). } If we develop in smaller batches using more or less constant agitation we } have no serious problems. } I } an } } 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 } } } } } } "Geoff Williams" {willi1gl-at-cmich.edu} 20/11/2003 5:48:47 } } } } } } ------------------------------------------------------------------------ -- } ---- } 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, } } I had mentioned in my Lacey grid review that the new batch of 4489 film } had made its way into rotation here. And after trying a few } suggestions, I did finally call Kodak. From some reports (from vendors } and users on the list) Kodak wasn't acknowledging a problem, and that } assumption contributed to a reluctance to prioritize calling and } complaining about the new film. } } I finally called them and was surpised. Sure enough! They've got a new } procedure for hand developing that is different than the printed } package. I tested the method this morning, and had another user test it } after me. } } And first looks seem to say: Back in business! (phew - no need to } design/construct a nitrogen bubbler system) } } I've quoted the pertinent parts of the procedure below. I might be } behind the ball on this but looking back at the list traffic it seems } this particular solution hasn't been announced here. } -quote- } PROCESSING } Develop: KODAK Developer D-19 diluted 1 part developer and 2 parts water } can be used for 4 minutes at 20°C (68°F). } Hand agitate by quickly inserting the film rack into the developer tank, } tapping the rack on the tank bottom to dislodge any air bubbles that may } cling to the film surface. Every 10 seconds, rapidly remove the film } rack completely from the developer tank and tip the film rack 45° to one } side of the tank and then tip the film rack 45° to the other side of the } tank, allowing the developer to drain into the tank (complete tipping } procedure is only a few seconds.) Rapidly return the film rack into the } tank. Repeat this process throughout the development cycle. Development } times longer than 4 minutes will result in a rapid increase in } background fog. } } Rinse: For 1 1/2 minutes in running water at 18 to 21°C (65 to 70°F) } with continuous agitation. Do not use conventional stop bath solutions } because they may produce a mottled appearance on films used for electron } exposures. } -quote- } Fix as normal.... } } I hope that helps, and I apologize to those who find this old news. } } Geoff Williams } Microscopy Facility Supervisor } } Central Michigan University Biology Department Microscopy Facility } http://www.cst.cmich.edu/centers/microscopy/ } } } } } } } } ______________________________________________________ } 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 Nov 19 17:04:08 2003
On Wednesday, November 19, 2003, at 10:36 AM, Jon Krupp wrote:
} Do bulbs have a finite life, ie., should they be changed at some } maximum } hours even if still lighting up and appearing OK? He has heard various } advice, like change after 200 hours or 250 hours. Change when the glass } gets dark, how dark? Is it true that bulbs with many hours may explode? } Dear Jon, I think that the bulbs you're talking about are low-pressure, so there is not a serious explosion hazard--more experienced people may correct me here. When I was using a high-pressure Hg lamp for photochemistry, rated for 100 hours, I turned on one that had been in use and was adjusting the air flow valve (to achieve the correct temperature) when there was a loud bang and a shower of quartz dust about 10 cm from my hand. The lamp housing protected me from injury, but there were several dents in the steel, one of the expensive quartz lenses was bruised (it could have been worse) and the reflector was destroyed. The company informed me that that was the usual failure mode. The next time, we set up all the experiments, turned the new lamp on, ran 24 hours a day until we were done--at about 400 hours--turned the lamp off, and carefully removed it, wrapped it in bubble paper, and left it in the back of a hood until the safety people could dispose of it. Temperature cycling is a lot worse for the lamp than running it continually, but for a low-pressure lamp, YMMV. Yours, Bill Tivol 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 Wed Nov 19 17:12:13 2003
Gang (Greg) Ning wrote: ========================================================== I'm looking for a supplier of carbon string (fiber) for making carbon films. Any recommendation/suggestion is greatly appreciated. ========================================================= Both carbon string and fiber is available from the main suppliers of EM consumables such as SPI Supplies. You can see the range of diameters offered on URL http://www.2spi.com/catalog/spec_prep/carbon-fiber.shtml
I would urge a word of caution: If you are contemplating using the string in a mechanical pumped vacuum system for the production of TEM support films , you will be disapointed with the result. You really do need a diffusion pumped system or better such as in a vacuum evaporator. And in that instance, carbon rods will give a better result than the string.
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 Nov 19 22:02:31 2003
Depending on which XL-30 you have, there may or may not be a difference. If you are using the thermal SFEG, it will definitely make a difference. With SFEG or non-SFEG systems, if resolution is good enough at high mag (} 75KX), you can see the Au coating as a web-like structure on the surface. If you coat over this with Au/Pd or Pt, it will just make the structure more obvious. Cold/magnetron coaters with Au/Pd targets do a very good job laying down an ultra fine film. Pt works well too. I use a Denton Desk II coater for semiconductor work up to 350KX. Au alone is not satisfactory even at 50KX since the Au structure distorts the image. I have used an Anatech Hummer VII coater before and also got good results. So it would seem to me that a good coater with an Au/Pd target will do a superior job over Au for high resolution, high mag imaging. Pt targets are more costly but are even better.
Use a low current, high vacuum setting. I'm running the Denton at 20mA, 60mT for between 30-40 seconds.
There are basic price point differences between Au/Pd and Pt. Pt is higher cost. However, keep in mind that many places sell targets with different thicknesses. A cheaper thin target will not last as long as a modestly more expensive thicker target. Used on a regular basis, my targets last over two years before needing replacement. The coating is only about 50-70A thick. My work ranges from 100X to 350KX. Au/Pd works for all but the most demanding specimens. If I were to choose one target, it would be Pt. I would use it for every specimen. But some specimens are better analyzed with EDS using Au/Pd, others with Pt. So it depends.
The other factor is to keep coated specimens under vacuum storage when not being used. Invariably, there will be Carbon polymerization that shows up as scan areas. These can be reduced by keeping oil out of the chamber and atmosphere away from the specimen. A small trap in the coater vacuum line helps keep the coater from backstreaming. Probably the ultimate coater is a turbo pumped system. But I have yet to find one that is user friendly enough and cost effective to justify purchasing.
gary g.
At 08:21 AM 11/19/2003, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 00:17:59 2003
Our technician Terry Fitton has summarised the current situation on our microscope. Here is his email to me...
Hi John. Thanks for the new emails. I cant get over what a great bucketful of talent you can call on. If we could get help like this on other instruments life would be much easier. I think we should clarify the current situation for others so we dont waste their time and cause confusion. Here is the situation at present as I see it. 1 Outputs from the stabiliser board to the tank are within spec. for HV, filament and bias. Voltages are OK and vary with adjustment of controls on console as required. 2 We have checked filament connections, hv cable continuity and its connections to the tank and filament and all appears OK. 3 Gun airlock was jammed closed and is currently being repaired. 4 We have indication of HV but no apparent filament current. 5 Still have to check vacuum values for column and camera. 6 Still have to check interlocks on specimen airlock and camera airlock. I am interested in Steve Chapman's idea of faulty diodes in the filament circuit inside the tank. Could he be persuaded to elaborate a little as we have no experience of tank insides. Can we do any checking without getting covered in stinking oil. Joel's suggestions have already been covered with all present and correct. That should do for starters. Cheers Terry
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 02:38:48 2003
Like Many others we suffered from the change in the formulation of our EM film. We tried many alternative methods of reducing the fogging and by far the best, and the easiest was changing to Ilford PQ universal developer. We don't even have to agitate to get smooth, repeatable results! Ken Blight Senior Scientific Officer Cancer Research UK London
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 04:47:46 2003
I am not familiar with the ACD you mention but am very familiar with electron guns so lets have a look at your problem?
Contamination in the gun area may usually be attributed to vacuum levels and the resulting high voltage discharge. Spoil the vacuum and you reduce the insulating effect such that high voltage discharge occurs and this cracks gasses that deposit around the gun chamber as a brown/orange layer. This layer seems to be very absorbent and acts like a gas sponge when you re apply a vacuum. The constant outgassing of course leading to even more contamination of the gun chamber. To get back to normal ALL this contamination must be removed
Within the cathode assembly the filament and its base will also give an indication of poor vacuum or poor quality filaments. I have set this out in detail on our interactive EM maintenance CD but this is a brief resume of what you will see.
1. The filament will have suffered from gas attack resulting in extreme thinning over the complete filament, not just at the tip as you would see in a normal failure. 2. The filament base will also become discoloured taking on the same orange/brown tint as the gun chamber. 3. Poor quality filaments that are made up of a high percentage of rubbish will very rapidly contaminate the cathode assembly as the rubbish evaporates. The filament break is normal but the life and the base will indicate problems; the base will be orange/brown.
I have worked with almost every type/make of EM and from time to time the manufacturers have a batch of filaments that are, through no fault of their own, of poor quality. Short life high contamination are instant indicators. I have probably mentioned before that if you have a good box of filaments NEVER use all of them, save two. In a situation where you believe you have a filament problem you may then go back to your good box and confirm that they run correctly. Pop the remainder of the bad box back to your supplier whom I am sure will provide you with a new box.
Good luck with your problem, is it a poor filament problem have you just opened a new box?
Steve Chapman Senior Consultant Protrain Electron Microscopy Training and Consultancy World Wide Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Mob 07711 606967 www.emcourses.com
----- Original Message ----- } From: "Bruce Brinson" {brinson-at-rice.edu} To: {microscopy-at-msa.microscopy.com} Sent: Wednesday, November 19, 2003 6:00 PM
Robert writes ...
} Listers I have a file in JPEG format. I need to reduce the file size. } Currently the file size is 22 Meg and too large to import into my } application. Is there a function in Paintshop that will perform a } file size reduction? Thank you
I cannot speak for Paintshop, but Irfanview should be able to load the JPEG and resave it with a lower quality compression setting. In the context of scientific imagery, I should warn you that resaving as JPEG is a lossy operation.
If you cannot make the file much smaller with Irfanview, you can change the print resolution (pixels/inch), and make the file size more appropriate for your printer. I.E., a 8x11" print (as JPEG) shouldn't be much larger than 3-5Mb (mileage may vary).
We used ILford PQ Universal with Ilford EM film then switched to 4489 and D19. When you develop 4489 with PQ Universal do you dilute 1+9 and develop for 4min, rinse 1min and fix for 4min (in Ilford Hypam)?
Dave
On Thu, 20 Nov 2003 08:47:31 +0000 Ken Blight {Ken.Blight-at-cancer.org.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 } ------------------------------------------------------------------------------- } } Like Many others we suffered from the change in the } formulation of our EM film. We tried many alternative methods of } reducing the fogging and by far the best, and the easiest was } changing to Ilford PQ universal developer. We don't even have to } agitate to get smooth, repeatable results! } Ken Blight } Senior Scientific Officer } Cancer Research UK } London } } } } 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"
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 09:45:00 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (osakhi2003-at-yahoo.co.in) from on Thursday, November 20, 2003 at 08:58:48 ---------------------------------------------------------------------------
Hello Sousan and Steve and all, Let me answer your questions.
We have a LaB6 Filament.
The Anti-Contamination Device (ACD) is a LN2 cooled metal (no charcoal) cryo pump that pretty much surrounds the sample. It accumulates molecules in this vicinity, most water vapor I am sure. When the ACD Heat (regeneration) cycle is initiated, the ion pump is valved off and the diffusion pump valve is opened to pump the molecules that are desorbed as the ACD is warmed up. The ion pump can not keep up with the gas load so if the ACD cycle is not run, the column pressure increases. I am not sure how high it goes but 10-4 Torr is probably not a unreasonable guestamation.
Yes we have a maintenance contract which includes bi-annual service.
Steve Chapman, Thank you, as you have probably seen provided a very detailed response. The obvious interpretation is that I trashed our gun. The explanation of the plasma effect is opposite is that I was thinking, that being that atomic hydrogen and oxygen from the ionized water would lean up existing carbon as in plasma etching with air and carbon cleaning experiments using H and O that I am aware of here at Rice. Ok so I am wrong. 100kV would tend to change things a bit :).
Thank you, Bruce
Steve Chapman wrote:
} Hi } } I am not familiar with the ACD you mention but am very familiar with } electron guns so lets have a look at your problem? } } Contamination in the gun area may usually be attributed to vacuum levels and } the resulting high voltage discharge. Spoil the vacuum and you reduce the } insulating effect such that high voltage discharge occurs and this cracks } gasses that deposit around the gun chamber as a brown/orange layer. This } layer seems to be very absorbent and acts like a gas sponge when you re } apply a vacuum. The constant outgassing of course leading to even more } contamination of the gun chamber. To get back to normal ALL this } contamination must be removed } } Within the cathode assembly the filament and its base will also give an } indication of poor vacuum or poor quality filaments. I have set this out in } detail on our interactive EM maintenance CD but this is a brief resume of } what you will see. } } 1. The filament will have suffered from gas attack resulting in extreme } thinning over the complete filament, not just at the tip as you would see in } a normal failure. } 2. The filament base will also become discoloured taking on the same } orange/brown tint as the gun chamber. } 3. Poor quality filaments that are made up of a high percentage of } rubbish will very rapidly contaminate the cathode assembly as the rubbish } evaporates. The filament break is normal but the life and the base will } indicate problems; the base will be orange/brown. } } I have worked with almost every type/make of EM and from time to time the } manufacturers have a batch of filaments that are, through no fault of their } own, of poor quality. Short life high contamination are instant } indicators. I have probably mentioned before that if you have a good box of } filaments NEVER use all of them, save two. In a situation where you believe } you have a filament problem you may then go back to your good box and } confirm that they run correctly. Pop the remainder of the bad box back to } your supplier whom I am sure will provide you with a new box. } } Good luck with your problem, is it a poor filament problem have you just } opened a new box? } } Steve Chapman } Senior Consultant Protrain } Electron Microscopy Training and Consultancy World Wide } Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Mob 07711 606967 } www.emcourses.com } } ----- Original Message ----- } From: "Bruce Brinson" {brinson-at-rice.edu} } To: {microscopy-at-msa.microscopy.com} } Sent: Wednesday, November 19, 2003 6:00 PM } Subject: [Microscopy] TEM, GUN? } } } } } } } -------------------------------------------------------------------------- } ---- } } 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 } } -------------------------------------------------------------------------- } ----- } } } } Bruce Brinson wrote: } } } } } Hello Microscopist, Ladies and gents, } } } } } } Let me set the stage, no pun intended. We have a Jeol 2010. I } } } managed and trained students on this instrument from 1993 to some time } } } } } } in 2000. During that time I had to tongue lash, in a nice way, users } } } for } } } not activating the ACD cycle at the end of the day. This happened ~ 20 } } } } } } times with the only failing being a bellows in the gun valve after ~ } } } 10,000 cycles. This will make since to you if you have a 2010 with the } } } } } } old gun valve design. } } } I still on occasion use the instrument. Please understand, I need } } } to } } } refer to specifics of day. This in not intended to be a assault on } } } Jeol. I am the culprit. } } } I logged onto the instrument a few days after an new filament was } } } installed. I found the image of the filament to be moving back and } } } forth, ~1 Hz, ~1mm at x100K.... This I interpreted as an instrument } } } charging } } } problem. I continued to survey samples, encountering 2 more technical } } } annoyances followed by a 4th that brought the session to an end. We } } } all } } } know these things happen. } } } Perhaps more frustrated than I realize, I left the instrument } } } without activating the ACD and with the HT at 100 kV. My mistake was } } } found the following morning. } } } It's been implied that my mistake caused a contamination of the } } } gun } } } so severe that 2 additional filaments were contaminated by it as fast } } } as they were installed. This I } } } find hard to believe but I suppose stranger things have happened and } } } what do I know. } } } I'll not share other thoughts or the opinions of others for now } } } least I } } } bias your response. } } } } } } Well what do you think? } } } 1. has anyone had this happen to their instrument? } } } 2. Has anyone ever heard of a gun contaminating a filament to this } } } extent?. } } } 3. other comments? } } } } } } } } } thanks, } } } Bruce Brinson } } } Rice U. } } } } } } } } } } } }
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 10:21:17 2003
I have developed the "new" 4489 with no problems using the same technique I used with the old film. I have never have problems with uneven development.
1. I make my D-19 from raw ingredients using Kodak's recipe. I don't use prepackaged D-19 because I do so little EM these days I wind up throwing most of it away. The prepackaged stuff only keeps 2 years anyway. 2. Use fresh developer! Stuff that has been sitting in a tank without a floating lid goes bad in just 2-3 days, even if it is covered. Even with a floating lid I never use developer that is more than 5 days old. Chemisty is the cheapest part of the process, a few sheets of film costs as much as all of the chemicals in D-19. 3. I agitate with the "tilt" method but more slowly, it takes me 8-10 seconds to remove-tilt left-reimmerse-tilt right-reimmerse. I do this every 30 seconds. 4. Rinse and fix as usual.
Geoff
Ken Blight 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 } ------------------------------------------------------------------------------- } } } Like Many others we suffered from the change in the formulation of } our EM film. We tried many alternative methods of reducing the } fogging and by far the best, and the easiest was changing to Ilford PQ } universal developer. We don't even have to agitate to get smooth, } repeatable results! } Ken Blight } Senior Scientific Officer } Cancer Research UK } London }
-- -- ********************************************** 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 Nov 20 11:48:49 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (rbremer-at-acpub.duke.du) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Thursday, November 20, 2003 at 11:31:41 ---------------------------------------------------------------------------
Email: rbremer-at-acpub.duke.du Name: Ron
Organization: Duke
Education: Graduate College
Location: Durham. NC
Question: Hey,
I work with an Olympus BH2 which for we recently got a blue filter set (filters both excitation and emission). When I view my specimens with my 20x objective I get a very nice signal over the microscopeís black background. If I use any of my other objectives (at least 10-40x), I get a blue haze over the whole field (that is even with no slide/specimen on the stage). I imagine this has to be a problem with the objectives, but since i've not done much blue fluorescence in the past I thouhgt I would check up on it a bit. All the objectives work fine when using red or green IR. Thoughts?
Gram-positive bacteria have relatively simple cell walls with a relatively high peptidoglycan content. Gram-negative bacteria have a more complex cell membrane system with relatively low peptidiglycan content. The crystal violet stain in the first step of Gram staining will bind to the peptidoglycan layer in Gram-positive cells effectively masking membrane counterstaining by safrannin. Gram-negative cells will bind relatively little crystal violet in the thin peptidoglycan layer, the rest being washed out in the decolorizing step. Cell membranes stained by safrannin in the last step will appear red in Gram-negatives. Hope this helps Robert Simmons
Dr Robert Simmons Program Director Biological Imaging Core Laboratory Georgia State University Atlanta, GA 30302-4010
404-651-3138 404-651-2509 FAX
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 12:21:49 2003
In reply to your request for more information on the high voltage tank. Yes you are correct it can be a messy task!
Disconnect all cables and move the tank to an area that allows plenty of space to work. Run the tank onto an expanse of plastic sheet at least nine times the size of the tank, i.e. plenty of floor space covered. The area selected should be as clean as possible with no through way for persons not involved with the task. The tank is built with all the electronics suspended from the top plate. Give the tank outsides a good clean and then remove the bolts holding top plate to tank. If you have a crane it helps but I have carried out this procedure with some willing helpers! Tank fluid ruins shoes so we always work bare footed with trousers rolled up! Get a picture of this?
Provided the HT has been off overnight you should have no residual charge problems. Very slowly lift the tank top allowing the fluid to drain back into the tank. Take your time here. Once clear of the tank and well drained you may place this unit on the protected floor but it must be on a CLEAN surface.
Its a long time since I went into the tank, but from memory if you have the circuit diagrams you should not have a problem finding the appropriate board. I assume you know how to check the diodes, lets hope there lies the fault?
Once fixed make sure all under surfaces are clean before you very very slowly lower the tank top back into place. The air needs to be driven out of the components so very gently rock the top as you lower it into place. Once in place and bolted down once again rock the tank but you may now be a little more violent.
Run the HT up very slowly, stay on the lowest kV for at least half an hour for the tank to warm, convection currents should ensue that no gas is trapped. Lets hope you are able to fix the beast?
Best of luck
Steve Chapman Senior Consultant Protrain Electron Microscopy Training and Consultancy World Wide Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Mob 07711 606967 www.emcourses.com
----- Original Message ----- } From: "John Brealey" {john.brealey-at-imvs.sa.gov.au} To: "Listserver" {Microscopy-at-MSA.Microscopy.Com} Sent: Thursday, November 20, 2003 6:27 AM
It was recently brought to my attention that a few members of the Microbeam Analysis Society (not Microscopy Society of America) did not receive their 2004 renewal membership forms and election ballots sent out a few weeks ago. If you are a current MAS (not MSA) member and did not receive this mailing, please contact me (rosslm-at-missouri.edu) and I will have it sent out to you.
We apologize for this unfortunate problem . Lou Ross MAS Membership Services -- 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 web: www.biotech.missouri.edu/emc
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 20 16:20:45 2003
Hi all, I'm giving up on Google: Does anyone know where I can get a cloth dust covers that would fit a RMC 6000XL ultramicrotome? thanks for any help, Beth -- ********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
Phone - (706) 542-1790 & FAX - (706) 542-1805
"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)
In a message dated 11/20/2003 5:28:29 PM Eastern Standard Time, beth-at-plantbio.uga.edu writes:
} Hi all, } I'm giving up on Google: } Does anyone know where I can get a cloth dust covers that would fit a } RMC 6000XL ultramicrotome? } thanks for any help, } Beth } -- } ************************************************************ } ********** } Beth Richardson } EM Lab Coordinator } Plant Biology Department } University of Georgia } Athens, GA 30602-7271 } } Phone - (706) 542-1790 & FAX - (706) 542-1805
Hi Beth,
I'll check on this tomorrow for you. Don't know if we have any in stock, but I think a dust cover for an MT-7 will work also. I'll let you know what I find.
Best,
Bob Chiovetti
Robert (Bob) Chiovetti, Ph.D. Senior Product Manager Boeckeler Instruments, Inc. 4650 South Butterfield Drive Tucson, AZ 85714 USA Tel. 520-745-0001 Fax 520-745-0004 www.rmcproducts.com www.boeckeler.com bob-at-boeckeler.com
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 21 05:02:36 2003
Hello all, Does anyone know of a stain for araldite resin to stain it more grey or white under the SEM ? I'm investigating little pieces of charcoal with a XL30SFEG and have some problems with the black background caused by the resin.
TIA, Erwin Vermeij Forensic Scientist Microtraces Netherlands Forensic Institute Rijswijk, The Netherlands
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 21 08:29:00 2003
Hi all, I am looking for a new inkjet printer and an considering the Epson Stylus Photo R800 due out in February, 2004. I was also considering using only continuous tone inks since this printer will be for EM prints only.
What I am looking for is a system that will really replace a good darkroom print. I want a true grayscale without the color toning or cast that you get with all inkjet printers that have color cartridges. I also want to maximize the shades of gray to capture fine nuances in images.
Does anyone have personal experience with the Piezography BW ink systems (www.piezography.com)? Which ink tones do you prefer, what paper do you use, and can you give me an idea of cost per 8x10 page? Also how many pages per cartridge set or is it better to bite the bullet and get the continuous feed system?
Thanks, 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
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 21 09:06:57 2003
If you don't mind "staining" before the fact, I had a similar problem examining coal in epoxy many years ago. Our solution was to dissolve iodaform in the epoxy resin before mixing with the hardener. We could dissolve up to about 18% iodaform by weight by using mild heating to speed the process. I think the iodaform would precipitate out if we tried to push it further. We were trying to do automated analyses and they worked reasonably well at that level of doping. If you just need to find particles by eye, then lower levels should be adequate.
For the record, this was a simple two-component epoxy such as is sold by Leco, Buehler and others. It seemed to reduce the need for hardener. Since it adds mass to the resin, you will need to recalculate the resin to hardener ratio. You should also experiment with the mix before adding your important samples. We lost some due to the severe exotherm of the new mixture.
Depending on the size of your charcoal, you might be able to add some filler to the araldite to help locate the particles. I just received a sample of Ni powder from Buehler intended to make the epoxy conductive. I have not tried it so I don't know what particle sizes are present.
Warren
At 05:06 AM 11/21/2003, you wrote:
} Hello all, } Does anyone know of a stain for araldite resin to stain it } more grey or white under the SEM ? I'm investigating little } pieces of charcoal with a XL30SFEG and have some problems } with the black background caused by the resin. } } } TIA, } Erwin Vermeij } Forensic Scientist Microtraces } Netherlands Forensic Institute } Rijswijk, The Netherlands
------------------------------------------- No files should be attached to this message ------------------------------------------- Warren E. Straszheim, Ph.D. Materials Analysis and Research Lab Iowa State University 46 Town Engineering Ames IA, 50011-3232
While evaluating the latest in digital SEM technology, I notice a few manufacturers are still specifying the size of their frame stores, and one even claims acquiring 2560x1920 with a 1280x960 frames store. My 1st question might be how is this accomplished?, ... but my primary question would be, with modern computer interface speed and RAM, why is a frame store even necessary?
tia & cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 21 09:54:58 2003
} } } -------------------------------------------------------------------------- ---- } 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 Friends, } } A user of the lab has asked me to separate urban myth from reality } regarding HBO bulbs used for fluorescence microscopy. } } His questions are: } } What is the difference between a 50W bulb and a 100W bulb? He says he set } up a demo, identical except one scope had a 50W bulb, the other a 100W } bulb, and he didn't think he could tell much difference. } } Do bulbs have a finite life, ie., should they be changed at some maximum } hours even if still lighting up and appearing OK? He has heard various } advice, like change after 200 hours or 250 hours. Change when the glass } gets dark, how dark? Is it true that bulbs with many hours may explode?
The HBO 100w lamps put out more light. There are issues of lamp alignment and condensing lens choices for the lamphouse which can have a dramatic influence on the amount of light that reaches the specimen. If your fluorophore is bright you may be able to use the 50 watt lamp and not notice much difference between the two. If there is not much fluorescence you may not be able to see it at all with a 50 watt lamp.
The 100 watt lamp is much more stable then the 50 watt lamp. There is nothing more irritating then using a microscope with the light intensity flickering wildly. The 100 watt lamp is not likely to do this.
The rated life of the lamps is 100 hours for the HBO 50w AC and 200 hours for the HBO 100. Osram says that you can safely exceed the rated life by 20% without danger of bulb explosion. The bulb will explode if run too long, based on several factors. The rated life is determined by the factory to be the point at which the light output has been reduced by 30% from the initial output of a new lamp. A lamp driven by a properly operating power supply should look new when you change it based on maximum allowable hours. How dark it is should not be a criteria. Some people run their lamps much longer then is safe. They may have power supplies which help them do this, they may get lucky... If not, the damage done inside the lamphouse by an exploding lamp will cost $300-500 to repair. You can purchase a lot of lamps for that much money and also avoid the dubious pleasure of breathing poisonous mercury fumes. These lamps have significant amounts of mercury in them, please dispose of them properly.
The lamps cost the same amount, but the life is different. You should be paying under $100 US for them. Shop around. It seems like quite a bargain to get twice the power of the 100w, for half the cost of a 50w (based on lamp life).
We have had the best luck with Osram lightbulbs based on the 3,000 or so we have purchased over the last 20 years, and have found no difference in price which would warrant using another brand. In general we have found no significant differences in performance either.
The lamp must be run for 20 minutes at a minimum every time it is turned on or it may be ruined. This is the only use issue with this type of low wattage mercury lamp. The newer power supplies are different then the old ones some people remember and so stories of what "used" to work are of little help today. The newer power supplies will run every lamp to its rated life if the power supply is working correctly. We see lots of power supply problems which cause short lamp life. HBO power supplies seem to be problematic for some reason. If you purchase a new system and experience short lamp life I would have the supplier of your system install and align a new lamp for you (showing you the proper way to install it). If the lamp still does not reach its rated life, insist that they swap out a known, good power supply for you to try. In my experience that invariably fixes short lamp life problems...
Good luck, David Burton Optical Specialist University of Washington
} } He has only used Osram bulbs, are there other brands available and what } criteria would one use to select one brand over another? } } 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 Fri Nov 21 13:26:36 2003
Before you buy a printer make sure the piezography (or MIS or Lyson) system will work with it. Several leading printer manufacturers have put proprietary chips on their ink cartridges so you are forced to buy their cartridges. In the past it has taken several months (at least) for the folks a piezography to 'engineer' a way around this. All of the inkjet black and white prints (piezography and others) I have seen have been on non-glossy paper so I don't know how good the final result will be for traditional EM work. From what I read in the photography press, talking to users and what I have seen a exhibts, there are a lot of variables (dye-based or pigment-based ink, paper, calibrating the monitor so what you see is what you get, etc.) to work out before getting photo-quality B&W prints that do not have a color cast. Study prints are another matter. If you are going to make a lot of prints a continuous ink system will save a lot of money in the long run. Have you ever worked out the cost of cartridge ink on a per ounce basis? Shocking! How about sending piezography a jpeg or tiff image and see what they can do for you before spending your money? They could probably answer your cost questions as well. By the way, I get very nice glossy B&W prints from my Epson stylus 1280 if I check the box that allows me to print without color ink. As much as I love a real silver print, inkjet printing is very fast and convenient. Since most journals are moving to digital submission of figures (or have already done so) I don't think the longevity issues with dye-based inks will be a problem.
Geoff
Debby Sherman wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-- -- ********************************************** 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 Fri Nov 21 14:16:40 2003
If not a frame store, then what option would be used?
The idea of a frame store, I believe, is that once an image is captured in the store, it can be annotated, measured and then saved to disk. Direct capture does not allow in-SEM annotation.
The other factor to consider is that most of the frame stores are the same aspect ratio of the Polaroid 4x5 film.
gary g.
At 07:39 AM 11/21/2003, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 21 15:26:01 2003
We're having a difficulties imaging CFP expressing live cells with a 405 nm laser because of autofluorescence, unhappiness with light and corrections (or lack of) in the objectives.
Has anybody tried the Pointsource 440 nm laser for CFP or have other suggestions for a laser source ideally in the 430 to 436 nm range?
Thanks.
____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 21 15:36:37 2003
Warren's suggestion to use iodoform is a good one.
Regarding his comment on commercial conductive embedding media. The problem with the conductive embedding media that I have used and/or read about is that they are have only macroscopic electrical conductivity. This means that the material will conduct a current but, on a microscopic scale, will contain expanses of nonconductive resin that will charge under the electron beam.
My suggestions are to use polymers that contain high loadings (~50 w%) of carbon black or other conductive particle with small particle size. I presented a paper at the 2003 M&M conference (see proceedings) on the subject. The approaches that I have used are (1) carbon black-filled epoxy resins and (2) carbon black-filled thermoplastics. Regarding the choice of medium, the epoxy component of the first material is available to infiltrate into porous samples. The higher viscosity, low crystallinity plastic component of the second preparation is unable to infiltrate into microporous samples and encapsulates the sample rather than infiltrating into it. One advantage of this prep is that a porous sample is not adulterated by the infiltration of an epoxy resin into it. An advantage of the first technique is that epoxy infiltration into the sample may allow microtomy of ultrathin sections for TEM.
Hope that this provides some guidance in finding a better preparation. Feel free to write off-line for more details.
Cheers,
Gary M. Brown ExxonMobil Chemical Company Baytown Technology & Engineering - West 5200 Bayway Drive Baytown, Texas 77520-2101 phone: (281) 834-2387 fax: (281) 834-2395 e-mail: Gary.M.Brown-at-ExxonMobil.com
Warren E Straszheim {wesaia-at-iastate.edu} To: Microscopy-at-msa.microscopy.com cc: Erwin Vermeij {e.vermeij-at-nfi.minjus.nl} Subject: [Microscopy] Re: Stain for araldite resin 11/21/03 09:13 AM
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If you don't mind "staining" before the fact, I had a similar problem examining coal in epoxy many years ago. Our solution was to dissolve iodaform in the epoxy resin before mixing with the hardener. We could dissolve up to about 18% iodaform by weight by using mild heating to speed the process. I think the iodaform would precipitate out if we tried to push
it further. We were trying to do automated analyses and they worked reasonably well at that level of doping. If you just need to find particles
by eye, then lower levels should be adequate.
For the record, this was a simple two-component epoxy such as is sold by Leco, Buehler and others. It seemed to reduce the need for hardener. Since it adds mass to the resin, you will need to recalculate the resin to hardener ratio. You should also experiment with the mix before adding your important samples. We lost some due to the severe exotherm of the new mixture.
Depending on the size of your charcoal, you might be able to add some filler to the araldite to help locate the particles. I just received a sample of Ni powder from Buehler intended to make the epoxy conductive. I have not tried it so I don't know what particle sizes are present.
Warren
At 05:06 AM 11/21/2003, you wrote:
} Hello all, } Does anyone know of a stain for araldite resin to stain it } more grey or white under the SEM ? I'm investigating little } pieces of charcoal with a XL30SFEG and have some problems } with the black background caused by the resin. } } } TIA, } Erwin Vermeij } Forensic Scientist Microtraces } Netherlands Forensic Institute } Rijswijk, The Netherlands
------------------------------------------- No files should be attached to this message ------------------------------------------- Warren E. Straszheim, Ph.D. Materials Analysis and Research Lab Iowa State University 46 Town Engineering Ames IA, 50011-3232
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (david.audette-at-sylvania.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, November 21, 2003 at 15:31:00 ---------------------------------------------------------------------------
Email: david.audette-at-sylvania.com Name: Dave Audette
Organization: OSRAM Sylvania
Title-Subject: [Microscopy] MListserver:
Question: I have an EDS dewar (Kevex) system which is not holding liquid nitrogen as long as it use to and was planning on warming it up and pumping on the inside chamber via the external valve. My question: Is there a, hopefully simple, way to regenerate the zeolite that I have heard is in side this chamber? Or is it not worth that much trouble?
I am also a high school teacher. I am not an expert, I lurk on this list to get ideas. I think I can answer this question if you haven't gotten other answers.
Do you have any more questions about viewing skin cells? They are pretty easy to spot after you figure out what's there! Using Methylene Blue, the nuclei should be stained. You can use Iodine (even Iodine soap works fine), and in this case the nuclei are stained brown.
Skin cells are pancake like, very thin and broad---which makes sense if you think about what they are. They are transparent, which is one reason to use stain.
You could try oblique illumination, if you have the same school microscopes that we have: the disk diaphragm is the circular disk underneath the stage. If not, you have an iris diaphram---better, but the disk works better for us for oblique illumination. You can darken the background to some extent by tweaking the disk to one side of one of the click stops, just a little bit. On our scopes, number 2 works best, on the disk, don't know what you have. The light that passes through the specimen is directed to the side of the objective, so the background is dark; but any light that is refracted or diffracted through certain parts of the specimens towards the objective now gives the appearance of a bright subject against the dark background. This works for amoebas too, and small plankton. It's a good way to see movement in a sample of water.
Good luck. Please don't hesitate to ask any more questions. The people on this list are experts, I'm not, but as a teacher like yourself, who, by the way, is a keen amateur microscopist, I am happy to help you. I use the microscope a lot in my biology classes. The students love it.
Alan Davis Marianas High School Saipan, N. Mariana Islands
On Mon, 17 Nov 2003 14:28:05 -0600 Donnarae48-at-AOL.com (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 (Donnarae48-at-AOL.com) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Sunday, } November 16, 2003 at 14:43:02 } --------------------------------------------------------------------- } ------ } } Email: Donnarae48-at-AOL.com } Name: Donna Morrow } } Organization: Moreau Catholic High School } } Education: 9-12th Grade High School } } Location: Hayward, California } } Question: My biology class did a lab using the microscopes. We are } able to magnify up to 400 magnification. We used a scraping from our } cheeks and dye to make a slide. We were asked to draw what we saw } and lable at least 2 things. I am not sure what I saw. At 400 } magnification how clear would an epithelial cell be and would I have } been able to see a nucleus. All I saw were blue circles and light } blue background and very small round things scattered around. I am } wondering if the larger dark blue circles were a nucleus and the } smaller specks the cell membranes even though they did not seem to be } uniform. The teacher said my slide looked good but I am not sure } what I was viewing. Also do you know of any other sight that is } available to help me with my lab work with the microscope. We have } been trying to find a sight that would show me what an epithelial } cell would look like at 400 magnification and I keep running in to } brick walls. Thank you for your help } } --------------------------------------------------------------------- } ------ } }
-- adavis-at-saipan.com 1-670-235-6580 Alan E. Davis, P. O. Box 506164 CK, Saipan, MP 96950, NMI
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From MicroscopyL-request-at-ns.microscopy.com Sat Nov 22 19:32:26 2003
Dave, Certainly, in part, it depends on what you're paying for LN2, but if the vacuum has deteriorated, then it will also aggravate any oil condensation on the detector snout. I haven't done this myself, but I've been told that if you put hot water in the dewar and use a hot air gun on the outside, you should be able to drive the water out of the zeolite. Room temperature takes care of the adsorbed gasses. Hopefully, someone who has done this can also respond. Ken Converse owner Quality Images third party SEM service Delta, PA
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From MicroscopyL-request-at-ns.microscopy.com Sun Nov 23 11:37:48 2003
During some testing and debugging today I inadvertently sent out a number of test messages, some of you will see a handful of test messages, which were not suppose to reach the world but were supposed to be "trapped" locally.
Please excuse the slip up.
Nestor Your Friendly Neighborhood SysOp
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 24 08:39:07 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (oneild-at-hfx.nrc.ca) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 24, 2003 at 08:26:14 ---------------------------------------------------------------------------
Question: Does anyone have any experience with the - Cressington 108 Auto or the Polaron/Quorum SC7620 "Mini" - Sputter Coaters that they are willing to share? Thanks
David O'Neil Institute for Marine Biosciences National Research Council of Canada 1411 Oxford St. Halifax, NS B3H 3Z1 ph. 902-426-8258 fax 902-426-9413 david.o'neil-at-nrc.ca http://www.imb.nrc.ca/techdev/microscopyfac_e.html
I am looking for protocols on how to prepare mushroom spores for scanning. I am very much interested in the practical side of things, e.g. in what to put the spores to pass them through the different solutions and critical point drying. All tips & tricks are welcome.
We have a researcher who needs to select and tack 2 to 5 micron particles on to a slide in a small spot over a 30 minute period, then put epoxy on the top, polymerize, polish, and analyze with SIMS. We need something that will stay sticky on the slide for the 30 minutes. I am not familiar with the methods for this. We've had nitrocellulose suggested, as well as B72. Would double sided sticky tape work? They must all be in the same plane as much as possible. Any suggestions? Many thanks, Nancy
Nancy Piatczyc Bronfman Science Center Williams College Williamstown, Ma
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 24 16:31:18 2003
Nancy Piatczyc wrote: =================================================== We have a researcher who needs to select and tack 2 to 5 micron particles on to a slide in a small spot over a 30 minute period, then put epoxy on the top, polymerize, polish, and analyze with SIMS. We need something that will stay sticky on the slide for the 30 minutes. I am not familiar with the methods for this. We've had nitrocellulose suggested, as well as B72. Would double sided sticky tape work? They must all be in the same plane as much as possible. Any suggestions? Many thanks, Nancy ===================================================== This sounds like the perfect application for the SPI Tacky Dot™ Slides, see URL http://www.2spi.com/catalog/new/tacky.shtml
You would use for the mounting apparatus, the Circle Analytical Mount which would align your particles within the two circular areas and then using the Teflon® Block Mount, embed the mounted particles, right on the slide with your peferred resin.
When the resin is cured, it will slide out of the Teflon mount and the particles will be perfectly aligned and parallel with the flat face of the plastic mount, which can then be polished down exactly as you have indicated The final result of that kind of cross-sectional polishing can be seen on URL http://www.2spi.com/catalog/new/tackdot_array.html
For particles in the indicated size range, I would suggest a 15 um size for the Tacky Dot Slides.
Disclaimer: SPI Supplies manufactures Tacky Dot Slide arrays and therefore has a vested interest in seeing them used by more workers.....
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 Mon Nov 24 17:39:01 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tgruber-at-phelpsdodge.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 24, 2003 at 13:12:47 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] MListserver: Accuview 789 Camera for Donation
Question: Dear Listers,
My company has a c. 1996 "Gatan Accuview Camera Model 789" available for donation to a suitable recipient. This is a 1.4 megapixel slow-scan format. It is interfaced via a camera interconnect cable to a Kodak Megaplus Camera Control Unit. The camera controller output port is 37-pin male and the output cable is provided. Camera, cables, and controller were working fine when their use was discontinued, and all will be donated together, in ìas-isî condition. This camera was formerly operated through Gatan software and a framegrabber board installed on a Macintosh computer (not available from us). The camera is c-mount and has a macro zoom lens mounted.
This donation is contingent upon my company being able to realize a charitable tax deduction relative to our depreciated book value of this equipment (c. $8,000). Non-profit institutions qualified to receive such a donation will be considered.
Eligible and interested parties may reply to me (OFF-LINE, PLEASE, at tgruber-at-phelpsdodge.com).
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (em_man_1-at-hotmail.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 24, 2003 at 17:16:13 ---------------------------------------------------------------------------
Email: em_man_1-at-hotmail.com Name: William McManus
Organization: J & M Analytical, Inc.
Title-Subject: [Microscopy] [Filtered] looking for EM equipment
Question: After 20 years, the EM facility which I managed at Utah State University has been closed. I am expanding my consulting company and looking for good used equipment. I am particularly interested in aquiring a Zeiss (LEO) 902 TEM. I am also looking for Ultramicrotomes and SEM prep equipment. If anyone has equipment that they would like to sell, please contact me. Bill 435-946-8739
Hi, I'm posting this enquiry for a gentleman who rang me today. He's acquired an LKB Nova Ultratome (circa 1985) through an auction, however, it's missing the accompanying electronics box (ie control box). Does anyone have a spare control box they are willing to part with?
Thanks,
John Brealey EM Unit Queen Elizabeth Hospital Adelaide, South Australia
(08) 8222 6612
From MicroscopyL-request-at-ns.microscopy.com Mon Nov 24 21:39:09 2003
We can writing alkanethiols onto a gold surface, by dip-pen nanolithography. Can we remove or alter the patten (like using an eraser) we had created if the sample is biological such as E-coli, cells or single peptide? If so how can we do that?
I do appreciate your kindness.
Sincerely yours,
Chris
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From MicroscopyL-request-at-ns.microscopy.com Tue Nov 25 02:16:40 2003
Thanks for the reply. I figured that this might work, but I also found this protocol:
Small pieces of the lamellae (herbarium material) were used. They were pretreated for 12h in water and ammonia. After a period of 1h in 70% ethanol, they were passed 2 x 30 min. in dimethoxymethane, before being submitted to the process of critical point-drying. The material was coated with gold (with Argon-gas, under 0.05 bar) for 3 min. 30 sec., until a layer, 15 nm in thickness, is covering the material. (The scanning electron microscopy was carried out with a JEOL 5800LV with tension of 25kVolt and a working distance op 7-8 mm.)
Can you tell me why or when one should use this protocol?
Regards,
Renaat
----- Original Message ----- } From: "Rosemary White" {rosemary.white-at-csiro.au} To: "Renaat Dasseville" {Renaat.Dasseville-at-UGent.be} Sent: Monday, November 24, 2003 11:06 PM
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (h.grenfell-at-geomarine.org.nz) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 24, 2003 at 18:21:41 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] MListserver: Any ideas?
Question: Hi, Can anyone help with info as to what these minute ?siliceous objects are? Please see images at http://homepages.ihug.co.nz/~bw.hayward/SEM%20images/
They are commonly found in estuarine samples that we are studying using foraminifera. Would be nice to know what these spheres are trying to tell us!!
They are not radiolaria and many tell us they are of sponge origin but no detail or images from the literature as yet.
Please note the final date for receipt of Abstracts for presentation at the Australian Conference for Microscopy and Microanalysis 2004 (http://www.deakin.edu.au/events/acmm18/) and for registration at the series of workshops scheduled to precede the conference is :
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Please also note that response to our call for papers has been outstanding! In order for abstracts to be considered for oral presentation we must have received them by close of business on Wednesday.
Abstracts received after this date may well not appear in the official proceedings, but form part of an addendum (if received in sufficient time).
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From MicroscopyL-request-at-ns.microscopy.com Tue Nov 25 07:51:19 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tholzheu-at-camet-lab.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 24, 2003 at 22:27:40 ---------------------------------------------------------------------------
Email: tholzheu-at-camet-lab.com Name: Thomas Holzheu
I am probably pushing my luck here, but then you nerver know. We have an EDXRF spectrometer with a kevex 8000 system and an iomega bernoulli box (scsi). Upon booting the DEC computer, the system is accessing the first drive, but is not loading the software. The issue here is, that I want to replace the bernoulli box with a superdisk drive (scsi), but just exchanging the drives is not working. I would appreciate any input and thoughts on this matter.
Hi Renaat, I would recommend not using any chemicals at all.
You can collect the Lactarius basidiospores by making a spore print: Remove the stalk. Place the cap on a piece of paper for several days (dark paper for light spore and vice versa). The spores will be discharged onto the paper. Remove the cap. Put a carbon sticky tab on a SEM stub and lightly touch the stub to the spore print.
Experiment with the spores after you mount them. Place some in a desiccator for a few days before coating them, and others try coating and scoping right away (when they are fresh like that it is harder on the scope - it takes longer to pump down).
Also, the scoping part depends on your scope. If you have a cryo-SEM then by all means use it. If you don't have the cryo attachment then coat the samples and use low kv (if possible) or scope them quickly at 10 or 15 kv so they don't collapse or look toasted.
have fun with fungi, Beth
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-- ********************************************************************** Beth Richardson EM Lab Coordinator Plant Biology Department University of Georgia Athens, GA 30602-7271
Phone - (706) 542-1790 & FAX - (706) 542-1805
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Thomas, Contact tech support for Thermo Kevex x-ray Tel: 831.461.2326 Fax: 831.438.5892 E-mail: chrisc-at-kxr.com Let them know what you are doing and ask them for tech support. Last year I had trouble with my 8000 and they put me in contact with a tech that really knowledgeable about the system. I think he is in Georgia, but they will connect you to him.
Rich L CMD Tempe
-----Original Message----- } From: by way of MicroscopyListserver [mailto:tholzheu-at-camet-lab.com] Sent: Tuesday, November 25, 2003 7:03 AM To: microscopy-at-ns.microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tholzheu-at-camet-lab.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, November 24, 2003 at 22:27:40 ---------------------------------------------------------------------------
Email: tholzheu-at-camet-lab.com Name: Thomas Holzheu
I am probably pushing my luck here, but then you nerver know. We have an EDXRF spectrometer with a kevex 8000 system and an iomega bernoulli box (scsi). Upon booting the DEC computer, the system is accessing the first drive, but is not loading the software. The issue here is, that I want to replace the bernoulli box with a superdisk drive (scsi), but just exchanging the drives is not working. I would appreciate any input and thoughts on this matter.
The deadline for all nominations for 2004 Microscopy Society of America awards is nearing (December 15, 2003). Please note the following announcement that was previously posted in October:
Call for Nomination of Individuals to be considered for Major Awards by the Microscopy Society of America.
Awards:
Distinguished Scientist Awards:
These Awards recognize preeminent senior scientists from both the Biological and Physical disciplines who have a long-standing record of achievement during their career in the field of microscopy or microanalysis..
Burton Medal:
The Burton Medal was initiated to honor the distinguished contributions to the field of microscopy and microanalysis of a scientist who is less than 40 years of age on January 1st of the award year.
Optical Imaging Association-MSA Outstanding Young Investigator Award:
This Award, initiated in 1999, recognizes the distinguished contributions in the field of optical microscopy made by a scientist who is less than 40 years of age on January 1st of the award year.
Outstanding Technologist Awards:
These Awards honor technologists from both the Biological and Physical Sciences who have made significant contributions such as the development of new techniques which have contributed to the advancement of microscopy and microanalysis.
Morton D. Maser Distinguished Service Award:
This Award was initiated to recognize outstanding volunteer service to the Society as exemplified by Mort Maser, who served the Society for many years with great dedication. This award is made to honor an MSA member who has provided significant volunteer service to the Society over a period of years.
Nomination Requirements:
The Distinguished Scientist, Burton Medal, OIA-MSA Outstanding Young Investigator and Outstanding Technologist Awards Nominations should include:
1) a letter from the primary MSA nominator describing the research accomplishments of the candidate with particular emphasis on the unique technical achievements in the Physical or Biological Sciences; and 2) supplemental letters of support from other members of the scientific community.
The Morton D. Maser Distinguished Service Award Nomination should include:
1) a letter from the primary MSA nominator describing the basis for the nomination; and 2) supplemental letters of support from other members of MSA.
The Deadline for receipt of Awards Nomination Packages is December 15, 2003.
Please contact the MSA Business Office for additional information.
Judy Janes, Administrative Manager Bostrom Corporation 230 E. Ohio Street, Suite 400 Chicago, IL 60611-3265 (800) 538-3672; Fax (312) 644-8557, jjanes-at-MSA.microscopy.org
Thanks,
William T. Gunning, Ph.D. Associate Professor of Pathology Medical College of Ohio Department of Pathology BHSB 140 3035 Arlington Avenue Toledo, Ohio 43614-5806 Phone: 419-383-5256 Fax: 419-383-3066 email: wgunning-at-mco.edu
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 26 03:34:10 2003
I have some questions regarding the quality of microplates for microscopy, mainly 96- and 384 well SBS-standard layout formats (1536 and 6144 layouts would be interesting too of course).
I have been doing some quality tests for certain types of multiwell plates which are very popular for use in "readers" with our 40x 0.75 N.A. objective and a 63x 0.8 objective in brightfield and fluorescence. These objectives have a very good spatial resolution and capture a lot of light for weak fluorescence (I ~ N.A.^4/M^2). However these objectives have a short working distance and as such the quality of the plate bottoms is important.
Our autofocus system only needs about 5 videoframes to gather enough information to find a focus level, so with a 40 millisec. frametime (PAL 25 fps.) and some time for mechanical movement, it does a complete image content based autofocus in 0.3 or 0.4 seconds (300 or 400 milliseconds). But this only is the case, when a focuslevel is found within a certain range, i.e. the travel range needed for 5 frames, which is in principle determined by the Nyquist sampling theorem for Z-slicing.
We also do a complete range check, as we want to focus in those microscopy modes which can cause multiple maxima in a focus algorithm (patented) in which case we focus on the highest peak/score found ( Geusebroek J.M., Cornelissen F., Smeulders A. W. M., and Geerts H., Robust autofocusing in microscopy, Cytometry, 36(1):1-9, 2000 ).
Using an image content based autofocus system, allows us to use very fine Z-level focusing into cells with a varying offset to the multiwell plate bottom.
I found out that when you plot the bottom profile of standard multiwell plates as a 3D landscape, these plates look more than a mountaneous region than having a flat profile.
I also found some microplate types with very thin plastic bottoms and a flat plate bottom, but they are of course a bit more expensive. Glass bottom plates are an alternative in some cases, but I heard that the glue which is used for these plate-bottoms is not compatible with some of the solvents used in farmaceutical research (DMSO, etc), so the bottoms tend to fall off the plate after 24 h. incubation, which would be rather inconvenient for our time-lapse work :-( ? So using good quality plastic bottom plates seems to be one possible solution ?
A possible solution for the variability of the plate bottoms is to do a prescan at lower mag./N.A., either with a plate bottom finding laser-system or a content-based autofocus and model the plate with a mathematical model, but this takes time.
Using Long Distance (L.D.) objectives is another possible solution, but these objectives have a lower N.A. and as such less spatial resolution, which hampers subcellular imaging and analysis. Also the reduced N.A. has a very dramatic efect on the amount of light captured by the objective (I ~ N.A.^4/M^2) and we always do our image capturing in real-time, we just use very sensitive cameras which allow imaging without the need to integrate frames. The image capturing must be done at 40 msec. (PAL 25 fps.), which is important when doing large volumes of multiwell plates in our screenings. Using the 0.75 and 0.8 N.A. objective in combination with the ultrasensitive camera allows us to dilute reagents up to 1000x, which is very good for our assay costs when doing large screens of subcellular phenomena :-)
I would like to know what is the experience of other researchers with the quality of multiwell plates on microscopes or similar (automated) readers ? Does anyone else uses high N.A. objectives on multiwell plates for screening ? As far as I understand the SBS-standards define XY-measures of multiwell plates, but leave the bottom offset and bottom thickness to the manufactureres of multiwell plates ?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (ajitsankar.mallik-at-epfl.ch) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, November 26, 2003 at 04:16:42 ---------------------------------------------------------------------------
Email: ajitsankar.mallik-at-epfl.ch Name: Ajit
Organization: EPFL .
Education: Graduate College
Location: Lausanne , Switzerland
Question: Dear Sir / Madam , 1)I want to know the best visualizing methods for Endothelial cells directly on an artery . 2)Could you please let me know the best fluroscent probes for Endothelial cells if i plan to visualize them on confocal laser scanning microscope ?(least back ground fluroscence ) 3) I would like to know the live and dead endothelial cells on an artery by microscopy . Please suggest the best method .
Postdoctoral Fellow Australian Key Centre for Microscopy and Microanalysis, The University of Sydney, Australia. Reference No. A46/004354
Applications are invited for a Postdoctoral Fellow in the Australian Key Centre for Microscopy and Microanalysis to work on the research project, "Mapping electronic structure and material properties with atomic resolution". This project will use electron energy loss spectroscopy (EELS) to map the bonding and electronic structure of InGaN quantum wells and other materials at the atomic scale. We will measure and correlate the local composition, strain and electronic structure variations within the wells in order to understand the optical emission in the system. The development of EELS spectrum imaging tools for the mapping of bonding states in materials in an important aspect of this project.
Essential criteria for successful applicants include: a PhD or equivalent qualifications in a relevant discipline; extensive experience in the operation of FEG TEM/STEM systems; experience in acquisition of EELS and EFTEM data; proven research ability; evidence of research potential; and the ability to work cooperatively with other members of the university community.
Experience in ab initio calculations of electronic structure and EELS spectra, computer programming experience and demonstration and teaching skills would be desirable.
The position is full-time fixed term for one year, subject to the completion of a satisfactory probation period for new appointees. There is the possibility of further offers of employment for up to twelve months, subject to funding and need. For further information, contact Dr Vicki J. Keast on 9351 4534, fax 9351 7682 or e-mail vicki.keast-at-emu.usyd.edu.au
Remuneration Package: $60,535 - $64,981 p.a. (which includes a base salary Level A $51,153 - $54,910 p.a., leave loading and up to 17% employer's contribution to superannuation)
Closing: 18 December 2003 Applications (five copies) should quote the reference no, address the selection criteria, and include a CV, a list of publications, the names, addresses, e-mail, fax and phone number of three confidential referees.
Forward applications to: The Personnel Officer, College of Sciences and Technology, Carslaw Building, (F07), The University of Sydney, NSW, 2006 ------------------------------------------------------------------------------ Dr. Vicki J. Keast Australian Key Centre for Microscopy and Microanalysis Madsen Building F09 University of Sydney Sydney NSW 2006 Ph: + 61 2 9351 4523 Fax: + 61 2 9351 7682 email: v.keast-at-emu.usyd.edu.au
For correspondence regarding the Australian Microscopy and Microanalysis Society, please reply to amms.secretary-at-emu.usyd.edu.au ------------------------------------------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 26 08:55:42 2003
I am trying to deliver some protein inhibitors into live worm sperm cell. The nice crawling cell has a long lamelipod and a cell body trailing behind. The lamelidod is about 30 micron in length and 10 micron in width. The tip of micropipette I used is about 1 micron in diameter. The injector I have is from Narishige and filled with mineral oil when used.
Every time when I touch the cell with the micropipette, the cells explode. Does anyone on the list have the same experience before?
Best regards,
Long
----------------------------------------------- Miao, Long Dept of Biological Science 334 Bio. Unit1 Florida State University Tallahassee, FL32306-4370 email: lmiao-at-bio.fsu.edu Voice: (850)644-9817 FAX : (850)644-0481 -----------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 26 11:52:44 2003
Hi, I have an investigator who is carrying out em immunolabeling on isolated dispersed filaments. The labeling is carried out on the grid but the filaments are not visible without negative staining. It would appear that the labeling procedures deposits considerable material on the grid which shows up as a thick layer and this obscures the filaments. Is this to be expected? Is it an inadequate washing procedure after labeling? Any thoughts?
Regards to this great resource as always
Chris
Christopher J. Gilpin Ph.D. Assistant Professor Director Molecular and Cellular Imaging Facility K1.246 Department of Cell Biology University of Texas Southwestern Medical Center 5323 Harry Hines Boulevard Dallas, Texas 75390-9039 +1 214 648 2827 Phone +1 214 648 6408 Fax christopher.gilpin-at-utsouthwestern.edu
From MicroscopyL-request-at-ns.microscopy.com Wed Nov 26 16:00:48 2003
Christopher- The solutions wouldn't ordinarily be dirty enough to deposit anything. I think he/she may be using too concentrated an antibody solution. Also, leaving the antibodies on for too long a time might accomplish the same thing.
Best of luck to you! Carol Heckman (Bowling Green State University)
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From MicroscopyL-request-at-ns.microscopy.com Wed Nov 26 19:06:45 2003
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jtd1-at-psu.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, November 26, 2003 at 08:56:20 ---------------------------------------------------------------------------
Email: jtd1-at-psu.edu Name: Tom Doman
Organization: Animal diagnostic Laboratory, Penn State University
Question: We are seeking to purchase a scanner to digitize 31/4" X 4" TEM negatives; the unit will be used as a dedicated film scanner for this format. The budget for the purchase is up to $700. We have selceted a Umax PowerLook 1000. Does anyone have any experinece with this scanner?
Chris In general, it's bad idea to do labelling on the grid. If you don't have very special reasons to do so, do it in vitro before grids. For good results, molar antigen-antibody ratio should be around 1:1, so you need to perform a serial dilution 1:2; 1:4; 1:8; 1:16 and so on to find optimal one. For negative staining, antibody should be homogeneous - IgG fraction preferably. It's also helpful to remove excess of material by gel-filtration etc... In general: for good consistent negative staining you need BIOCHEMICALLY (at least) homogeneous sample. If it's filaments, it should be filaments, not cell lysate. If it's specific IgGs - it should be IgGs, not serum, etc. Sergey
At 12:03 PM 11/26/2003 -0600, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Thu Nov 27 03:12:34 2003
Hello, I use to do immunolabelling followed by negative staining on virus particles and do not have this kind of problem. You can see the procedure I follow in this paper : Spehner, D., R. Drillien, F. Proamer, C. Houssais-Pecheur, M. A. Zanta, M. Geist, K.Dott, and J. M. Balloul. 2000. Enveloped virus is the major virus form produced during productive infection with the modified vaccinia virus Ankara strain. Virology 273:9-15 Don't hesitate if you wish more informations good luck danièle
-----Message d'origine----- De : Christopher Gilpin [mailto:christopher.gilpin-at-utsouthwestern.edu] Envoyé : mercredi 26 novembre 2003 19:04 À : Microscopy-at-MSA.Microscopy.Com Objet : Immunolabeling and negative staining
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Hi, I have an investigator who is carrying out em immunolabeling on isolated dispersed filaments. The labeling is carried out on the grid but the filaments are not visible without negative staining. It would appear that the labeling procedures deposits considerable material on the grid which shows up as a thick layer and this obscures the filaments. Is this to be expected? Is it an inadequate washing procedure after labeling? Any thoughts?
Regards to this great resource as always
Chris
Christopher J. Gilpin Ph.D. Assistant Professor Director Molecular and Cellular Imaging Facility K1.246 Department of Cell Biology University of Texas Southwestern Medical Center 5323 Harry Hines Boulevard Dallas, Texas 75390-9039 +1 214 648 2827 Phone +1 214 648 6408 Fax christopher.gilpin-at-utsouthwestern.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Nov 27 08:52:46 2003
We are looking for a micromanipulator to handle FIB manufactured TEM samples. We have references from Narishige (distributes in the USA by Tritech Research) but we need a europeen distributor.
By the way, advices from people with some experience in this domain are welcome.
Answer directly to Jacques.Werckmann-at-ipcms.u-strasbg.fr
Thanks
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
Dear all, Does anyone know much about low-loss EELS spectra of gold or silver. I am looking for information on bulk plasmon and surface plasmon energies and energy loss spectra of the region to about 30 or 40 eV I already tried the EELS database at CEMES in France (http://www.cemes.fr/~eelsdb/) and searching the literature but didn't find much. This is to try and explain spectra and energy filtered images of gold or silver nanoparticles.
Thanks for any clues you can give.
Best wishes
-- Ian MacLaren Technische Universität Darmstadt Material- und Geowissenschaften Petersenstr. 23 64287 Darmstadt Germany http://www.tu-darmstadt.de/fb/ms/fg/sf/projekte/maclaren-Dateien/maclaren.html
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 28 02:07:57 2003
have you tried the EELS Atlas by Ahn and Krivanek, published by Gatan in 1983. (Copies available from Gatan or from the Arizona State University. I can give you adresses.)
Old publications: M. Creuzburg (1966), Zeitschrift fuer Physik 196, 433-463 H. Froehlich and H. Pelzer (1955), Proc. of the Physical Society A, 68, 6, 525 -529
Philip
Philip Koeck Svdertvrns Hvgskola and Karolinska Institutet Dept. of Bioscience at Novum S-14157 Huddinge Sweden phone: +46-8-6089186 fax: +46-8-6089290 http://www.biosci.ki.se/em
The phrase 'We have always done things this way.' is as much a reason to change as a reason not to. - Dartwill Aquila _______________________________________
----- Original Message ----- } From: "Ian MacLaren" {maclaren-at-tu-darmstadt.de} To: "Microscopy Listserver" {Microscopy-at-sparc5.microscopy.com} Cc: "Rik Brydson" {mtlrmdb-at-leeds.ac.uk} ; "Alan Craven" {a.craven-at-physics.gla.ac.uk} Sent: Thursday, November 27, 2003 4:55 PM
Dear all, Does anyone know much about low-loss EELS spectra of gold or silver. I am looking for information on bulk plasmon and surface plasmon energies and energy loss spectra of the region to about 30 or 40 eV I already tried the EELS database at CEMES in France (http://www.cemes.fr/~eelsdb/) and searching the literature but didn't find much. This is to try and explain spectra and energy filtered images of gold or silver nanoparticles.
Thanks for any clues you can give.
Best wishes
-- Ian MacLaren Technische Universitdt Darmstadt Material- und Geowissenschaften Petersenstr. 23 64287 Darmstadt Germany http://www.tu-darmstadt.de/fb/ms/fg/sf/projekte/maclaren-Dateien/maclaren.ht ml
From MicroscopyL-request-at-ns.microscopy.com Fri Nov 28 15:49:12 2003
Our company is considering buying an automatized fluorescence microscope (motorized fluorescence filter wheel, bright field illuminator, nosepiece, condenser, and equipped with a motorized stage).
Considering the quality of optics and reliability of motorized pieces (precision, durability), is there a manufacturer that would be recommended?
Thank you.
Marie-Claude Bélanger
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