Do you have access to an ion-beam coater? I should think there would be one in your neck of the woods -- Ohio State, or Wright Patterson AFB, if there isn't one in Cleveland. We routinely use one with a platinum target for high resolution SEM of subcellular structures, gels, etc., and I don't coating effects until 300 - 400,000X normally. Depending on thickness -- normally we use 1 - 2 nm. Phil
} I am trying to look at a native silica aerogel in a Hitachi S4700 } FE-SEM. I want to look at a fractured specimen in a 100 to 200 } nanometer range. The problem seems to be optimal sample preparation. } I can view the sample with a gold or gold/palladium sputter } coat but in order to view at the magnification I'm looking for the } coating is substantial enough that it fills in some of the pores or } open pathways and totally coats over the primary silica particles. } I have a comparison of gold sputtered onto a glass slide with a } Balzer Coater, 40 ma and 1.5 inches to the top of the jar at 10, 20 } and 30 seconds. The ten second sputter coating viewed by SEM at } x250k and 200nm somewhat mimics photographically the channels and } porosity of the native silica. At the higher sputter times of 20 and } 30 seconds the pores and channels begin to close or fill in. } I have also tried a carbon deposition coating. It's lighter } in density not filling in pores and channels and not obliterating } primary particles but doesn't seem to be conductive enough to } dissipate charging in a manner that lends itself to anything but low } magnification photos. I've tried mounting on carbon tape, and/or } carbon paint but the isopropanol dissolves the sample. I've tried } varying degrees of the carbon and gold coat. I've tried silver } epoxy, I've even tried, after the coating running a thin line of the } carbon paint up the side of the sample onto the area to be viewed. } Of course with the native silica being so easily degraded the carbon } paint was actually eating away the silica aerogel and breaking the } contact of the existing coating. I've tried running a thin copper } wire or copper tape from the face of the sample to the sample } holder. The problem being there is no good method of adhesion to the } sample. I have simply run out of ideas. Does anyone please have a } soluton? } } Linda } } } } Linda S. McCorkle } Ohio Aerospace Institute } Materials Division, Polymers Branch } NASA John H. Glenn Research Center at Lewis Field } 21000 Brookpark Road } Cleveland, OH 44135 } } Tel.: (216) 433-3689 } Fax: (216) 977-7132 } e-mail: Linda.S.McCorkle-at-grc.nasa.gov
-- 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)
} Is there a way to convert an RGB image to a CMMYK image } without having the colors go "haywire"? I ask on behalf of a colleage } who has a request to submit an image for publication in the CMMYK } format, an image that he made in RGB. If you reply, please be sure to } include his email (cardenas-at-bio.umass.edu) because he is not a } subscriber.
I am always surprised when publishers ask for CMYK. Since RGB=} CMYK conversions need CMYK (ink) definitions, how do they expect you to know what the definitions are? Or, they should at least ask you to convert using specific definitions as provided by Photoshop (e.g., Euroscale coated, U.S. sheetfed coated, ...).
As you may now know, ... since RGB=} CMYK involves CRT color definitions (or working color space definitions) going to ink+paper color definitions, there is no way to keep the colors from going "haywire" (i.e., out of gamut).
Sure, I do it all the time for Microscopy Today--thanks to Jerry Sedgewick's Photoshop book!*
Here's what to do: In RGB mode, before conversion, under view, select gamut warning (ctrl-shift-y)**. The out-of-gamut colors in the RGB image will turn gray. Open the 'image/adjust/hue and saturation' box and adjust saturation, color by color (in the drop down box) until the gray overlay disappears, bump up the lightness and then increase saturation a bit until you find the saturation and lightness compromise that doesn't effect contrast too much. Flip in and out of gamut warning to view your progress. You may have to cycle through the color drop down box more than once. Convert to CMYK when finished (Not before!). Try it with copies of images until you become skilled.
Ron
* "Quick Photoshop for Research" Kluwer Academic/Plenum ISBN: 0-306-47375-5. 2002.
** Gamut refers to the range of colors visible in any color system. There are more hues and saturations possible in RGB vs. CMYK. Colors that you see in RGB that you can't see in CMYK are "out of gamut" with respect to CMYK.
-----Original Message----- } From: Tobias Baskin [mailto:Baskin-at-bio.umass.edu] Sent: Monday, June 30, 2003 4:59 PM To: Microscopy-at-sparc5.microscopy.com Cc: cardenas-at-bio.umass.edu
Group, Is there a way to convert an RGB image to a CMMYK image without having the colors go "haywire"? I ask on behalf of a colleage who has a request to submit an image for publication in the CMMYK format, an image that he made in RGB. If you reply, please be sure to include his email (cardenas-at-bio.umass.edu) because he is not a subscriber.
there are many different formats that are being used by different programs. TIFF (or TIF, Tagged Image File Format) is one of the most flexible and seems to be a de-facto standard now. It is now in revision 6, I believe. The format allows not only for data, but also for meta-data, such as magnification, etc., to be stored. It also allows for multiple images in a file, compression, and other goodies. There is a set of "public" tags, which include magnification, and each user can define "private" tags, which are simply ignored by other programs. There are also some drawbacks. For example the tag "magnification". If you put in a large number there, some programs (Word used to do it), would interpret this as information about how big you wanted the image displayed in a document. If the tag was, let's say "1000", Word would interpret this as "the image is 1000 times as large as the original", and try to print it at the original size, resulting in an image a few pixels across. Not good. There are other image formats, some of them require royalty payments (GIF), other store only image data (BMP, JPG). As I mentioned above, TIF seems to be the de-facto standard.
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: jo verbeeck [mailto:joverbee-at-ruca.ua.ac.be] Sent: Tuesday, July 01, 2003 1:16 AM To: MSA listserver
Hi,
I was wondering wether there is a non-commercial file format for storing microscopy images with additional data (like magnification etc) attached to it in a standardized way. There is an MSA EELS format which stores EELS spectra, but I never came across an image format which has the same goal. I`ve read some comments on TIFF images on this list (which seem a good way to go to me). Is anybody working on this, or does it exist already?
Jo -- Dr. J. Verbeeck Electron Microscopy for Materials Research University of Antwerp, Belgium
No relevant experience still .... Perhaps you could get access to a sputter coating unit for a metal which gives a finer coat eg chromium. Some companies have some sort of osmium "coating" device.
Dave
On Mon, 30 Jun 2003 18:24:59 -0500 Becky Holdford {r-holdford-at-ti.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 } -----------------------------------------------------------------------. } } } Linda: you didn't say what kV accelerating voltage you were using. You might try } varying the accelerating voltage to see if you can find a voltage that results in a } non-charging (or very minimally-charging) sample. You might try anywhere from 0.5kV } to 2 kV. Another alternative is to find a colleague with a variable-pressure SEM with } a secondary electron detector that functions in the VP mode (as opposed to the BSD). } } "Linda S. McCorkle" wrote: } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } I am trying to look at a native silica aerogel in a Hitachi S4700 FE-SEM. I } } want to look at a fractured specimen in a 100 to 200 nanometer range. The } } problem seems to be optimal sample preparation. } } I can view the sample with a gold or gold/palladium sputter coat but in } } order to view at the magnification I'm looking for the coating is } } substantial enough that it fills in some of the pores or open pathways and } } totally coats over the primary silica particles. } } I have a comparison of gold sputtered onto a glass slide with a Balzer } } Coater, 40 ma and 1.5 inches to the top of the jar at 10, 20 and 30 } } seconds. The ten second sputter coating viewed by SEM at x250k and 200nm } } somewhat mimics photographically the channels and porosity of the native } } silica. At the higher sputter times of 20 and 30 seconds the pores and } } channels begin to close or fill in. } } I have also tried a carbon deposition coating. It's lighter in density not } } filling in pores and channels and not obliterating primary particles but } } doesn't seem to be conductive enough to dissipate charging in a manner that } } lends itself to anything but low magnification photos. I've tried mounting } } on carbon tape, and/or carbon paint but the isopropanol dissolves the } } sample. I've tried varying degrees of the carbon and gold coat. I've tried } } silver epoxy, I've even tried, after the coating running a thin line of the } } carbon paint up the side of the sample onto the area to be viewed. Of } } course with the native silica being so easily degraded the carbon paint was } } actually eating away the silica aerogel and breaking the contact of the } } existing coating. I've tried running a thin copper wire or copper tape from } } the face of the sample to the sample holder. The problem being there is no } } good method of adhesion to the sample. I have simply run out of ideas. Does } } anyone please have a soluton? } } } } Linda } } } } } } Linda S. McCorkle } } Ohio Aerospace Institute } } Materials Division, Polymers Branch } } NASA John H. Glenn Research Center at Lewis Field } } 21000 Brookpark Road } } Cleveland, OH 44135 } } } } Tel.: (216) 433-3689 } } Fax: (216) 977-7132 } } e-mail: Linda.S.McCorkle-at-grc.nasa.gov } } -- } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ } Becky Holdford (r-holdford-at-ti.com) } 972-995-2360 } 972-648-8743 (pager) } SC Packaging FA Development } Texas Instruments, Inc. } Dallas, TX } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ } } } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
One of the other responders to your post mentioned the technique of moving around on the specimen to minimize localized charging on the specimen surface. This technique takes a little practice, but is the best way to manage a specimen like Silica Aerogel.
At Lawrence Berkeley Laboratory, we sometimes wrapped the specimen in aluminum foil, with a small hole approximately 2~3mm in diameter through which we would view the specimen. This worked well for Auger analysis too.
Hope this helps, Ken Gaugler
Linda S. McCorkle wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } I am trying to look at a native silica aerogel in a Hitachi S4700 } FE-SEM. I want to look at a fractured specimen in a 100 to 200 nanometer } range. The problem seems to be optimal sample preparation. } I can view the sample with a gold or gold/palladium sputter coat but } in order to view at the magnification I'm looking for the coating is } substantial enough that it fills in some of the pores or open pathways } and totally coats over the primary silica particles. } I have a comparison of gold sputtered onto a glass slide with a Balzer } Coater, 40 ma and 1.5 inches to the top of the jar at 10, 20 and 30 } seconds. The ten second sputter coating viewed by SEM at x250k and 200nm } somewhat mimics photographically the channels and porosity of the native } silica. At the higher sputter times of 20 and 30 seconds the pores and } channels begin to close or fill in. } I have also tried a carbon deposition coating. It's lighter in } density not filling in pores and channels and not obliterating primary } particles but doesn't seem to be conductive enough to dissipate charging } in a manner that lends itself to anything but low magnification photos. } I've tried mounting on carbon tape, and/or carbon paint but the } isopropanol dissolves the sample. I've tried varying degrees of the } carbon and gold coat. I've tried silver epoxy, I've even tried, after } the coating running a thin line of the carbon paint up the side of the } sample onto the area to be viewed. Of course with the native silica } being so easily degraded the carbon paint was actually eating away the } silica aerogel and breaking the contact of the existing coating. I've } tried running a thin copper wire or copper tape from the face of the } sample to the sample holder. The problem being there is no good method } of adhesion to the sample. I have simply run out of ideas. Does anyone } please have a soluton? } } Linda } } } } Linda S. McCorkle } Ohio Aerospace Institute } Materials Division, Polymers Branch } NASA John H. Glenn Research Center at Lewis Field } 21000 Brookpark Road } Cleveland, OH 44135 } } Tel.: (216) 433-3689 } Fax: (216) 977-7132 } e-mail: Linda.S.McCorkle-at-grc.nasa.gov } }
A while ago I posted a message on this issue (pasted below). Here's a brief of what I could get to the moment.
At least TIFF and JPG formats have more or less standardized tagged fields to store metadata (= data on the image itself). The appropriate place to store microscope and image setting data seems to be the EXIF fields. (See Digital Still Camera Image File Format Standard. Version 2.1. JEIDA-49-1998. http://www.kodak.com/global/plugins/acrobat/en/service/digCam/exifStandard.pdf. A more recent draft in www.dpnet.com.cn/download/software/exif22.pdf.)
Several programs let you view and/or access the data stored on the EXIF fields, e.g., Photostudio (freeware), IMatch (shareware, $50). Using Photostudio, I can see that our SEM FEI XL stores all the microscope settings and image data in one EXIF field (#8778).
Digital cameras use the more or less standardized EXIF fields for date, resolution, size, etc., while storing other specific data (exposure, flash, diafragm, etc.) in non-standard, proprietary fields. For accessing and managing such data, you need a filter that tells you what to look for in what field. The funny thing is that camera makers tend to keep this information secret (I presume they trade the format specifications with software makers over a commercial agreement). There are several non-official filters that will permit you to read, manage, and edit these camera-specific EXIF fields. I only know those of IMatch (http://www.photools.com/, manual in http://www.photools.net/bin/imatchdoc.zip).
Some microscope makers store all image and microscope settings in only one EXIF field, while leaving the more standardized fields empty. For example, the EXIF field 8778 in our SEM FEI XL looks like this:
Some microscopes produce one text file for every image, with all this data. For example, the Hitachi S4700 FE-SEM produces a file with the same name as the image and .txt extension, looking like this:
In either case, one could easily extract this information with a script in a text editor (i.e., building your own filter), and then load the data in your system of choice.
It seems that other microscope makers went a step beyond, and are way more cryptic. The SEM LEO 1430 VP stores in the EXIF filed 8546 something like this:
In some cases the software that controls the microscope has basic capabilities to build databases with this information. But that machine is typically heavily used by many users on a tight schedule, and it won't help if you collaborate with other researchers using different microscopes.
The microscope image managing programs have specific filters for each microscope type. For instance, the one by Soft Imaging (http://www.soft-imaging.net/) has filters for many microscope makers. If you install the freeware Soft Imaging Viewer, it seems that only the filter for the FEI is activated, the rest are blocked. You can read the metadata from each image, but not really manage it. The full version supposedly has all these filters active and permits data manipulation--but it is expensive.
Reading the EXIF fields or the text files associated with the image, copying to the clipboard, and pasting fragments in a database (or in more meaningful EXIF fields) could work for a few images, but it is not a solution for large batches and for the everyday work.
The question is how we can efficiently manage the data associated with the images, if we don't have the resources for expensive microscope image software.
First of all, knowing the specifications for metadata storage by each microscope is a great start. (For instance, do somebody know how to read the SEM LEO 1430 VP data?)
Ideally, there should be a standard format such that one knows where to look for magnification, KV, etc., in any image coming from a microscope. But if camera makers cannot converge on a standard, I doubt that microscope makers, which seem to be moving in the opposite direction, will converge ever. Perhaps the microscopy community could come to agree on a standard (I think there are such standards in the health sciences) that we all can use to properly document, transmit, and manage our data--it's a lot of work though.
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
} Date: Wed, 19 Feb 2003 14:17:12 -0300 } Hi all, } } I wonder if there are standard (or usual) ways for storing setting data } from electron microscopes (magnification, working distance, acceleration } V, etc.) into the image file itself, such that they can be automatically } imported to a database. Some other devices (like digital cameras) } automatically use the IPTC or EXIF fields for this. } } Any general idea about how preserve and manage these data together with } the images will be very welcome. } } Martín J. Ramírez
} Hi, } } I was wondering wether there is a non-commercial file format for storing } microscopy images with additional data (like magnification etc) attached } to it in a standardized way. } There is an MSA EELS format which stores EELS spectra, but I never came } across an image format which has the same goal. I`ve read some comments } on TIFF images on this list (which seem a good way to go to me). Is } anybody working on this, or does it exist already? } } Jo } -- } Dr. J. Verbeeck } Electron Microscopy for Materials Research } University of Antwerp, Belgium
I have a researcher who dissected out a beetle stomach, digested much of the tissue away with KOH and wants to examine the contents to identify the pollen it was feeding on. There was still a sheath of tissue around it so we ran it through alcohol, placed it and a drop of alcohol between 2 slides, and froze it in liquid nitrogen. When I snapped it apart it just fractured in cross section rather than the longitudinal plane I had hoped. I then stuck a piece of tape over and ripped it off. Basically we ended up just destroying everything, but one of the pieces looked like it might have been some kind of pollen grain at one time or other.
Has anyone done this before? The entire stomach is about 4-500um long and under the light microscope we can see several grains that look like they could be pollen. Any suggestions on how to get them out and captured for SEM examination?? I only have a couple more so experimentation is out and I am running out of time.
Thanks
Scott Whittaker Laboratories of Analytical Biology Smithsonian Institution National Museum of Natural History PO Box 37012 MRC104 Washington DC 20013-7012 202-357-1651
When creating images in Photoshop in RGB space that will be converted to CMYK later, there is a Preview in CMYK mode that should be turned on. The reason is that CMYK is for four color separation ink printing and cannot reproduce many of the more saturated colors in particular, especially the greens, of RGB.
What I do is to keep the image RGB so that info isn't lost, go into the CMYK preview mode and readjust the image to something suitable but admittedly less vivid, and then convert to CMYK.
} Group, } Is there a way to convert an RGB image to a CMMYK image without } having the colors go "haywire"? I ask on behalf of a colleage who has a } request to submit an image for publication in the CMMYK format, an image } that he made in RGB. If you reply, please be sure to include his email } (cardenas-at-bio.umass.edu) because he is not a subscriber. } } Thanks, } Tobias
____________________________________________________________________________ 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/
Is there anyone out there in ListserverLand that has found a way to bring TIF images into Visual Basic 6.0 and work with them? Since I'm doing this for myself, I don't want to buy a commercial package because they pretty expensive. If you have an example of programming code, it would be greatly appreciated.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472
We also have an Oxford EDS, but we have a Hitachi S-2460N rather than a Zeiss.
We have made numerous holders over the years for our scope to accommodate various sample forms. It is certainly nice to have a top referencing holder. It would be one less thing to be concerned about. However, we normally cannot count on all of our samples being at the same working distance even then. We still have to do a little focusing as we move from sample to sample. But when we do the focusing, we try to do most, if not all, of it by using the Z control. We initially set the focal length of the beam at that distance best for x-ray analysis, and then bring the samples to that focal plane.
Therefore, a top referencing holder would eliminate gross adjustments in Z moving from sample to sample, but it is more of a convenience than a necessity. I would probably give up the idea of the top reference and just have the shop machine a holder for a standard and a common thickness of slide plus sample and expect to make fine adjustments in Z as I move between them.
Warren
At 10:33 AM 6/30/2003 -0500, you wrote:
} Dear Listserv members: } } I am looking for help in finding and/or designing a petrographic slide } holder for our SEM. We have a Zeiss DSM960A with a 4-axis mechanical } stage (100x100mm travel). We currently have a 'homemade' holder with room } for one slide (26x46mm) OR one standard block (25mm diam.). I would like } a top-referencing holder for use with our Oxford EDS system. } } I want to come up with something similar JEOL's multi-specimen holder } (model KR-6M150-58) and have considered buying this and modifying it for } our machine. Our main problem in our own designs is the machining } involved (we can do it in-house) and specifically, how do we top-reference } a slide; springs or clamps of some sort or maybe a hinged top plate or...?? } } How have other users tackled this problem when a specific holder is not } produced by your instrument manufacturer? I did not see any messages } relating to this in the archives. Any information or advice would be } tremendously appreciated. } } Thanks for your time. } } Jeff Thole } } Jeff Thole - Geology Laboratory Supervisor and Instructor } Geology Department, Macalester College } 1600 Grand Avenue, St. Paul, MN 55105 } (ph. 651-696-6426, fax. x6122, email thole-at-macalester.edu) } Web: http://www.macalester.edu/geology } ------------------------------------------
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
An excellent and free library for use in Visual Basic is available through the XnView imaging package by Pierre Gougelet. Some of the features include import of ~360 graphic file formats, LUT transforms, filters and a range of tools. It is available at http://perso.wanadoo.fr/pierre.g/xnview/engfl.html
Cheers, Paul Baggethun Alcoa Technical Center
-----Original Message----- } From: Walck, Scott D. [mailto:walck-at-ppg.com] Sent: Tuesday, July 01, 2003 5:12 PM To: Microscopy (E-mail)
Is there anyone out there in ListserverLand that has found a way to bring TIF images into Visual Basic 6.0 and work with them? Since I'm doing this for myself, I don't want to buy a commercial package because they pretty expensive. If you have an example of programming code, it would be greatly appreciated.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472
I believe some of the later printers try to "compensate" for the flat screen and other monitor effects. I know this is a feature of my HP7550 and it got a good review from the latest issue of popular photography magazine. Now, however, I can't say I've done it. But the same article did describe the conversion to CMYK from GB as a necessity for every photographer for the obvious reasons. Photoshop was also discussed, but it didn't allude to any real problems with the conversion. Perhaps one of the digital photography magazines can lend further insight. I just can't imagine there isn't a solution to such a widespread application, just gotta find the people who know the tricks...maybe.
Regards, Ed ----- Original Message ----- } From: "David Elliott Ph.D." {David.Elliott-at-yale.edu} To: {cardenas-at-bio.umass.edu} ; "Microscopy ListServer" {Microscopy-at-sparc5.microscopy.com} Sent: Tuesday, July 01, 2003 4:26 AM
When in doubt, read the destructions! The following are extracts from Adobe Photoshop 5's online help ========================== Reproducing Colour Accurately. *see also: About calibration About ICC profiles Choosing a color management module About the RGB, Grayscale, and CMYK Setup dialog boxes Calibrating your monitor Entering RGB setup information Entering CMYK setup information Using ICC profiles to define the CMYK color space Using the Built-in option to define the CMYK color space Adjusting separation options Printing a color proof Calibrating the screen image to the proof Converting to CMYK Managing ICC profiles in files Converting the color space of open images =========================== Using ICC Profiles to define the CMYK color space.
The CMYK Setup dialog box lets you specify the CMYK color space based on the ICC profile of the printer you select. The CMM then maps the colors in the image to the profiled printer's color gamut, or range of printable colors.You can choose the method (called rendering intent) that is used to translate the colors to the printed gamut.
To use ICC printer profiles to define the CMYK color space:
1 Choose File } Color Settings } CMYK Setup. 2 For CMYK Model, select ICC. 3 Select Preview to display a preview of your changes. A flashing bar under the option indicates a preview is being created. 4 For Profile, choose the printer profile you want to use. If the printer you use is not listed in the Profile menu, contact your printer manufacturer for the appropriate printer profile or create one using third-party printer profiling software. 5 For Engine, choose the CMM you want to use. Built-in refers to Photoshop's built-in CMM.
Note: This option is not the same as choosing Built-in for CMYK Model.
6 For Intent, choose one of the following:
. Perceptual (Images) to maintain the relative color values among the original pixels as they are mapped to the printer gamut. This method preserves the relationship between colors, although the color values themselves may change. . Saturation (Graphics) to maintain the relative saturation values of the original pixels. Out-of-gamut colors are converted to colors that have the same saturation but fall just inside the gamut. . Relative Colorimetric to leave colors that fall inside the gamut unchanged. This method usually converts out-of-gamut colors to colors that have the same lightness but fall just inside the gamut.
. Absolute Colorimetric to disable white point matching when converting colors. This option is not generally recommended.
7 If desired, choose Black Point Compensation to map the darkest neutral color of the source's color space to the darkest neutral color of the destination's color space rather than to black when converting colors. 8 Click OK. ======================== About ICC profiles
One of the methods Photoshop can use to manage color is based on the use of ICC profiles. An ICC profile is a color space description. The ICC profile format was defined by the International Color Consortium (ICC) as a cross-application standard. ICC profiles help you reproduce colors accurately across different platforms, devices, and ICC-compliant applications (such as Adobe Illustrator and Adobe PageMaker®). Adobe Photoshop uses a Color Management Module (CMM) to interpret the ICC profiles that describe the RGB and CMYK color spaces you are using in your system. You can select from existing ICC profiles or create your own. These profiles can then become part of your image files. The CMM interprets the ICC profiles to automatically manage color issues among different color models as well as color issues between your monitor, other monitors, and the final print image. Although you do not have to use ICC profiles, it can greatly simplify managing color.
Important: To ensure that color management works correctly on your system, change the color management settings every time you change printing devices. ========================= hth Chris
Dr. Chris Jeffree Inveresk Cottage 26, Carberry Road Inveresk Musselburgh Midlothian EH21 8PR Tel: +44 131 665 6062 FAX +44 131 653 6248 Mobile 07710 585 401
I had just received some information on 3-CCD cameras when I saw your message (as below). DuncanTech lists some cameras at http://www.duncantech.com/area_scan_cameras.htm
Apparently DuncanTech have been acquired by Redlake ( http://www.redlake.com ) and there are 3-CCD models listed under their website. Not sure of the C-mount though.
=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-= Dr Thor Bostrom Acting Director, Analytical EM Facility; and School of Physical and Chemical Sciences, Queensland University of Technology (QUT) GPO Box 2434, Brisbane, QLD 4001, Australia Ph: +61 7 3864-2351 FAX: +61 7 3864-5100 http://www.sci.qut.edu.au/aemf/ CRICOS No. 00213J =-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=-=
I am looking for information about which database to use to organize my pictures (from the microscope, SEM, EM, confocal, etc.). What database do you use? Why?
:-) Torsten
Torsten Fregin
Universität Hamburg - Zoologisches Institut Abt. Neurophysiologie AG Wiese - Raum 413 Martin-Luther-King-Platz 3 20146 Hamburg, Germany Telefon *49-(0)40-42838-3931 Fax *49-(0)40-42838-3937 eMail Torsten.Fregin-at-zoologie.uni-hamburg.de Torsten-at-Fregin.de
} ... TIF, Tagged Image File Format) is one of the most flexible and } seems to be a de-facto standard now. ... } There are also some drawbacks. For example the tag "magnification". } If you put in a large number there, some programs (Word used to do } it), would interpret this as information about how big you wanted } the image displayed in a document. If the tag was, let's say } "1000", Word would interpret this as "the image is 1000 times as } large as the original", and try to print it at the original size, } resulting in an image a few pixels across. Not good.
This is an interesting point. My own opinion would ask if "magnification" being anything meaningful in the first place. Of course it is in the context of the images original acquisition, but many acquistion softwares, while writing magnification to the file, pay absolutely no attention to "print size" (or print resolution). That is, "mag" and "size" should be interdependent, and there is no provision maintaining the relationship. My first experience with SEM software would write the magnification to the TIF file, but pay absolutely no attention to the size neccessarily being 4 by 5. Therefore the mag was useless, unless it was subsequently always printed correctly.
This would be an interesting plugin for Photoshop ... that is, would recognize the "mag tag", and understand (and maintain) the relationship with print size. It would also be interesting if anyone was aware of current software for maintaining this relationship ... I am not aware of any.
cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com (in progress)
} At least TIFF and JPG formats have more or less standardized } tagged fields to store metadata (= data on the image itself). } The appropriate place to store microscope and image setting } data seems to be the EXIF fields. } (See Digital Still Camera Image File Format Standard. } Version 2.1. JEIDA-49-1998. } {http://www.kodak.com/global/plugins/acrobat/en/service/digCam/exifStandard. pdf} } A more recent draft in www.dpnet.com.cn/download/software/exif22.pdf.) } } [...] } } the EXIF field 8778 in our SEM FEI XL looks like this: } } [DatabarData] } flAccV = 20000.000 } flSpot = 6.000 } [...]
} the Hitachi S4700 FE-SEM produces a file with the same } name as the image and .txt extension, looking like this: } } [SemImageFile] } InstructName=S-4700 } FileName=xxxx 371bALSleft.tif } SampleName= } DataNumber= } } [...] } } In either case, one could easily extract this information } with a script in a text editor (i.e., building your own filter), } and then load the data in your system of choice.
Thanx for this info ... I'll at least know where to explore for more info. I did notice references to "micron bar", but knowing what it meant wasn't obvious. And, I did see references to "magnification", but never did see any instance which would have implied a "print size"(?)
cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com (in progress)
I am into my 6th box of 250 sheets of the new formulation. Up until this 6th box, everything was fine. We use nitrogen burst (2 sec every 10 sec) and Kodak HRP developer. The 6th box, however, is a nightmare. Streaking, uneveness and even darker than the other 5.
Has anybody experienced this variation among the boxes? I can't find any lot number on the boxes to see if there might be variation among the lots.
Grrrr!
Paula :-P
Paula Sicurello George Washington Univ. Medical Center Electron Microscope Lab Washington, DC 20037 202-994-2930 phone 202-994-2518 fax
I would just like to say "Thank you" to everyone that responded to my inquiry on native silica. My biggest problem is the charging issue. When I do get a good image of it at required magnifications, there is too much drift to photograph it. I have received a lot of responses and some offers to get in touch with individuals over the phone. I'm sorting through the information and intend to try some of the suggestions and get in touch with some of the individuals. Once again Thank you Linda
Linda S. McCorkle Ohio Aerospace Institute Materials Division, Polymers Branch NASA John H. Glenn Research Center at Lewis Field 21000 Brookpark Road Cleveland, OH 44135
I am looking for input from all of you experts in quantitative SEM/EDS analyses in trying to optimize conditions for EDS analyses of soda-lime type glass. I am currently trying to estimate the compositions in high alumina (Al2O3 ~ 20 wt%) soda lime glass with B2O3 levels anywhere from 0-15wt%. I am trying to account for some recent results which have been less than satisfactory. I am open to suggestions, indications of pitfalls, and misconceptions involving reliable quantitative analyses of glass by EDS. My objective is to get an estimate of the B2O3 levels with the balance from 100% in non-normalized results. Am I expecting too much from EDS methods by trying to treat the balance of totals below 100% as an indication of levels of light elements not detectable by EDS methods? The instrumentation conditions and procedures I have followed are listed below (in no particular order).
-Utilizing EDAX DX-4 (ver 2.3) system with AmRay 1830 SEM and tungsten filament. -Samples and standards mounted in Struers epoxy/resin mount. -Samples and standards cleaned with 1-micron diamond (to remove previous carbon coat) and methanol. I stopped using acetone, as I believe it leaves a film. -Glass sample and glass standard mounts both coated with carbon at the same time to hopefully insure identical carbon coat thickness. -Beam current closely monitored. Currently 1 nanoamp at 20kv operating voltage. I know lower operating voltage is desirable, but I need to check for titanium and iron. -Unknown glass is referenced to the NBS 620 and NBS 621 glass standards as well as a high alumina, zirconia-bearing in-house standard. -Working distance is closely monitored (frequent degaussing of lenses) to insure it is the same between the sample and standards. -Use box raster scan at ~ 800x for standard and ~1500-2500x in unknown (glass pools at grain boundaries). Should these be the same? -Zero degree tilt (checked with small level) and 24 degree TO angle.
Previously my data gave totals close to 100% in non- borosilicate glass. In borosilicate glasses, the amount less than 100% was consistent with the expected B2O3 levels. Now the results are inconsistent.
Thank you in advance for your anticipated input.
Kevin Selkregg Microanalytical Vesuvius Monofrax Inc
Yes, our JEOL 5600 has a similar txt file that comes along with all the images. I wrote a little program to extract this info in batch mode for import into a spreadsheet, database, or whatever. It's posted at:
http://www.mta.ca/~jehrman/software.htm
Simple to do. I have the (Turbo Pascal, I think) source code if anybody is interested.
Jim
--
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Martin Ramirez writes ... } } } At least TIFF and JPG formats have more or less standardized } } tagged fields to store metadata (= data on the image itself). } } The appropriate place to store microscope and image setting } } data seems to be the EXIF fields. } } (See Digital Still Camera Image File Format Standard. } } Version 2.1. JEIDA-49-1998. } } } {http://www.kodak.com/global/plugins/acrobat/en/service/digCam/exifStandard. } pdf} } } A more recent draft in www.dpnet.com.cn/download/software/exif22.pdf.) } } } } [...] } } } } the EXIF field 8778 in our SEM FEI XL looks like this: } } } } [DatabarData] } } flAccV = 20000.000 } } flSpot = 6.000 } } [...] } } } the Hitachi S4700 FE-SEM produces a file with the same } } name as the image and .txt extension, looking like this: } } } } [SemImageFile] } } InstructName=S-4700 } } FileName=xxxx 371bALSleft.tif } } SampleName= } } DataNumber= } } } } [...] } } } } In either case, one could easily extract this information } } with a script in a text editor (i.e., building your own filter), } } and then load the data in your system of choice. } } Thanx for this info ... I'll at least know where to explore for more info. } I did notice references to "micron bar", but knowing what it meant wasn't } obvious. And, I did see references to "magnification", but never did see } any instance which would have implied a "print size"(?) } } cheerios ... shAf :o) } Avalon Peninsula, Newfoundland } www.micro-investigations.com (in progress)
without trying to sound too commercial, but you asked specifically this question: You may want to take a look at our analySIS software.
As you mentioned (rightly so!), magnification is really a misleading quality, as it always involves the print size. Our approach is to keep track of the calibration (i.e., nm/pixel or similar). This is independent of the display or print size. Of course, if you do display the image, this can then be converted back into a magnification. For example, when you print a report in our software, you can tell it to print the magnification. This will then be the true magnification of that print.
Of course, if you put it on a copier and blow it up 2x, the magnification printed is wrong again.
A scale bar is a much better approach, which we normally use.
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 shaffer [mailto:michael-at-shaffer.net] Sent: Wednesday, July 02, 2003 6:02 AM To: MSA listserver
Mike Bode writes ...
} ... TIF, Tagged Image File Format) is one of the most flexible and } seems to be a de-facto standard now. ... } There are also some drawbacks. For example the tag "magnification". } If you put in a large number there, some programs (Word used to do } it), would interpret this as information about how big you wanted } the image displayed in a document. If the tag was, let's say } "1000", Word would interpret this as "the image is 1000 times as } large as the original", and try to print it at the original size, } resulting in an image a few pixels across. Not good.
This is an interesting point. My own opinion would ask if "magnification" being anything meaningful in the first place. Of course it is in the context of the images original acquisition, but many acquistion softwares, while writing magnification to the file, pay absolutely no attention to "print size" (or print resolution). That is, "mag" and "size" should be interdependent, and there is no provision maintaining the relationship. My first experience with SEM software would write the magnification to the TIF file, but pay absolutely no attention to the size neccessarily being 4 by 5. Therefore the mag was useless, unless it was subsequently always printed correctly.
This would be an interesting plugin for Photoshop ... that is, would recognize the "mag tag", and understand (and maintain) the relationship with print size. It would also be interesting if anyone was aware of current software for maintaining this relationship ... I am not aware of any.
cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com (in progress)
Suggest that you contact Dr. Ken Piel in our office. He is a world class expert on Pollen. E: kenpiel-at-mme1.com P: 413-746-6931
Hope this is helpful.
Barbara Foster Microscopy/Microscopy Education 125 Paridon Street, Suite 102 Springfield, MA 01118 PH: 413-746-6931 FX: 413-746-9311 Web: www.MicroscopyEducation.com
Will you be at M&M in San Antonio? If so, don't forget the Tuesday night seminar on Fluorescence Calibration. Also, join the tradition of over 10,000 microscopists: participate in our survey at any time during the meeting and receive a "sweet thank you". Booth #218 -at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at-
At 01:41 PM 7/1/03 -0400, Scott Whittaker wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Foregut, midgut, or hindgut? The exact answer will depend a little on this, but arthropod gut contents are really pretty simple. Your questions reads like the midgut was removed. Fix, dehydrate, and critical point dry the intact gut + contents. HMDS could work here, especially for pollen. Mind, I'd suggest avoiding KOH digestion anyway. Dissect and fix, or, if the critter is too small, fix and dissect. Put double sticky tape on a SEM stub (I would've said double-sticky carbon tab, but given the ones sold now aren't very sticky anymore I won't). Or else use Al foil tape -- glue the Al side down with Ag paint. Keep the paint handy. Stick the dried gut on the tape glue ("sticky tabs" or sticky "paint" aren't sticky enough). Cut (tear) a line along the gut to open it. Use a #0 insect pin for this, smaller or larger depending on the beetle. Roll the gut along the tape. The contents will be exposed as the gut is stuck to the tape. Carefully use a fine insect pin to spread the gut contents out on the tape. Not a brush or anything similar, or the sample will be lost in the brush. Take the Ag paint and using a sharpened applicator stick (or the like) paint a ring close around the gut contents, then a line from this ring to the stub. This will leave less insulting surface (tape glue) between the sample and the conductive path to ground. Gold coat. Note that the foregut is chitinous, part of the exoskeleton. So it's more robust and easier to handle than contents from the midgut or hindgut, which are often in a fine, membranous structure. I think this is what you describe, but I'm not entirely certain you mean this "bag", and not the gut itself. The method works regardless of which you have, you just have to be more careful the farther back you go in the beetle, or if you have the "bag" and not the gut itself. You might want to get a pair of flexible, "fine" point dried insect handling forceps for handling the guts. Fine Science Tools carries these, but I'm sure other companies do also (especially entomology supply houses). I used this method a fair amount to look at amphipod guts, which are smaller than the beetle stomachs you describe. Just cut back on the coffee, or have a beer for lunch, before starting.
Phil
} I have a researcher who dissected out a beetle stomach, digested much of the } tissue away with KOH and wants to examine the contents to identify the } pollen it was feeding on. There was still a sheath of tissue around it so we } ran it through alcohol, placed it and a drop of alcohol between 2 slides, } and froze it in liquid nitrogen. When I snapped it apart it just fractured } in cross section rather than the longitudinal plane I had hoped. I then } stuck a piece of tape over and ripped it off. Basically we ended up just } destroying everything, but one of the pieces looked like it might have been } some kind of pollen grain at one time or other. } } Has anyone done this before? The entire stomach is about 4-500um long and } under the light microscope we can see several grains that look like they } could be pollen. Any suggestions on how to get them out and captured for SEM } examination?? I only have a couple more so experimentation is out and I am } running out of time. } } } Thanks } } Scott Whittaker } Laboratories of Analytical Biology } Smithsonian Institution } National Museum of Natural History } PO Box 37012 MRC104 } Washington DC 20013-7012 } 202-357-1651
-- 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)
I'm still having problems with uneven negatives. It really varies a lot. There are days that the problem is not as obvious and at times really bad negatives.
I wonder if there is an alternative brand? Does Fuji or Ilford make any EM film? If so where can one purchase them?
Rajesh Patel Robert Wood Johnson Medical School Department of Pathology 675 Hoes Lane Piscataway, NJ 08854
} } Fellow microscopists. } } } } Due to the imminent departure of my colleague Corinna Wauchope for the } } Land Down Under, I am in need a a person to help me run the North } } Campus EMAL at the University of Michigan } } (http://emalwww.engin.umich.edu/emal/fset.html). QUALIFIED } } individuals } } should respond to the job posting which is at: } } http://websvcs.itd.umich.edu/jobnet/ } } job_posting.php?postingnumber=3D032406&searchwhat=3Dcurrent } } DO NOT send resumes or applications to me personally they will NOT get } } processed. Please follow the instructions on the Job Posting Web } } Site. } } } } (http://www.umich.edu/~jobs/). } } } } } } } } John Mansfield PhD MInstP } } North Campus Electron Microbeam Analysis Laboratory } } 417 SRB, University of Michigan } } 2455 Hayward, Ann Arbor MI 48109-2143 USA } } Phone: (734) 936-3352 } } FAX (734) 763-2282 } } Cell. Phone: (734) 834-3913 } } (Leaving a phone message at 936-3352 is preferable to 834-3913) } } Email: jfmjfm-at-engin.umich.edu } } URL: http://emalwww.engin.umich.edu/people/jfmjfm/jfmjfm.html } } Location: Lat. 42=B0 16' 48" Long. 83=B0 43' 48"
Just a thought - make certain that the surface is really free of 1 micron alumina. At 20kV these particles can be nearly transparent to the beam depending on the surface beneath, especially if you are working in the backscatter mode. Their presence could inflate the apparent response from aluminum, throwing off the calculation by difference.
The boron content (presumably by weight?) sounds extremely high and potentially subject to volatilization. I understand that you are rastering the beam which should reduce that problem. However, you don't indicate what the beam diameter is. You might consider spreading out the beam as an additional means of avoiding volatile losses.
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } I am looking for input from all of you experts in quantitative SEM/EDS analyses in trying to optimize conditions for EDS analyses of soda-lime type glass. I am currently trying to estimate the compositions in high alumina (Al2O3 ~ 20 wt%) soda lime glass with B2O3 levels anywhere from 0-15wt%. I am trying to account for some recent results which have been less than satisfactory. I am open to suggestions, indications of pitfalls, and misconceptions involving reliable quantitative analyses of glass by EDS. My objective is to get an estimate of the B2O3 levels with the balance from 100% in non-normalized results. Am I expecting too much from EDS methods by trying to treat the balance of totals below 100% as an indication of levels of light elements not detectable by EDS methods? The instrumentation conditions and procedures I have followed are listed below (in no particular order). } } -Utilizing EDAX DX-4 (ver 2.3) system with AmRay 1830 SEM and tungsten filament. } -Samples and standards mounted in Struers epoxy/resin mount. } -Samples and standards cleaned with 1-micron diamond (to remove previous carbon coat) and methanol. I stopped using acetone, as I believe it leaves a film. } -Glass sample and glass standard mounts both coated with carbon at the same time to hopefully insure identical carbon coat thickness. } -Beam current closely monitored. Currently 1 nanoamp at 20kv operating voltage. I know lower operating voltage is desirable, but I need to check for titanium and iron. } -Unknown glass is referenced to the NBS 620 and NBS 621 glass standards as well as a high alumina, zirconia-bearing in-house standard. } -Working distance is closely monitored (frequent degaussing of lenses) to insure it is the same between the sample and standards. } -Use box raster scan at ~ 800x for standard and ~1500-2500x in unknown (glass pools at grain boundaries). Should these be the same? } -Zero degree tilt (checked with small level) and 24 degree TO angle. } } Previously my data gave totals close to 100% in non- borosilicate glass. In borosilicate glasses, the amount less than 100% was consistent with the expected B2O3 levels. Now the results are inconsistent. } } Thank you in advance for your anticipated input. } } Kevin Selkregg } Microanalytical } Vesuvius Monofrax Inc
Paula: We were among the first ones to have problems with the "new formulation" and as my people used to say it is a "limon" or a lemon in English. This film has never worked properly for us, even as we accommodate our processing to fit its particular needs. We go through 200 to 250 plates in a week to week and a half, so we see the differences in each box and lot. Each box of film is like a box of melted chocolates. You never know the mess you will get. We use Kodak D19 at 2:1 water/developer ratio, for developing and each time I develop, new surprises come to life. I'm trying to contain the comments that this film deserve in reality. The old formulation worked great and it was consistent from box to box. But now this film is just making life very interesting in the dark room. I'll just say, is just a matter of time and patience to develop the "new Formulation".
Omayra Velez EM Specialist Patholy Department Weill Cornell Medical College NY,NY. 10021 Phone: 212-746-6437
--- Paula Sicurello {patpxs-at-gwumc.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 Listers, } } I am into my 6th box of 250 sheets of the new } formulation. Up until this 6th box, everything was } fine. We use nitrogen burst (2 sec every 10 sec) } and Kodak HRP developer. The 6th box, however, is a } nightmare. Streaking, uneveness and even darker } than the other 5. } } Has anybody experienced this variation among the } boxes? I can't find any lot number on the boxes to } see if there might be variation among the lots. } } Grrrr! } } Paula :-P } } Paula Sicurello } George Washington Univ. Medical Center } Electron Microscope Lab } Washington, DC 20037 } 202-994-2930 phone } 202-994-2518 fax } }
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (bop00rav-at-sheffield.ac.uk) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 2, 2003 at 16:59:13 ---------------------------------------------------------------------------
Email: bop00rav-at-sheffield.ac.uk Name: BOB VASEY
Organization: UNIVERSITY OF SHEFFIELD
Education: Graduate College
Location: sheffield, uk
Question: Hello,
My name is Robert Vasey and I am a PhD student at the university of Sheffield, England. I am working on Arabidopsis roots. I am currently trying to make root sections to stain for lignin and cellulose. I have been using LR White resin but it does not allow my stains to penetrate (phloroglucinol for lignin). I have also tried staining with Methylene Blue followed by counterstaining with Fuchsin Red, without success.
To stain for cell wall components, specifically lignin and cellulose, can anyone advise me on:
(a) What is the appropriate resin to use (b) What stains can I use - I would be grateful for recipes and how to apply them. (c) Are there any good counterstaining techniques or metachromatic stains to stain multiple cell components (apart from Toluidine Blue).
I would be grateful for any advice as I am struggling with this.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (gaston.garcia-at-123.cl) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Tuesday, July 1, 2003 at 14:25:02 ---------------------------------------------------------------------------
Email: gaston.garcia-at-123.cl Name: gaston
Organization: leon prado school
Education: Graduate College
Location: santiago, chile
Question: I have a microscope with a reticle eyepiece. I want to know the scale of reticle. I see for an objetive 40X/0.65 and the scale is a 100 parts. Can you help me please?
Dear Sir or Madam: We take the liberty to write to you and will appreciate your attention on this mail. We have a foundry engaging in brake disc casting and have been in this field for years and have gained the fame for our products step by step.
With the CNC lathe from Japan, we are able to keep the run out below 0.04mm and the surface roughness to be 1.6.
Now we export about 50 containers of brake rotor and other casting parts annually to the US market, but our ability is not fully used and we are now seeking for new buyers.
Quotation by drawings or samples will be ok and we will quote in 3days after reception of your inquiry.
You can refer to our website for details: www.szautoparts.com and www.bacoengineering.com
Thanks and best regards
China machinery imp/exp co., ltd Tel: 86-512-65325936 Fax: 86-512-65321605 Korin Tian the sales manager
Dear Sir or Madam: We take the liberty to write to you and will appreciate your attention on this mail. We have a foundry engaging in brake disc casting and have been in this field for years and have gained the fame for our products step by step.
With the CNC lathe from Japan, we are able to keep the run out below 0.04mm and the surface roughness to be 1.6.
Now we export about 50 containers of brake rotor and other casting parts annually to the US market, but our ability is not fully used and we are now seeking for new buyers.
Quotation by drawings or samples will be ok and we will quote in 3days after reception of your inquiry.
You can refer to our website for details: www.szautoparts.com and www.bacoengineering.com
Thanks and best regards
China machinery imp/exp co., ltd Tel: 86-512-65325936 Fax: 86-512-65321605 Korin Tian the sales manager
Dear reader, We used for years in combination with TEM the Reflection Contrast Microscope (RCM)for a overview of the ultrathin sequentional sections (EPON,Spurr, Lowicryl or Cryo) Al you need is a fluorescence microscope with 3 extra parts: A central stop, antiflex objective and a polarization block for more infomation see the website or mail
At the NESM Woods Hole Meeting in May, the Kodak rep suggested adjusting the nitrogen burst to 2sec every 8sec (instead of 10). I have no clue if this will help.
Roger Moretz, Ph.D. Dept of Toxicology BI Pharmaceuticals
-- Where the world is only slightly less weird than it actually is. } ------------------------------------------------------------------------ } 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 Listers, } } I am into my 6th box of 250 sheets of the new formulation. Up until this 6th } box, everything was fine. We use nitrogen burst (2 sec every 10 sec) and Kodak } HRP developer. The 6th box, however, is a nightmare. Streaking, uneveness and } even darker than the other 5. } } Has anybody experienced this variation among the boxes? I can't find any lot } number on the boxes to see if there might be variation among the lots. } } Grrrr! } } Paula :-P } } Paula Sicurello } George Washington Univ. Medical Center } Electron Microscope Lab } Washington, DC 20037 } 202-994-2930 phone } 202-994-2518 fax } }
Greetings, As for part a, while I have no specific experience with histo stains on LR White, I can say that using Butyl-methyl methacrylate produces blocks that section much more easily than does LR White, and certainly stain well with many things as a rule. As for part b, a typical stain for celluose is calcofluor white (sometimes called cellufluor). It is not strictly specific for cellulose as it will stain some other beta glucans, but it is easy and widely used. You can also use polarized light which avoids staining altogether and can be quantitative.
Good luck, Tobias Baskin } } } Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (bop00rav-at-sheffield.ac.uk) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, } July 2, 2003 at 16:59:13 } --------------------------------------------------------------------------- } } Email: bop00rav-at-sheffield.ac.uk } Name: BOB VASEY } } Organization: UNIVERSITY OF SHEFFIELD } } Education: Graduate College } } Location: sheffield, uk } } Question: Hello, } } My name is Robert Vasey and I am a PhD student at the university of } Sheffield, England. I am working on Arabidopsis roots. I am } currently trying to make root sections to stain } for lignin and cellulose. I have been using LR White resin but it } does not allow my stains to penetrate (phloroglucinol for lignin). I } have also tried staining with Methylene Blue followed by } counterstaining with Fuchsin Red, without success. } } To stain for cell wall components, specifically lignin and } cellulose, can anyone advise me on: } } (a) What is the appropriate resin to use } (b) What stains can I use - I would be grateful for recipes and how } to apply them. } (c) Are there any good counterstaining techniques or metachromatic } stains to stain multiple cell components (apart from Toluidine Blue). } } I would be grateful for any advice as I am struggling with this. } } Bob. } } } } ---------------------------------------------------------------------------
The reticle scale can sometimes be arbitrary. Regardless of whether it is arbitrary or not, you need to calibrate the reticle. There is an item available called a "microscope slide", which is a special glass slide that you place on the microscope stage. The slide has measureable divisions of known length appropriate for calibrating high magnifications. The calibration would need to be made for each objective lens on the microscope.
Stu Smalinskas Metallurgist SKF USA Plymouth, Michigan
-----------------------------------
Gaston wrote:
Email: gaston.garcia-at-123.cl Name: gaston
Organization: leon prado school
Education: Graduate College
Location: santiago, chile
Question: I have a microscope with a reticle eyepiece. I want to know the scale of reticle. I see for an objetive 40X/0.65 and the scale is a 100 parts. Can you help me please?
__________________________________ Do you Yahoo!? SBC Yahoo! DSL - Now only $29.95 per month! http://sbc.yahoo.com
Good day to all, I have been a member of MSA for over 5 years now - I first joined in Oklahoma. In Oklahoma, the local society was quite busy and I really enjoyed being a part of a group of people that enjoyed the same things I did! My current problem is that I live in North Carolina where the local chapter seems to have completely died - or at least no one will let me know anything about meetings etc even after repeated request! My question - is there any other state society near NC that would allow me the opportunity to join or does anyone in the national MSA society have suggestions as to how to start a local chapter or revive the one that is suppose to be in NC? Thanks a whole bunch, Connie Cummings
Connie,
The Appalacian Regional Microscopy Society (AREMS) is an active chapter of MSA and serves microscopists in VA, WV, NC, SC, TN, and GA. You can find more about AReMS on-line at www.uvawise.edu/arems/index.html or contact me if you wish to become a member. My numbers are below.
Thank you,
Rich
Richard Fiore NC State University Analytical Instrumentation Facility 2410 Campus Shore Drive 318 EGRC, Campus Box 7531 Raleigh, NC 27695 Tel: 919-515-2348 Fax: 919-515-6965 Email: rafiore-at-ncsu.edu
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I forgot to meantion that the next AReMS meeting is October 2-3, 2003 in Boone, NC at the Broyhill Inn & Conference Center.
Rich
Original Message -------- Subject: MSA help
Good day to all, I have been a member of MSA for over 5 years now - I first joined in Oklahoma. In Oklahoma, the local society was quite busy and I really enjoyed being a part of a group of people that enjoyed the same things I did! My current problem is that I live in North Carolina where the local chapter seems to have completely died - or at least no one will let me know anything about meetings etc even after repeated request! My question - is there any other state society near NC that would allow me the opportunity to join or does anyone in the national MSA society have suggestions as to how to start a local chapter or revive the one that is suppose to be in NC? Thanks a whole bunch, Connie Cummings
Connie,
The Appalacian Regional Microscopy Society (AREMS) is an active chapter of MSA and serves microscopists in VA, WV, NC, SC, TN, and GA. You can find more about AReMS on-line at www.uvawise.edu/arems/index.html or contact me if you wish to become a member. My numbers are below.
Thank you,
Rich
Richard Fiore NC State University Analytical Instrumentation Facility 2410 Campus Shore Drive 318 EGRC, Campus Box 7531 Raleigh, NC 27695 Tel: 919-515-2348 Fax: 919-515-6965 Email: rafiore-at-ncsu.edu
A position is available at Texas Instruments in Dallas TX as a Physical Failure Analyst. Details are provided below. To apply e-mail your resume to faresume-at-list.ti.com with "DMFA" in the subject line of the e-mail.
Texas Instruments Product Engineer SiTD CMOS1
You'll develop the latest technology and will have many learning opportunities in this dynamic engineering position.
As a CMOS1 Product Engineer, you'll be responsible for PFA and EFA of C021 material. Other responsibilities include: SRAM cell debug CO21 F/A of low K dielectrics Metal gate failure analysis Leading edge product failure analysis on wireless, DSP, and microprocessors Pareto creation Baseline F/A Scrap assignment
The ideal candidate will have 5-10 years of related experience. Process knowledge is a must. PFA candidates with Cu experience on 130nm or 90nm nodes are preferred. Experience with reliability fails and constructional analysis is also preferred.
Additional information follows Primary Responsibilities: PFA and EFA of C021 Products and understanding of process limitations through physical and electrical characterization Supporting & Secondary Responsibilities: Baseline pareto creation and development. Interaction with Process Integration engineers, Process Engineers, and other Product Engineers in order to ensure that the correct solutions are implemented for yield and reliability learning. Management /Organization Skills: Detail individual with a desire to derive statistically significant backup for the improvement of our baseline process, and the evaluation of new processes. Opportunities for Improvements: Electrical Failure Analysis, device physics understanding, process knowledge, micromanipulator tool knowledge, automated FIB, TEM, SEM, statistical analysis, copper interconnect, low-K dielectrics, metal gates, etc. Complex Tasks: Failure analysis of processes that have never been F/A before. Fail mechanisms that do not have any history behind them. Fail mechanisms that will push past the common failure analysis techniques. Technical Abilities: 130nm/90nm process, deprocessing techniques, electrical isolation techniques, SEM use, TEM use, FIB use Team & People Skills: Work with a team of a handful of failure analysis engineers as part of a group working on the qualification of TI's 65nm technology. Interaction with business units ranging from the wireless products to microprocessors. Projects & Deliverables: Failure pareto, constructional analysis, process characterization, design rules and yield limiters, scrap assignment, etc. Additional skills and experience (include years of required experience): 5-10 years List unique selling features of this position, team or project: Dynamic team involved on the development of TI's 65nm technology. Opportunity to gain visibility within the development team, TI's manufacturing facilities, and business units. Special Work Environment: F/A lab with chemicals, SEM, FIB, TEM, polish wheels, etc. EFA lab with all the necessary tools for electrical isolation.
E-mail your resume to faresume-at-list.ti.com with "DMFA" in the subject line of the e-mail.
Kim Christensen Texas Instruments, Inc 13560 N Central Expressway Dallas, TX 75243 Ph: 972-995-3855 E-mail: kchristen-at-ti.com
What you need to find is something you know the size of that you can look at through the microscope. You can then calibrate the grid you have for each objective size. Red blood cells are of a known size and can give you an approximation of the scale of the grid. What you really need is called a "stage micrometer". This is an accurate scale mounted on a slide which will allow you to accurately calibrate your grid. If you can, take a look at this web site for more information:
Dr. Allerton: I am the President/CEO of a company in Natick, MA that specializes in Optical coatings. We specialize in antireflection coatings of optics. I am going to have to look a little bit further in to the exact specifications of the AR coating on your microscope objectives but I believe we should be able to strip and recoat the lenses with a durable coating with specifications at least as good as the original.
Bob O'Leary - President, CEO Optical Coating Corporation 01760 6R Mercer Road Natick, MA P: 508.655.1650 F: 508.653.0729 http://www.opticalcc.com BOLeary-at-opticalcc.com
----- Original Message ----- } From: "Ron Allerton" {rsallerton-at-caribsurf.com} To: {Microscopy-at-sparc5.microscopy.com} Cc: {zaluzec-at-sparc5.microscopy.com} Sent: Tuesday, July 01, 2003 3:41 PM
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Hi listers,
Perhaps I didn`t put my question very clearly. I am aware that tiff can hold extra info like mag (and calibration and scale bar etc). But my main question is: is there a STANDARD (convention) way of naming these tags. It seems to me that a lot of tags could be defined so they are useful in TEM, SEM, STM, LM etc etc. Defining a set of rules like this could maybe stop commercial companies of inventing their own file formats (which is a bad idea for scientists in all respects except for the company monopoly)
The good thing about that format (the MSA-tiff?) is that it is still viewable with a normal image viewer but to get more info you need special software. There are good GNU imageviewers available that allow to easily include this new image format and give them the functionality (and better) of other commercial (very expensive) programs.
kind regards,
Jo
On Wed, 2003-07-02 at 19:33, James M. Ehrman 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 } -----------------------------------------------------------------------. } } } Yes, our JEOL 5600 has a similar txt file that comes along with all } the images. I wrote a little program to extract this info in batch mode } for import into a spreadsheet, database, or whatever. It's posted at: } } http://www.mta.ca/~jehrman/software.htm } } Simple to do. I have the (Turbo Pascal, I think) source code if anybody } is interested. } } Jim } } -- } } James M. Ehrman } Digital Microscopy Facility } Mount Allison University } Sackville, NB E4L 1G7 } CANADA } } phone: 506-364-2519 } fax: 506-364-2505 } email: jehrman-at-mta.ca } www: http://www.mta.ca/~jehrman } } } } michael shaffer 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 } } -----------------------------------------------------------------------. } } } } Martin Ramirez writes ... } } } } } At least TIFF and JPG formats have more or less standardized } } } tagged fields to store metadata (= data on the image itself). } } } The appropriate place to store microscope and image setting } } } data seems to be the EXIF fields. } } } (See Digital Still Camera Image File Format Standard. } } } Version 2.1. JEIDA-49-1998. } } } } } {http://www.kodak.com/global/plugins/acrobat/en/service/digCam/exifStandard. } } pdf} } } } A more recent draft in www.dpnet.com.cn/download/software/exif22.pdf.) } } } } } } [...] } } } } } } the EXIF field 8778 in our SEM FEI XL looks like this: } } } } } } [DatabarData] } } } flAccV = 20000.000 } } } flSpot = 6.000 } } } [...] } } } } } the Hitachi S4700 FE-SEM produces a file with the same } } } name as the image and .txt extension, looking like this: } } } } } } [SemImageFile] } } } InstructName=S-4700 } } } FileName=xxxx 371bALSleft.tif } } } SampleName= } } } DataNumber= } } } } } } [...] } } } } } } In either case, one could easily extract this information } } } with a script in a text editor (i.e., building your own filter), } } } and then load the data in your system of choice. } } } } Thanx for this info ... I'll at least know where to explore for more info. } } I did notice references to "micron bar", but knowing what it meant wasn't } } obvious. And, I did see references to "magnification", but never did see } } any instance which would have implied a "print size"(?) } } } } cheerios ... shAf :o) } } Avalon Peninsula, Newfoundland } } www.micro-investigations.com (in progress) } } } -- Dr. J. Verbeeck Electron Microscopy for Materials Research University of Antwerp, Belgium
by sparc5.microscopy.com (8.9.3+Sun/8.9.3) id VAA14149 for dist-Microscopy; Fri, 4 Jul 2003 21:23:40 -0500 (CDT) Received: from njz_spm_filter (sparc5 [206.69.208.10]) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) with SMTP id VAA14146 for "MicroscopyFilteredEmail4-at-msa.microscopy.com"; Fri, 4 Jul 2003 21:23:09 -0500 (CDT) Received: from mta7.pltn13.pbi.net (mta7.pltn13.pbi.net [64.164.98.8]) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) with ESMTP id VAA14139 for {microscopy-at-sparc5.microscopy.com} ; Fri, 4 Jul 2003 21:22:56 -0500 (CDT) Received: from nigelhomepc (adsl-64-175-40-97.dsl.pltn13.pacbell.net [64.175.40.97]) by mta7.pltn13.pbi.net (8.12.9/8.12.3) with SMTP id h652GgeS024709 for {microscopy-at-msa.microscopy.com} ; Fri, 4 Jul 2003 19:16:42 -0700 (PDT) Message-Id: {4.1.20030704190401.00ae4960-at-yellow.ucdavis.edu} X-Sender: nigel-at-yellow.ucdavis.edu X-Mailer: QUALCOMM Windows Eudora Pro Version 4.1
POSTDOCTORAL POSITIONS IN ELECTRON MICROSCOPY
DEPARTMENT OF CHEMICAL ENGINEERING AND MATERIALS SCIENCE UNIVERSITY OF CALIFORNIA-DAVIS
Two postdoctoral positions are available in the Electron Microscopy Group at the University of California-Davis (UCD). Research in the newly formed Electron Microscopy Group focuses on the use of atomic resolution imaging and analytical techniques in electron microscopy, coupled with theoretical simulations, to determine the structure-property relationships at internal interfaces on the fundamental atomic scale. Current research programs involve catalysts, ferroelectrics, high-Tc superconductors and optoelectronic/high-power semiconducting materials and devices. The positions that are currently available are for the study of metal-oxide interactions in heterogeneous catalysts and the vacancy mediated fatigue mechanisms in ferroelectric thin films. The experimental analyses will primarily be performed using the facilities in the National Center for Electron Microscopy at Lawrence Berkeley National Laboratory. In particular, extensive use will be made of the new 200kV monochromated FEI G2 Tecnai that has recently been installed and the Cs-corrected VG HB501 dedicated STEM that will be installed later this year. There will also be the opportunity for both postdoctoral appointees to be involved in the development of in-situ stages for the recently funded 200kV field emission TEM to be installed at UC-Davis next year. Successful candidates will be recent Ph.D. graduates in physics, metallurgy, or materials science with a sound background in the relevant materials issues and an ambition to be part of a developing program pushing at the frontiers of interface science. Please send a resume and publication list to Professor Nigel D. Browning at the address below. Prior experience in STEM or TEM is essential. However, consideration will be based on the candidates overall potential for success in the field and applicants with prior experience in related fields are encouraged to apply. Positions are for one year initially, normally renewed for a second year with possibilities existing for further years. Salary is commensurate with experience.
Nigel D. Browning
Department of Chemical Engineering and Materials Science University of California-Davis One Shields Ave Davis, CA 95616
AND
National Center for Electron Microscopy, MS 72-150 Lawrence Berkeley National Laboratory Berkeley, CA 94720
EPSRC funded Ph.D. Research Project on the Chemistry and Biological Activity of Nanoparticles using Aberration-Corrected Scanning Transmission Electron Microscopy (SuperSTEM).
Institute for Materials Research (IMR) at the University of Leeds and Daresbury Laboratories
Supervisors: Dr Rik Brydson & Dr Andy Brown, Institute for Materials Research, University of Leeds, Leeds LS2 9JT, U.K. Please Contact mtlrmdb-at-leeds.ac.uk or a.p.brown-at-leeds.ac.uk or phone +44 (0)113 343 2348/2369 for further details.
What is SuperSTEM ? The SuperSTEM project involves the development of two aberration-corrected STEM instruments in a five-year, £4.2 million project funded as a result of the NW Science Review. The microscopes have been installed in a specially designed and purpose built facility on the Daresbury Laboratory site in Cheshire, and ultimately they will be available as a national resource. SuperSTEM provides a world-leading atomic imaging and analysis facility capable of chemical spectroscopy from single atoms and atomic columns further details may be found by visiting www.superstem.org.uk. The SuperSTEM project is led by five scientists from four partner universities [Peter Goodhew, Mick Brown, Alan Craven, Rik Brydson and Gordon Tatlock]. The full-time research team comprises a Technical Director [Dr Andrew Bleloch], two research microscopists [Dr Uwe Falke and Dr Meiken Falke], a computer specialist [Will Costello] and a technician [Peter Shiels]. This project is part of a set of six studentships recently funded by the Engineering and Physical Sciences Research Council and as such is fully funded (fees and maintenance) for both UK and EC students. The project will be based at the IMR in Leeds but will involve extensive travel to Daresbury Laboratories.
The studentship will examine a limited range of metal oxide nanoparticles of varying mean particle size and size distribution. Systems of current interest include: (a) bare, carboxylic acid stabilized and charge stabilized Fe3O4 nanoparticles produced by standard colloidal routes at Leeds. (b) g-Al2O3 and -g-Al2O3/ CeO2 catalyst supports produced at Leeds or obtained commercially.
Atomic resolution HAADF and EELS measurements using the SuperSTEM I facility will be undertaken. With an aberration corrected probe size of around 0.14 nm and below it will be possible to resolve the oxygen sub-lattice. A combination of Z contrast imaging and EELS measurements at particle surfaces (both direct and aloof beam measurements to maximize surface excitations) and within the particle core will give structural and spatially resolved chemical/ electronic information. It is intended that results will be compared with theoretical calculations, both ab-initio total energy/ electronic structure calculations as well thermodynamic predictions.
An extension of this project, pertaining directly to the life sciences, would involve an investigation of iron oxide nanoparticles in human liver biopsies in collaboration with Dr Alice Warley and Dr Jonathan Powell at King¹s College London. -- _____________________________ Dr. Rik Brydson, Leeds Electron Microscopy and Spectroscopy (LEMAS) Centre Institute for Materials Research, University of Leeds, Leeds LS2 9JT, U.K.
New MSc in Nanoscale Science and Technology Visit http://www.ee.leeds.ac.uk/nanomsc/ ______________________________
UK SuperSTEM facility - Atomic resolution Analysis http://www.superstem.org.uk
_______________________________
Bedtime reading.......... Series: Microscopy Handbooks - Electron Energy Loss Spectroscopy by Rik Brydson Published by BIOS September 2001 £29.99 ISBN: 1859961347
************************** Join email discussion list for all aspects of Electron Microscopy and Analytical Spectroscopy (LEMAS)
visit website http://www.jiscmail.ac.uk/lists/lemas.html and follow instructions **************************
===================================== EELS and X ray database : http://www.cemes.fr/~eelsdb/ =====================================
I don't think there is or will be ever a standard that can anticipate and provide tags for all possible fields. Let's face it: the imaging field is HUGE, and microscopy is a tiny part of that. You have many fields, where magnification is a meaningless quantity. For example, take all your vacation photos. You can't assign a magnification to them. Then there is the medical field. Do you want to provide a field for each disease of each bone for the X-rays? Dental images, Forensics, Space imaging, Military Imaging, etc., all with their specific requirements. TIF proides SOME standard fields, but they are metadata about the image itself (size, magnification, etc.) If you need to store more information, you need to use an archiving system (here again a pointer to our software, but other software does similar things).
If I remember corectly, the TIF tags are just numbers. Certain numbers are defined in the TIF specs and have specific meaning (for example "Number of pixels in X direction"). Other numbers are not defined and can be used by other software.
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: jo verbeeck [mailto:joverbee-at-ruca.ua.ac.be] Sent: Friday, July 04, 2003 1:49 AM To: MSA listserver
My name is Bob Vasey and I am currently conducting light microscopy on Arabidopsis using LR White resin. I want to stain root sections for cell wall components, specifically lignin and cellulose. I would also like to try some counterstains to stain multiple cell wall components on a single section.
Could anyone help with the following:
(a) A protocol for Methylene Blue/Fuchsin counterstaining that works in LR White (including recipes for stains). When I try, the Fuchsin seems to wash out the Toluidine Blue. (b) Suggest any other polychromatic stains (apart from Toluidine Blue) or counterstains. (c) I have tried phloroglucinol for lignin but it does not seem to be taken up, probably because it is made up in ethanol. Are there any other stains/protocols that work for lignin in LR White. Will phloroglucinol work in epoxy resins, e.g. Spurr’s or Araldite? (d) Am I using the best resin – do other resins allow a greater diversity of stains to be used. Does LR White give the best quality (intensity) of stain – sometimes my sections look a bit pale when Toluidine Blue is used. (e) I have heard a stain called Paragon mentioned – could anyone tell me what it is and how to make and use it? (f) In the literature, some people fix in glutaraldehyde, some in formaldehyde and some in both. Could anyone tell me the difference between them and which is the best to use?
I would be grateful for any advice as I am a PhD student in a department with limited experience of conducting staining of resin sections, so I am having to work it out as I go along!
My email is { HYPERLINK "mailto:bop00rav-at-sheffield.ac.uk" }bop00rav-at-sheffield.ac.uk. The two “00” in the middle are zeros and not capital “o”s!
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (bbaxter-at-westminstercollege.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Sunday, July 6, 2003 at 13:18:04 ---------------------------------------------------------------------------
Email: bbaxter-at-westminstercollege.edu Name: Bonnie K. Baxter
Organization: Westminster College
Education: Undergraduate College
Location: SLC, UT
Question: I am attempting some microscopy of microbes from Great Salt Lake. Our best scope is a Nikon inverted phase contrast. The problem I'm having is doing iol immersion with an inverted scope. ANy pointers for doing this upside down and actually keeping the bugs in focus? thanks!! Bonnie
We have a Denton Vacuum DFE-3 Freeze Etch Apparatus with DRM-1 Resistance Monitor (for precision C and C-Pt film deposition) available to anyone interested.
You must pick up or arrange for packing and shipping.
For further information contact Sue Gruber, (segruber-at-mtholyoke.edu).
Marian Rice Assoc. Dir. Labs Dept Biological Sciences Mount Holyoke College South Hadley, MA 01075
We have a Sorvall MT-2 ultramicrotome and we are missing the glass knife holder. Does somebody know of a service that provides spare parts for old microtomes? Thanks you very much. Tea
-- *************************************** Tea Meulia Research Scientist and Director Molecular and Cellular Imaging Center Ohio State University/OARDC 1680 Madison Ave. Wooster OH 44691
We need to move a JEOL 35C out of our facility. Your only cost would be shipping. It is fully functional, except it does need a turbo pump. We purchased this new in 1980 and has been under service contract with JEOL service until one year ago when we stopped using it. It was purchase loaded with about every option they had including a four crystal spectrometer. We do have five crystals for that if you are interested. Give me a call or reply to this message if you are interested.
Mark Windland Honeywell Minneapolis, Minnesota 763-954-2845 mark.j.windland-at-honeywell.com {mailto:mark.j.windland-at-honeywell.com}
We are doing immunogold labeling for electron microscopy on adult drosophila eyes. I would like to chat with someone who has done successful labeling on this tissue. Off-line replies are preferred. Thank you very much.
Hong
Hong Yi Emory EM Tel: (404) 727-8692 (Office), (404) 712-8491 (Lab) Fax: (404) 727-3157 Email: hyi-at-emory.edu
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We have a Sorvall MT-2 ultramicrotome and we are missing the glass knife holder. Does somebody know of a service that provides spare parts for old microtomes? Thanks you very much. Tea
-- *************************************** Tea Meulia Research Scientist and Director Molecular and Cellular Imaging Center Ohio State University/OARDC 1680 Madison Ave. Wooster OH 44691
Hello, I've tried a few times now to do cryo SEM after freeze fracturing and cold coating samples. Each time I end up having the fracturing removing the sample from the stage/hat/TEM grid, or ending up melting upon transfer from the freeze fracture device into the cryo SEM wand. I am working with a Hitachi S5000 SEM, a Balzers 301 Freeze fracture unit, a gatan 626 53h31 cryotransfer system, a gatan dry pumping station, a gatan 626DH cryoholder, and either clipring holders for TEM grids, or gatan specimen capsules (626.04352).
I really dislike using TEM grids with samples, fracturing and then trying to load the TEM grids into the gatan clipring holder. I want to try using either larger hats, or the gatan specimen capsules, but I'm either unsure how to mount the hats into the gatan cryoholder, or I'm unsure how to mount the gatan specimen capsules into the Balzers.
Could anyone give me some advice into doing a successful freeze fracture, transfer, and then image with the cryo sem holder? The manuals that came with the Gatan hardware are pretty vague at many many levels. Thanks for the help.
\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\ Gordon Ante Vrdoljak Electron Microscope Lab ICQ 23243541 http://nature.berkeley.edu/~gvrdolja 26 Giannini Hall gvrdolja-at-nature.berkeley.edu UC Berkeley phone (510) 642-2085 Berkeley CA 94720-3330 fax (510) 643-6207 cell (510) 290-6793
I am interested in cutting the seeds of crops such as rice, corn, cotton, and soybeans, and identifying protein, lipid, and starch components in SEM. Can you tell me what approaches and methods I could use?
Good morning We are experiencing some contamination issues in our process. These particulates are poly clothing, dust rubber compounds and other "stuff". With the exception of a few metal particles none of these items are identifiable with the SEM although it is excellent for actual physical structure identification. My question is what optical microscopy techniques do you use for identifying these particulates such as filters, polarizers etc.. I do have most filters at my disposal. I just need the right combination to get positive identification. Also the base the materials are mounted to view these particulates may be important? I am currently using a white background but this may have some effect due to the translucence of some of the materials. Is there a standard background material that you use? Thank you in advance for your input.
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.315 Fax: (770) 487-1460 email: rfowler-at-tdktca.com www.component.tdk.com
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To all subscribers, Due to circumstances beyond my control I must downsize/close my metallurgical engineering consulting practice and move toward retirement. As a first step I am liquidating all of my metallographic laboratory equipment which includes a JEOL T-330 SEM and a full compliment of Buehler metallographic sample preparation/examination equipment. All of the items were purchased new and have been used only by myself in my practice. The T-330 SEM has always been and is currently under a manufacturers (JEOL) service agreement. For anyone interested in
a list of the items, specifications and photographs of same please contact me offline at kmuszar-at-iquest.net.
Karl E. Muszar Jr. M.E.I., Inc. P.O. Box 50366 6161 E. 75th Street Indianapolis, Indiana 46250 (O) 317-842-1906 (F) 317-595-7606
I am looking for a PC file conversion possibility of our old 1993 Voyager EDS sun system (.eds) to a ASCII or text format. Or does somebody know the file structure of data saved in .eds-format ?
Many thanks in advance,
Carsten Ronning
************************************* Dr. Carsten Ronning School of Materials Science and Engineering Georgia Institute of Technology 771 Ferst Drive Atlanta, GA 30332-0245 USA
on leave: II. Insitute of Physics University of Goettingen Bunsenstr. 7-9 37073 Goettingen, Germany ****************************************
We manufacture the JB-4, which was once a Sorvall product. Contact us and we will try to be of assistance.
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: tea meulia [mailto:meulia.1-at-osu.edu] Sent: Sunday, July 08, 2007 10:36 AM To: Microscopy-at-sparc5.microscopy.com
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We have a Sorvall MT-2 ultramicrotome and we are missing the glass knife holder. Does somebody know of a service that provides spare parts for old microtomes? Thanks you very much. Tea
-- *************************************** Tea Meulia Research Scientist and Director Molecular and Cellular Imaging Center Ohio State University/OARDC 1680 Madison Ave. Wooster OH 44691
Any suggestions on how to avoid getting small bubbles in mixed epoxy?
My application is in cementing ultrasonic transducers to glass. The transducers look like very thick washers, about 1-inch in diameter. I am getting a cavitation ringing in the applied voltage (10-volts, at 20-30 KHz). I suspect the bubbles are causing this, but do not know for sure.
I've tried slowly mixing the resin and catalyst beads, but the bubbles persist.
Any suggestions are welcome.
Nathan Haese Moraga, CA
__________________________________ Do you Yahoo!? SBC Yahoo! DSL - Now only $29.95 per month! http://sbc.yahoo.com
After mixing mounting medias in our lab (epoxy, OCT, ProLong, etc.), we place the container under vacuum to get the bubbles to rise out of the media. The amount of time spent under vacuum will depend on how quickly the media dries. This could possibly work for your application.
----------------------------------------------------------------------- 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
You could try putting the epoxy in a vacuum for a short time or curing under vacuum.
Ron L
-----Original Message----- } From: Nathan Haese [mailto:nathanhaese-at-yahoo.com] Sent: Wednesday, July 09, 2003 1:27 PM To: Microscopy-at-sparc5.microscopy.com
List members,
Any suggestions on how to avoid getting small bubbles in mixed epoxy?
My application is in cementing ultrasonic transducers to glass. The transducers look like very thick washers, about 1-inch in diameter. I am getting a cavitation ringing in the applied voltage (10-volts, at 20-30 KHz). I suspect the bubbles are causing this, but do not know for sure.
I've tried slowly mixing the resin and catalyst beads, but the bubbles persist.
Any suggestions are welcome.
Nathan Haese Moraga, CA
__________________________________ Do you Yahoo!? SBC Yahoo! DSL - Now only $29.95 per month! http://sbc.yahoo.com
Nathan: have you tried outgassing your epoxy under vacuum before you use it on the transducers? This works well for embedding and backfilling epoxies. It would depend on the cure time of your epoxy; if it cures too fast (under 15 minutes) this wouldn't be a good thing to try as the epoxy would be on its way to setting before all the bubbles were out. Let me know how it goes.
Nathan Haese 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 } -----------------------------------------------------------------------. } } List members, } } Any suggestions on how to avoid getting small bubbles } in mixed epoxy? } } My application is in cementing ultrasonic transducers } to glass. The transducers look like very thick } washers, about 1-inch in diameter. I am getting a } cavitation ringing in the applied voltage (10-volts, } at 20-30 KHz). I suspect the bubbles are causing } this, but do not know for sure. } } I've tried slowly mixing the resin and catalyst beads, } but the bubbles persist. } } Any suggestions are welcome. } } Nathan Haese } Moraga, CA } } __________________________________ } Do you Yahoo!? } SBC Yahoo! DSL - Now only $29.95 per month! } http://sbc.yahoo.com
-- ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Becky Holdford (r-holdford-at-ti.com) 972-995-2360 972-648-8743 (pager) SC Packaging FA Development Texas Instruments, Inc. Dallas, TX ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
I agree with Christopher. Be carefully for the time under vacuum vs. polymerisation time. Also be careful with the rate of pumping down. You should not have trapped gas in your samples but expanding gas can be destructive. I prefer to "see" the bubbles raising to prevent "boiling" of the resin. If this occur your whole vacuum system is contaminated. I left a sample to cure under vacuum once. Big mistake! You need it to cure in air. The reassure of the atmosphere will close the bubbles that are left over. Depending on the resin/epoxy you can heat it to 40 degrees prior to exposing it to the vacuum to decrease viscosity allowing a shorter time in the vacuum.
-----Original Message----- } From: Nathan Haese [mailto:nathanhaese-at-yahoo.com] Sent: Wednesday, July 09, 2003 7:27 PM To: Microscopy-at-sparc5.microscopy.com
List members,
Any suggestions on how to avoid getting small bubbles in mixed epoxy?
My application is in cementing ultrasonic transducers to glass. The transducers look like very thick washers, about 1-inch in diameter. I am getting a cavitation ringing in the applied voltage (10-volts, at 20-30 KHz). I suspect the bubbles are causing this, but do not know for sure.
I've tried slowly mixing the resin and catalyst beads, but the bubbles persist.
Any suggestions are welcome.
Nathan Haese Moraga, CA
__________________________________ Do you Yahoo!? SBC Yahoo! DSL - Now only $29.95 per month! http://sbc.yahoo.com
If my memory serves me correctly, there is a conversion routine in the Voyager software that converts the file to an emsa format which can then be read by programs such as Excel. I have not used the system in a long time and I can't remember exactly where the command is. The version of the software I used was 2.4.
Good luck.
Colin Veitch
Instrumentation Scientist
Late Stage Innovation Group
CSIRO Textile and Fibre Technology
PO Box 21, BELMONT, Vic. 3216. Australia.
E-mail: colin.veitch-at-csiro.au
Web: http://www.tft.csiro.au
Tel: +61 (0) 3 5246 4000
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The information contained in this e-mail message may be privileged or confidential information. If you are not an intended recipient, you may not copy, distribute or take any action in reliance on it. If you have received this message in error, please telephone CSIRO Textile and Fibre Technology on +61 3 5246 4000.
-----Original Message----- } From: Carsten Ronning [mailto:Carsten.Ronning-at-phys.uni-goettingen.de] Sent: Thursday, 10 July 2003 2:10 AM To: Microscopy-at-sparc5.microscopy.com
Dear All,
I am looking for a PC file conversion possibility of our old 1993 Voyager EDS sun system (.eds) to a ASCII or text format. Or does somebody know the file structure of data saved in .eds-format ?
Many thanks in advance,
Carsten Ronning
************************************* Dr. Carsten Ronning School of Materials Science and Engineering Georgia Institute of Technology 771 Ferst Drive Atlanta, GA 30332-0245 USA
on leave: II. Insitute of Physics University of Goettingen Bunsenstr. 7-9 37073 Goettingen, Germany ****************************************
*From: "Christopher S. Zurenko" {czurenko-at-mail.khri.med.umich.edu} *Organization: Kresge Hearing Research Institute *To: Microscopy-at-sparc5.microscopy.com *Date sent: Wed, 09 Jul 2003 13:59:34 -0400 *Subject: Re: Epoxy: how to avoid small bubbles *Send reply to: czurenko-at-umich.edu *Priority: normal
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Well, in my practice we did once something like this and the oil in rotary pump had to been replaced. However it was big volume of epoxy so maybe in case of small one it wouldn't be so serious damage.
Best regards,
Witold Zielinski
*After mixing mounting medias in our lab (epoxy, OCT, ProLong,
*etc.), we place the container under vacuum to get the bubbles to rise *out of the media. The amount of time spent under vacuum will *depend on how quickly the media dries. This could possibly work for *your application. * *----------------------------------------------------------------------- *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 * * * :) :) :) :) :) :) :) :) :) :) :) :) :) :) :) :) ;)
Witold Zielinski, Ph.D. Warsaw University of Technology Department of Materials Science and Engineering 02-507 Warszawa, Woloska 141 POLAND
} I am looking for a PC file conversion possibility of our old 1993 } Voyager EDS sun system (.eds) to a ASCII or text format. Or does } somebody know the file structure of data saved in .eds-format ? } I have written a multi-purpose application to read all Noran Voyager (and Vantage) file formats to enable, among a lot of other features, data to be exported in ascii format. Noran do not (will not?) give any information on their file structures so this has been the result of many hours of hacking. Unfortunately, I have found that some files from other users do not read on my application. Send me some of your files (.eds, .grey, etc) and I will see if they read OK.
David Vowles Electron Microscope Unit Dept of Materials Science and Metallurgy University of Cambridge Pembroke St Cambridge UK CB2 3QZ Tel: +44 1223 334325 Fax: +44 1223 334567 Email: djv23-at-cam.ac.uk
It has been my experience doing this that one needs to evacuate the container (vacuum oven, desiccator, whatever)by slowly opening the valve, allowing the bubbles rise to the top, close the valve, etc, effectively cycling the system until the valve can be completely opened to permit removal of the few remaining bubbles.
Roger Moretz, Ph.D. BI Pharmaceuticals, Inc Dept of Toxicology Ridgefield, CT
-- Where the world is only slightly less weird than it actually is. } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } *From: "Christopher S. Zurenko" {czurenko-at-mail.khri.med.umich.edu} } *Organization: Kresge Hearing Research Institute } *To: Microscopy-at-sparc5.microscopy.com } *Date sent: Wed, 09 Jul 2003 13:59:34 -0400 } *Subject: Re: Epoxy: how to avoid small bubbles } *Send reply to: czurenko-at-umich.edu } *Priority: normal } } *------------------------------------------------------------------------ } *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, } } Well, in my practice we did once something like this and the oil in } rotary pump had to been replaced. However it was big volume of epoxy } so maybe in case of small one it wouldn't be so serious damage. } } Best regards, } } Witold Zielinski } } } *After mixing mounting medias in our lab (epoxy, OCT, } ProLong, } } *etc.), we place the container under vacuum to get the bubbles to rise } *out of the media. The amount of time spent under vacuum will } *depend on how quickly the media dries. This could possibly work for } *your application. } * } *----------------------------------------------------------------------- } *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
If you have access to an acoustic microscope [SonooScan or Sonix] you can image through the transducer and detect voids in the epoxy non-destructively. I assume the transducer is a solid piezoelectric device.
Peter Tomic Agere Systems
-----Original Message----- } From: Nathan Haese [mailto:nathanhaese-at-yahoo.com] Sent: Wednesday, July 09, 2003 1:27 PM To: Microscopy-at-sparc5.microscopy.com
List members,
Any suggestions on how to avoid getting small bubbles in mixed epoxy?
My application is in cementing ultrasonic transducers to glass. The transducers look like very thick washers, about 1-inch in diameter. I am getting a cavitation ringing in the applied voltage (10-volts, at 20-30 KHz). I suspect the bubbles are causing this, but do not know for sure.
I've tried slowly mixing the resin and catalyst beads, but the bubbles persist.
Any suggestions are welcome.
Nathan Haese Moraga, CA
__________________________________ Do you Yahoo!? SBC Yahoo! DSL - Now only $29.95 per month! http://sbc.yahoo.com
This year there is a program for spouses and companions at M&M. (Details on page 34 of EXPO, if you have it - mine arrived today).
So far only a few people have registered for this program and we will have to cancel it unless more people sign up. If you or your companion intend to take part in this program but have not yet registered, please do so immediately.... Or else...
My apologies for the threatening and harassing tone but we have to make a decision at once.
Alwyn Eades -- .......... Alwyn Eades Department of Materials Science and Engineering Lehigh University 5 East Packer Avenue Bethlehem Pennsylvania 18015-3195 Phone 610 758 4231 Fax 610 758 4244 jae5-at-lehigh.edu
You'll need freezing methods for lipids -- cryo fixation, cryo coating and cryo SEM, or you'll have to do these by 'subtraction'. That is, do something like a chloroform extraction, and look at the result, assuming the holes are where the lipids were. Dangerous, because the holes could've been air or water, too. And you'd be assuming that the chloroform (and EtOH during dehydration) removed *all* of the lipids. Proteins can be done somewhat on morphology, but also by labeling: use gold-conjugated antibodies to the proteins of interest. Starches typically are as smooth grains, so they're pretty easy to ID by morphology. At least the ones I've looked at are. But if you need to ID "starch A" vs "starch B" ... well. There might be something in the literature, but I don't know of a method.
Phil
} I am interested in cutting the seeds of crops such as rice, corn, } cotton, and soybeans, and identifying protein, lipid, and starch } components in SEM. Can you tell me what approaches and methods I } could use? } } Thanks, } } Raja Rao
-- 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)
You might also try centrifuging the mixed epoxy, which would avoid the pump problems other listers have mentioned.
------------------------------------------------ Kevin Frischmann Microscopy & Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
At 03:17 PM 7/9/03 -0400, Ron L'Herault wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Good Day. On behalf of the Local Arrangements Committee of the 2003 Microscopy and Microanalysis Meeting in San Antonio, Texas, I would like to invite you to the 2003 Golf Outing to be held Sunday, August 3 at the Pecan Valley Golf Club. Pecan Valley is a majestic golf course, located only six miles from downtown San Antonio and the Riverwalk. Pecan Valley was the site of the 50th Anniversary PGA Championship in 1968 when Julius Boros edged Arnold Palmer on the 18th hole and has also hosted three Texas Opens. It has been rated in Golf Digest¹s Top 50 Public Courses and #1 Public Golf Course in the State of Texas for 2002 (http://www.golftexas.com/pecan1.htm Rating:74.5 - Slope: 136 - Yards:7,071). The cost will be $70.00 and will include greens fees, cart, transportation to and from the Convention Center, Driving range, lunch buffet and awards banquet. We will leave the Convention Center at 6:45 - 7:00am (to beat the heat). Callaway club rental is available onsite at $45.00 per set. Another addition to this year¹s outing is the inclusion of various types of sponsorships. We are also looking for additional raffle prizes and / or giveaways for the event. Any company or individual that becomes a sponsor will be promoted in the following ways: * On the hole of their choice (for hole sponsorships) * On the lunch placemats * In each ³goodie² bag that every golfer will receive at the course
If you are interested in playing in the outing, please register on the web at http://www.microscopy.com/MSAMeetings/MMMeeting.html or contact me directly (msanders-at-cbs.umn.edu). If you have a foursome, please let me know and include all four names. If you do not have a foursome, that is fine as well we can pair you into a foursome at the course. If you are interested in becoming a sponsor, please contact me directly and I can get further details to you.
I would also like to announce that there will be 2 ³free² slots for students (preferably). These will be in the name of a former MSA member, golfer and a good friend of mine who passed away a few years ago, Joe Polak.
Contact Mark Sanders with any questions (msanders-at-cbs.umn.edu)
Thanks in advance for your consideration and participation in this matter!
At Pecan Valley you can truly ³Play where Champions Have Left Their Footprints.²
My Best Regards,
Mark Sanders MSA/MAS Golf Tournament at the Pecan Valley Golf Club
I have no suggestions from personal experience, but interestingly, shortly after reading your question, I opened up my most recent copy of Scanning, and the first paper was:
Kohler A, Høst V, Enersen G, Ofstad R: Identification of Fat, Protein Matrix, and Water/Starch on Microscopy Images of Sausages by a Principal Component Analysis-Based Segmentation Scheme Scanning 25, 3, 109-115 (2003)
It's not an SEM paper, but if you have access to a fluorescence scope, you might try it (at least for checking SEM results).
In short, samples were sectioned (10um) with a cryostat, and stained with Nile red (9-diethylamino-5H-benzophen-oxazin-5-one) dissolved in acetone (0.1 mg/ml overnight at 4 deg C). Stained sections were examined using a fluorescence microscope with a blue excitation filter (450-490nm). After staining, lipids show strong fluorescence, proteins weak fluorescence, and starch near zero fluorescence.
Granted, sausages and seeds are a bit dissimilar, but it seems like it ought to work.
------------------------------------------------ Kevin Frischmann, Laboratory Manager Microscopy & Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
At 07:59 AM 7/9/03 -0500, RAO, RAJA S [AG/1000] wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I don't think you can avoid getting bubbles in epoxy, but you can remove them after mixing if it's not a quick-setting epoxy. Place the mixed epoxy under a light vacuum for about five to ten minutes, or until there are no more small bubbles rising in the mix. If the vacuum is too strong, the epoxy will start to outgas - large bubbles rising from the bottom of the vessel, which interferes with the later polymerisation.
Lesley Weston.
on 09/07/2003 10:27 AM, Nathan Haese at nathanhaese-at-yahoo.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 } -----------------------------------------------------------------------. } } } List members, } } Any suggestions on how to avoid getting small bubbles } in mixed epoxy? } } My application is in cementing ultrasonic transducers } to glass. The transducers look like very thick } washers, about 1-inch in diameter. I am getting a } cavitation ringing in the applied voltage (10-volts, } at 20-30 KHz). I suspect the bubbles are causing } this, but do not know for sure. } } I've tried slowly mixing the resin and catalyst beads, } but the bubbles persist. } } Any suggestions are welcome. } } Nathan Haese } Moraga, CA } } } } } __________________________________ } Do you Yahoo!? } SBC Yahoo! DSL - Now only $29.95 per month! } http://sbc.yahoo.com }
My practice for years has been to mix the components less the catalyst/accelerator and place under vacuum (15-20 in Hg) overnight or for no less than 4 hr. If the room is too cold, I precondition the vacuum oven in which I have done this to about 22 degrees C (i.e., just a little warm). To prevent excessive air entrainment, I use only 2 sticks to stir the materials and I do so slowly without any attempt to 'beat' the mixture as one does with egg whites. In some cases, I have mixed by very slow rolling or tumbling for 4-6 hr. NEVER shake to mix, because that only ensures air entrainment, and remember, low temperature will exacerbate the solution of atmospheric gases. The final alternative is to place large volumes under light vacuum ( {15 in Hg) for a sufficiently long time (e.g., 24 hr) to insure that they are clear of entrained air. Then mix and expose to light vacuum again. One manufacturer suggests warming all components to 60 degrees before mixing.
These amount to an admission that I have had the same problems. The issue with your problem is that you must be particularly careful to degas the system because you appear to be using it to build a sandwich on the glass.
Finally, I believe you should use a very slow cure. Rapid curing is will known to cause bubbles. The best example is methacrylate plastics polymerized in bulk. In most cases the 'pour' is not made until the amount of monomer has been significantly reduced.
Suggestion. If you were to use a 90 min epoxy (two part available at a local hardware store) you would notice that both components exhibit a lot of viscosity. You will also note that you hardly ever get bubbles in the seal if even a little care is exercised. You might want to try a simple solution like that. If you are using EPON, I would suggest a partial cure at 45 degrees for 4-6 hr. At that temp a lot of degassing will have occurred and polymerization will be well on its way. Then transfer only what you need to your job and complete the cure at 55. In this manner, the HOT part of polymerization would likely be done during the preliminary cure and no cavities should be produced. Then, at the end, turn the oven off and let it slowly come to room temperature before you remove your devices.
Rationale/Guess? Consider the two substances, glass and transducer material, and their coefficients of expansion. Most resins remain somewhat fluid at higher temperatures, if two layers with different coefficients of expansion are connected by a flowing plastic at 60 degrees and the system is cooled rapidly, what happens when, and if, the polymer reaches its glass transition, and tries to hold two materials that are contracting at different rates? Cavities, I bet! Perhaps you want a polymer that will survive the stress at operating temperatures above its glass transition.
http://www.warmglass.com/COESummary.htm
http://www.psrc.usm.edu/macrog/tg.htm
http://www.dymax.com/pdf/SPIE_Paper_Appendix.pdf
If I have gotten it all wrong, please feel free to reply with more info about the specific epoxy you are using.
Frederick C. Monson, PhD Center for Advanced Scientific Imaging Mail Drop: Geology West Chester University West Chester, PA, 19383 http://darwin.wcupa.edu/casi/ Phone/FAX: 610-738-0437
-----Original Message----- } From: Nathan Haese [mailto:nathanhaese-at-yahoo.com] Sent: Wednesday, July 09, 2003 1:27 PM To: Microscopy-at-sparc5.microscopy.com
List members,
Any suggestions on how to avoid getting small bubbles in mixed epoxy?
My application is in cementing ultrasonic transducers to glass. The transducers look like very thick washers, about 1-inch in diameter. I am getting a cavitation ringing in the applied voltage (10-volts, at 20-30 KHz). I suspect the bubbles are causing this, but do not know for sure.
I've tried slowly mixing the resin and catalyst beads, but the bubbles persist.
Any suggestions are welcome.
Nathan Haese Moraga, CA
__________________________________ Do you Yahoo!? SBC Yahoo! DSL - Now only $29.95 per month! http://sbc.yahoo.com
I have a guy about to go on a cruise to collect coccolithophorids. He wants to bring them back and examine them using SEM. He asked for advice on collection and preservation techniques.
I guessed at standard SEM fixation with glutaraldehyde and keep track of the pH to prevent dissolution of the coccoliths. It is important that they be preserved intact, ie. individual cells retain all their coccoliths and not fall apart. He wants to count coccoliths/cell from different collecting stations.
Any coccolithophorid experts out there with practical experience on how to bring them back in one piece, or at least dead but still looking alive?
Thanks
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
The Journal of Microscopy publishes top quality review articles, original research papers, short technical notes, short communications, rapid publications and letters to the Editors, covering all aspects of microscopy and high-energy in situ beam analysis. Papers that emphasize the application of microscopical techniques or specimen preparation procedures in an investigation are also welcome.
} From August 1st, 2003 it will be possible to submit your papers online to the Journal of Microscopy, simply go to http://jmi.manuscriptcentral.com and follow the online instructions.
This system will enable us to cut down on decision and publication time, whilst ensuring the same high standards of peer review.
We look forward to receiving your contributions to this ever-growing journal.
Dr Ilaria Meliconi, Executive Editor Journal of Microscopy 37/38 St Clements, Oxford OX4 1AJ, UK Tel +44 (0)1865 248768 Fax +44 (0)1865 791237 http://www.blackwell-science.com/jmi
Note to anyone with experience in chick embryonic tissue and/or eye tissue, I recently did some TEM on chick embryonic conjunctival epithelium and mesenchyme and discovered some odd cells that need identifying.
The cells are generally in clumps, with regular edges, very very few organelles (a few mitochondria and many many free ribosomes) - no other organelles can be identified (no RER or SER or anything else). The cells within the mesenchymal tissue. We suspect the cells have recently divided and are beginning to differentiate but we are not sure into what. Anyone with insight into what these cells could be, please email me at {mailto:tfranzod-at-dal.ca} tfranzod-at-dal.ca - I can also send a few images to you if required/wanted. Dr. Tamara Franz-Odendaal, Halifax, Canada
Hello. We found a staining procedure for ruthenium hexaammine trichloride and we would like to try it. Has anybody used this chemical in their labs? I have questions regarding safety and handling of it. Is it easy to dissolve? MSDS says that it is highly reactive and air sensitive. How do you store it when a bottle is open?Label indicates that it should be stored under nitrogen, so what does this mean? How do you dispose of it? Can stock solution be made ahead of time and safely store in the fridge? Thanks Dorota
} Microscopy and Microanalysis 2003 } San Antonio } } This year there is a program for spouses and companions at M&M. } (Details on page 34 of EXPO, if you have it - mine arrived today). } Another meeting event will interest spouses who are teachers. Project MICRO will present a workshop on the use of "Microscopic Explorations" in the classroom. It's SC-11, Sunday Aug. 3,from 2-5. Unlike most Sunday courses, it's free - and it's an outstanding opportunity. Look it up! -- Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
Has anyone done EDX analysis and/or EM imaging on gold corrosion or dendritic growth in the presence of a halogen like bromine? I'd like to compare SEM's and spectra with anyone involved in this.
I have what looks like to be the complete document set (instructions, schematics, spare parts list, etc.; multiple copies of some or all) for the Philips 200 TEM. Anybody interested?
Be the first caller (well, emailer) for these fine collectibles and I'll throw in two (two!) plate magazines complete with cassettes and three (three!) receiver boxes, all from the EM 200.
But wait, there's more! At no extra charge, you also receive the vintage EM 200 "Simplified Cross-Section" poster, suitable for framing or covering up that nasty hole in your wall.
Don't wait! All of this can be yours, for a modest shipping fee (I'm guessing maybe 30 pounds worth of stuff).
Contact me off-list. If I don't have any bites in a week or so, I'm going to toss the whole lot.
Jim
--
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
I read your email. First of all you need to state the epoxy you are using. I am sure it is not Epon. You need to test each component under vacuum to see what happens. Apply the vacuum in stages to see how the bubbles form and their sizes.
Below is a paragraph from an article I wrote for Microscopy Today about capillary forces and epon epoxy but have not sent in yet. Anyway below is the part I want you to consider. The capillary forces part would seem to be irrelevant because you are not wetting microporous membranes or nano-porous agglomerates.
Referring to the Epon Clone epoxy component DDSA for some perspective:
"Try this experiment. Pour DDSA into a long test tube and fill it about half full. Put the test tube in a beaker and pull a vacuum on both using a bell jar. You will notice the formation of small bubbles. These are entrained or dissolved air. Before they reach the surface and break, release the vacuum. Notice what happens to the air bubbles. They just shrink in place. Next immediately pull a vacuum and notice they reappear in the same place. Let them rise to the top surface, combine to form larger bubbles and then break them by applying and releasing the vacuum. Eventually only larger bubbles will form under increased rotary pump vacuum. These are not air bubbles but vaporizing DDSA or whatever other high boiler is in DDSA. After you only get the large bubbles, release the vacuum. Using a micro-spatula, stir up the DDSA and entrain some air. With the spatula, break the large bubbles that rise to the surface. Pull a vacuum on the DDSA again. Small bubbles will form again. The larger DDSA bubbles will follow these smaller bubbles. This will give you insight as to what really happens in a microporous membrane or porous agglomerated powder during vacuum impregnation." My DDSA from Pella foams under vacuum. I think all DDSA does but I haven't tried them all.
I can't know how your epoxy system will react but try this experiment. It will tell you how your epoxy components and air in your components behave under vacuum. I realize you are probably using a small amount of epoxy as a thin film. Use a larger amount of each component to determine where the bubbles are coming from in your samples.
What about the transducer surface as a source? Do you have sub-micron particles forming nucleation sites in a non-transparent epoxy? Can you try that special automotive glass adhesive used for attaching rear view mirrors? It is very thin but my not be strong enough for you. In any case, it might last long enough to show the elimination of the ringing.
I hope this gives you some help or gives you some ideas that lead you to a solution.
Sincerely,
Paul Beauregard Senior Research Associate Electron Microscopy PPG Industries
} List members, } } Any suggestions on how to avoid getting small bubbles } in mixed epoxy? } } My application is in cementing ultrasonic transducers } to glass. The transducers look like very thick } washers, about 1-inch in diameter. I am getting a } cavitation ringing in the applied voltage (10-volts, } at 20-30 KHz). I suspect the bubbles are causing } this, but do not know for sure. } } I've tried slowly mixing the resin and catalyst beads, } but the bubbles persist. } } Any suggestions are welcome. } } Nathan Haese } Moraga, CA } } } } } __________________________________ } Do you Yahoo!? } SBC Yahoo! DSL - Now only $29.95 per month! } http://sbc.yahoo.com } } }
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-- [ From: Garber, Charles A. * EMC.Ver #3.1a ] --
Dorita Wadowska wrote: ============================================================================ ==== We found a staining procedure for ruthenium hexaammine trichloride and we would like to try it. Has anybody used this chemical in their labs? I have questions regarding safety and handling of it. Is it easy to dissolve? MSDS says that it is highly reactive and air sensitive. How do you store it when a bottle is open?Label indicates that it should be stored under nitrogen, so what does this mean? How do you dispose of it? Can stock solution be made ahead of time and safely store in the fridge? Thanks ============================================================================ ====== So far as I know, all of these (organic) ruthenium compounds are quite unstable. That is why it is difficult to find someone who would ship for example, ruthenium tetroxide.
For staining, we have offered for some years a ruthenium tetroxide staining kit. It is described on URL www.2spi.com/catalog/chem/chem1c.shtml
The concept of the kit is that is allows you to make up tiny amounts of the tetroxide, in tiny amounts, which is the "active" species that does the staining. The ingredients that are used to produce the much more dangerous (actually potentially explosive tetroxide) are relatively inert and the amount of tetroxide that is produced is quite small, thereby minimizing any risk to the user.
If you think that there is some kind of specificity of the staining unique to ruthenium hexaammine trichloride, relative to the more straight-forward tetroxide, I (and probably others) would be very eager to hear more about it
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Hi all;
I am interested in quick freezing and freeze substitution of human Red Blood Cells (I am looking for high quality membrane architecture). I have the freezing all set up, but am looking for the freeze sub protocol. Could anyone send me one, or point me in the correct direction?
Thanx David ____________________
David Elliott Ph.D.
Yale University School of Medicine 333 Cedar Street PO Box 208022 New Haven, CT 06520-8022
Can you clarify: is Ruthenium tetroxide unstable when it is in solution, or only in the crystalline form? Thanks.
Marie
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Dr. Marie E. Cantino Director, Electron Microscopy Laboratory Associate Professor of Physiology and Neurobiology University of Connecticut Unit 3242 Storrs, CT 06269-3242 Phone: 860-486-3588 Fax: 860-486-6369
Regarding ruthenium tetroxide, I use the stain routinely for polymer microscopy and make it fresh as needed. An alternative (not better or worse, just different) to the periodate preparation of which Charles speaks was first described for use in microscopy by Montezinos. I have only slightly modified his procedure, using 10 % sodium hypochlorite instead of bleach. I refer you to G. M. Brown and J. H. Butler, "New method for the characterization of domain morphology of polymer blends using ruthenium tetroxide staining and low voltage scanning electron microscopy (LVSEM)", Polymer 38 (15), 3937 (1997). The appendix provides detailed descriptions for preparing and using the stain for low voltage SEM as well as TEM, and how to safety handle and dispose of the ruthenium tetroxide. Also, the Montezinos paper is referenced in this paper.
Feel free to contact me off-line with any questions.
Cheers,
"The statements and opinions expressed here by Gary M. Brown represent neither those of ExxonMobil Corporation nor its affiliates."
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
"Garber, Charles A." To: MICROSCOPY BB {Microscopy-at-sparc5.microscopy.com} {cgarber-at-2spi.com} cc: Subject: Ruthenium staining
07/12/03 07:16 PM
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-- [ From: Garber, Charles A. * EMC.Ver #3.1a ] --
==== We found a staining procedure for ruthenium hexaammine trichloride and we would like to try it. Has anybody used this chemical in their labs? I have questions regarding safety and handling of it. Is it easy to dissolve? MSDS says that it is highly reactive and air sensitive. How do you store it when a bottle is open?Label indicates that it should be stored under nitrogen, so what does this mean? How do you dispose of it? Can stock solution be made ahead of time and safely store in the fridge? Thanks ============================================================================
====== So far as I know, all of these (organic) ruthenium compounds are quite unstable. That is why it is difficult to find someone who would ship for example, ruthenium tetroxide.
For staining, we have offered for some years a ruthenium tetroxide staining kit. It is described on URL www.2spi.com/catalog/chem/chem1c.shtml
The concept of the kit is that is allows you to make up tiny amounts of the tetroxide, in tiny amounts, which is the "active" species that does the staining. The ingredients that are used to produce the much more dangerous
(actually potentially explosive tetroxide) are relatively inert and the amount of tetroxide that is produced is quite small, thereby minimizing any risk to the user.
If you think that there is some kind of specificity of the staining unique to ruthenium hexaammine trichloride, relative to the more straight-forward tetroxide, I (and probably others) would be very eager to hear more about it
Dear Sir; I worked about stucture analysis of single crystal with rigaku afc7s single crystal diffractometer in my master thesis.Now I have been studying shape memory alloys since 2000 in my doctorate thesis and responsible user from JEM3010 Transmission Electron Microscopy since 1999.I was heard "tem single crystal sample investigation to Darmstat university in germany" by my teacher in University of Ankara.I put on grid my single crystal sample and I observe.But In my opinion this method may be dangerous if sample fall in microscopy. Sincerely.
***************************************************** ***************************************************** ** Research Assistant ** ** PhD.Student ERDEM YASAR ** ** University of Kirikkale ** ** Department of Physics ** ** Electron Microscopy Laboratory ** ** 71450 KIRIKKALE/TURKEY ** ** erdem.yasar-at-physics.org ** ** tem_sem-at-hotmail.com ** ** http://turkuaz.kku.edu.tr/~yasar/erdem.html ** ***************************************************** *****************************************************
On Fri, 25 Apr 2003, darryl krueger wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } } } } From: darryl krueger {dkruege-at-clemson.edu} } } Subject: TEM single crystal sample } } } } I have a friend that is doing Diffraction, who is looking for a source for } } single crystals on TEM grids. If there is source someone knows of could } } you please let us know. TIA } } } } Darryl } } }
HELP!!! I have an old MT 5000 and a MT2B both in need of repair. Does anyone have a number for someone working on these microtomes? I am located in the south Texas area. Thanks in advance.
Lauren E. Chesnut Technical Director Electron Microscopy Lab UTHSCSA Dept. of Pathology MC-7750 7703 Floyd Curl Drive San Antonio, TX. 78229-3900 (210)567-4052 CHESNUTL-at-UTHSCSA.EDU
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Hi everybody. Ruthenium that I am using is a hexamine trichloride. I already dissolve it in cacodylate buffer and it almost immediately turns yellow upon addition of glutaraldehyde. Alter a few hours is dark brown. I am following procedure of Michael D. Buschmann et al "Ruthenium hexaammine trichloride chemography for aggrecan mapping in cartilage is a sensitive indicator for matrix degradation", The Journal of Histochemistry and Cytochemistry, vol 48(1): 81-88, 2000. The information I was able to retrieve form the Internet sites indicates that it is very reactive compound, degrades under air and is slightly radioactive hence my concerns. Thanks for your inputs. Dorota
we have to prepare a polymere (elastomere) which contains Nanoparticle (SiO2) in low concentration. The aim of the investigation is to get a nanoparitcle size distribution?
Hi All, It is a beast, isn't it? We have had somewhat consistent results with the new film by watching the temperature carefully and using a constant gentle bubbling with my nitrogen burst system (rather that the 2 second burst every 10 seconds that Kodak recommends). I am also careful to make sure that the film does not rest on the support bar that runs the length of the film racks, but is raised a bit to allow fluid flow. Of course I realize that I may just have a "good" box and all bets may be off with the next one, but since we've have nice results 4 times in a row, I thought it was worth passing it on. Lee -- Lee Cohen-Gould Electron & Optical Microscopy Facilities Weill Medical College of Cornell U. (212)746-6146 Rms A-105, LC-207
Werner, you omit the important point of likely particle size, but if it is nanoscale, let's assume it is somewhere in the 30-100nm range. I'm also assuming you plan get the distribution via low voltage FE-SEM. The simplest method by far is visit your local medical people at NMI and ask for the person who runs their diamond knife ultramicrotome (used for preparing ultrathin sections of tissue and the like for TEM) to prepare and 'face off' (ie put a flat on the end) a block of your material. Many polymers are soft enough to require cryosectioning, but that is standard for medical applications (and many biological ones as well, so they should have that capability. SInce you say the SiO2 is 'low concentration', wear of the diamond knife edge should not be a problem, which is something that the person will immediately be concerned about. I recommend a fairly high sectioning speed since you want to fracture your way through the particles, not pull them out of the polymer.
If you want thin TEM sections (by not having access to an FE-SEM), matters are trickier in general re picking up the section without having all the sectioned particles fall out, but then you would still have the holes as a guide. If the particles are {30nm in size, you've got an additional problem, as mixed phase materials tend to be difficult to section at thicknesses less than about 20-30nm, at least in the few soft matrix/hard particle systems that I've looked at. But it would be worth a try if you can find that microtome and a willing operator. I can't see mechanical polishing being very successful with your system, but if you have access to a focused ion beam (FIB) system, that could be tried as well.
Best of luck and let me know how it turns out.
Tom
Dr. Tom Malis Scientist Advisor Mineral Technology Branch Natural Resources Canada 555 Booth St., Ottawa, Ontario K1A 0G1 Tel.: (613) 995-7358 FAX: (613) 947-6606 malis-at-nrcan.gc.ca
-----Original Message----- } From: Werner Dreher [mailto:Dreher-at-nmi.de] Sent: Tuesday, July 15, 2003 8:11 AM To: Microscopy-at-sparc5.microscopy.com
Hi all,
we have to prepare a polymere (elastomere) which contains Nanoparticle (SiO2) in low concentration. The aim of the investigation is to get a nanoparitcle size distribution?
Kodak Eastman Motion Picture Film 5302 fine grain release positive film cat 166 7229.
At 09:20 15-07-2003 -0400, Eleana Sphicas wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Postdoctoral Position Transmission Electron Microscopy Acadia University
The Center for Microstructural Analysis is seeking a postdoctoral research associate to perform transmission electron microscopy of ferromagnetic shape memory alloys. The work will involve collaboration with an interdisciplinary group (physicists, metallurgists and chemists) at Acadia University, Dalhousie University and Defence Research and Development Canada. Duties will include the operation of a Philips CM30 300 kV analytical STEM equipped with EDS and PEELS and specimen preparation using a Hitachi FIB 2000A. Occasional travel to a reactor facility to assist in the performance of neutron diffraction experiments may also be required.
The successful applicant will have a doctoral degree in Physics, Metallurgy or Materials Science with demonstrated expertise using TEM. Specific knowledge of high resolution imaging techniques and prior experience working with shape memory alloy systems would be considerable assets. Applicants must also possess good organizational and communication skills.
The position is funded for 18 months at $41000 CDN per annum, including salary and benefits. The start date is planned as soon as possible after September 1, 2003.
Review of applications will begin July 25, 2003. Interested applicants should submit a cover letter highlighting their qualifications, a CV including a list of publications, and contact information for three references to:
Dr. Craig Bennett Physics Department Acadia University Wolfville, NS B4P 2R5
On Sunday, July 13, 2003, at 12:03 PM, Erdem Yasar wrote:
} I worked about stucture analysis of single crystal with rigaku } afc7s } single crystal diffractometer in my master thesis.Now I have been } studying } shape memory alloys since 2000 in my doctorate thesis and responsible } user } from JEM3010 Transmission Electron Microscopy since 1999.I was heard } "tem } single crystal sample investigation to Darmstat university in germany" } by } my teacher in University of Ankara.I put on grid my single crystal } sample } and I observe.But In my opinion this method may be dangerous if sample } fall in microscopy. } Dear Erdem, Depending on the nature of your single-crystal specimen, there may be reliable methods either to attach it to a grid or to sandwich it between two grids (or halves of a folding grid). For a crystal that is only about 10nm thick and a few hundred nm wide--reasonable dimensions for a SAED structural study--there might be little problem if the sample drops, depending on where it ends up, which depends on the construction of your microscope. It is, of course, best if the specimen stays on the grid. 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
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-- [ From: Garber, Charles A. * EMC.Ver #3.1a ] --
Werner Dreher wrote: =================================================== we have to prepare a polymere (elastomere) which contains Nanoparticle (SiO2) in low concentration. The aim of the investigation is to get a nanoparitcle size distribution? ======================================================= To do this reliably, and with the greatest efficiency, you will have to separate the isolate and then concentrate the nanoparticles from the polymer matrix and this is most easily done by putting the filled polymer into a plasma etcher using oxygen (which will produce an oxygen plasma, which will etch away the organics) leaving the inorganic nanoparticles behind.
We would recommend the SPI Supplies Plasma Prep II etcher for this kind of work, but there are other etchers on the market that should do the same thing. The Plasma Prep II etcher is described on URL http://www.2spi.com/catalog/instruments/etchers1.shtml
You can then pick up the nanoparticles on either a carbon supported TEM grid or if the grain size of the carbon is such that it interferes with the analysis of the SiO2 nanoparticles, you can consider using the SPI silicon nitride membrane window grids, see URL http://www.2spi.com/catalog/instruments/silicon-nitride.shtml since they are both structureless and featureless.
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 ######################## ============================================
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listers
St. Lawrence University is seeking to fill a full-time, 12 month academic support staff position as a MICROSCOPY SPECIALIST. A masters degree or equivalent experience is required as is experience with confocal microscopy as well as electron microscopy (TEM and/or SEM). Mechanical and laboratory aptitude, computer experience, a desire to learn and teach new methodologies, and a positive work ethic is sought. The successful candidate will help oversee a developing interdisciplinary, multi-user microscopy/imagery center, will assist faculty and student researchers in advanced microscopy techniques, help teach microscopy methods and provide for the routine maintenance of the instrumentation infrastructure. The major instruments at the facility are on service contracts and the candidate will be expected to develop good working relationships with the professional service personnel. The facility is housed in the biology department and the successful candidate will also serve other science departments.
Applicants should send a letter of application, a current curriculum vitae (including references), a statement of any relevant research and teaching interests related to microscopy, and have three letters of recommendation forwarded to Dr. T. Budd, Biology Department, St. Lawrence University, Romoda Drive, Canton, NY 13617. The search committee will begin reviewing applications on July 21 and the search will remain open until filled.
-- Dr. T. Budd Chair of Biology St. Lawrence University Canton, NY 13617 Phone = 315-229-5640 Fax = 315-229-7429 E-mail = tbudd-at-stlawu.edu
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (sswaffe-at-abv.bg) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 16, 2003 at 00:38:42 ---------------------------------------------------------------------------
Email: sswaffe-at-abv.bg Name: Veselin
Organization: 54
Education: 6-8th Grade Middle School
Location: Sofia Bulgaria
Question: How can I clean my immersion oil lenses after work with immersion oil ?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (alessandro-at-polymer.kth.se) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 16, 2003 at 06:15:00 ---------------------------------------------------------------------------
Question: Dear Sir I would be grateful if you can provide some advice in order to carbon coat my replicas for TEM. I am using a Jeol Vacuum evaporator. I would like to know: -which kind of geometry should I use for the carbon rods? -how big should be the distance between the tips? -which current density should I use (25-30A as for evaporating metals?)? Thank you very much Regards Alessandro Matttozzi
I have the possibility of obtaining an Cambridge S90 that includes the factory installed Digital Image Store, that will allow Frame Averaged, Frame Integrated or Line Integrated video at tv rate.
Does anyone have any experience of connecting the tv output of the SEM to a PC TV card to allow the image to be saved on the computer?
Or do I need to spend more money in buying a digital imaging system.
I welcome any suggestions or recommendations
many thanks
Kevin
Electron Microscope unit School of Biological Sciences University of Aberdeen Aberdeen AB24 2TZ
Tel 01224-272847 Fax 01224-272396 ------------ Kevin Mackenzie k.s.mackenzie-at-abdn.ac.uk
Light-Microscopist There is a full-time opening for a core facility light/confocal microscopist available immediately. The encumbent must demonstrate total facility with current technology and sample preparations. They will be responsible for 1) providing direct hands-on access to light microscopy in a core facility setting, 2) overseeing daily facility operations, 3) consulting with users on experiment design, materials and methods, and instrument optimization and assist in data acquisition and analysis, 4) troubleshooting problems with microscopes or samples and 5) assuring instrument integrity. Must maintain a current understanding of the microscopy field as necessary to implement new technologies on campus. Applicants must have a strong desire to participate actively as a member of a team, possess excellent written and oral communication skills, be comfortable working independently, communicate effectively with service users to understand their microscopy needs, be facile with PC and Mac computer platforms and demonstrate effective troubleshooting skills. M.S. in biological sciences with five years research experience in molecular biology, immunohistochemistry and cytology is required. Previous work in a core facility is preferred.
The Jackson Laboratory is one of the world's foremost centers for mammalian genetics research. Located in Bar Harbor, Maine, the lab is adjacent to Acadia National Park. Mountains, ocean, forests, lakes, and trails are all within walking distance. If you are looking for a more natural environment, this could be the opportunity you've been searching for.
Interested applicants should send cover letter and resume to: jax/210 Human Resources, Box 27 The Jackson Laboratory 600 Main Street Bar Harbor, Maine 04609 Fax: (207) 288-6106 Email (preferred option): jobs-at-jax.org The Jackson Laboratory is an Affirmative Action/Equal Opportunity Employer.
Lesley S. Bechtold Supervisor, Biological Imaging The Jackson Laboratory 600 Main St. Bar Harbor, ME 04609 207-288-6191
Dear readers I need some advice in order to optimize he carbon coating process of my replicas for TEM. In particular I need some tips about: -geometry of carbon rods, sharpening, distance between the tips, etc; -current density; -use of carbon ropes: how do they work and how has to be changed the set up I would be really grateful if You can help me. Regards
Alessandro Mattozzi Dept. of Fibre and Polymer Technology
Please note that the Core Facility Management session at M&M 2003 has been rescheduled. It originally was scheduled for Thursday August 7 from 1:00pm to 3:00pm. NEW TIME is Thursday August 7 from 8:00am to 10:00am in room 202B. Please change the time in your meeting schedules.
Topics for this year are:
How to Increase the Use of the Core Facility...with a Corresponding Increase in Revenue. Facilitator: Elaine Humphrey, U. of British Columbia
Defining the Roles of the Lab manager/Director and the Advisory Committee of a major Core Facility. Facilitator: Debby Sherman, Purdue University.
The goal of this session to to promote discussion of topics of interest to facility managers/directors. Therefore attendees are encouraged to bring information relative to the topics that they would like to share. In this case it may include examples of ways to advertise facility services (brochures, web sites, etc). Or perhaps you have a mission statement that helps clarify roles of staff and advisory committee members. Please contact me if you would like to share information and what sort of audio-visual equipment you would like to have available. Although it helps to have advanced notice to better organize the session, last minute inclusions are always welcome.
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
Dear Colleagues, On behalf of the Organizing Committee, we have the pleasure to inform you that the 4th ASEAN Microscopy Conference and the 3rd Vietnam Conference on Electron Microscopy will be held during 5th - 6th January, 2004 in Hanoi. We would like to ask for your participation with an oral presentation and / or a poster session. The deadline for article submission will be on 30, August 2003. The submitted and approved articles will be put in the Proceedings of the Conference For registration and for Hotel reservation, please use the second announcement and call for paper that you have already received. Looking forward for your answer and we will be happy to welcome you in Hanoi.
Chairman of the Organizing Committee Prof. Nguyen Van Man M.D., D.M.Sc
If you need more informations, please contact at: Assoc. Prof Nguyen Kim Giao Electron Microscopy Unit National Institute of Hygiene and Epidemiology 1- Yersin Str - HaiBaTrung Distr - HaNoi-VietNam Tel: 84.4.9715434 Fax: 84.4.8210853 Email: emlad-at-hn.vnn.vn or emunihe-at-vol.vnn.vn
Omega Optical, the worldwide leading supplier of optical filters for fluorescence microscopy, is seeking to fill a position to sell our products. Interested applicants may apply by sending a resume to:
rgorham-at-omegafilters.com
Ruth Gorham Houle V.P. of Business Development Omega Optical, Inc. 210 Main St., PO Box 573 Brattleboro, Vermont 05301 ph:802-254-2690 X-127 t.f.ph:866-488-1064 fax:802-254-3937
Most EM's I know use PCI Quartz. The output of your EM will not be a standard frame rate of 30 frames/sec. [60 fields per sec.] but something less, so you simply can't put it into a video "frame grabber" card that only accepts NTSC [RS-170] video.
Oops, I just noticed you are in the UK! Video frame rates are different there, more scan lines per field but less fields/sec. Nonetheless, PCI Quartz sould do the job.
I have no interest in PCI Quartz other than as a user.
Hope this is helpful.
Peter Tomic Agere Systems Allentown, PA USA
-----Original Message----- } From: Kevin Mackenzie [mailto:k.s.mackenzie-at-abdn.ac.uk] Sent: Wednesday, July 16, 2003 9:00 AM To: Microscopy-at-sparc5.microscopy.com
I have the possibility of obtaining an Cambridge S90 that includes the factory installed Digital Image Store, that will allow Frame Averaged, Frame Integrated or Line Integrated video at tv rate.
Does anyone have any experience of connecting the tv output of the SEM to a PC TV card to allow the image to be saved on the computer?
Or do I need to spend more money in buying a digital imaging system.
I welcome any suggestions or recommendations
many thanks
Kevin
Electron Microscope unit School of Biological Sciences University of Aberdeen Aberdeen AB24 2TZ
Tel 01224-272847 Fax 01224-272396 ------------ Kevin Mackenzie k.s.mackenzie-at-abdn.ac.uk
The accepted method of cleaning oil from immersion lenses is with lens tissue paper wetted with xylene. The lens tissue paper should be available from any camera store.
I am trying to explain and compare the similarities and differences of ESEM equipment and VPSEM equipment in plain language to our Administrators. I know that the 2 types of equipment share some characteristics but others are unique. I also understand that the 2 types of equipment are unique in that individual companies produce one or the other, but not both.
I know that there have been some discussions about this which are archived, but with the pace that technology changes, I wanted to ask these questions again to be sure that I could provide the most up to date information possible to my Administrators.
Could you please help?
Thank you in advance.
Kind regards,
Paula.
Paula M. Allan-Wojtas Research Scientist - Food Microstructure / Chercheur scientique - microstructure des aliments Food Safety and Quality Team / Salubrité et qualité des aliments Agriculture and Agri-Food Canada /Agriculture et Agroalimentaire Canada Telephone / Téléphone: 902-679-5566 Facsimile / Télécopieur: 902-679-2311 32 Main Street / 32 rue Main Kentville, Nova Scotia / Kentville (Nouvelle-Écosse) B4N 1J5 allanwojtasp-at-agr.gc.ca
The Spouses Program offered as a separate social event at the M&M 2003 meeting in San Antonio has been canceled. The number of people registering for the event fell far short of the 35 person minimum we needed. Refunds can be arranged at the meeting in the registration area by contacting Monica Eggers at the registration desk, or Ev Osten or Dwight Erickson at the Local Arrangements Committee booth.
At the Local Arrangements Committee booth we can give you information on some of the venues that were part of the program and some of the things you can do on your own.
Ev Osten
Local Arrangements Committee Chair Microscopy & Microanalysis 2003 August 3 - 7, San Antonio
efosten-at-mmm.com 651-736-0104 fax: 651-733-0648
3M Company Corporate Analytical Technology Center 3M Center, 201-BE-16 St. Paul, MN 55144-1000 USA
Good News. Almost all of the lost master tapes from The MSA video collection have been recovered. So, some of the titles that have not been available for a while can now be provided in VHS or DVD format. The current catalog is available as a link on the MSA web page. Mastercard and VISA are now accepted. We also accept institutional purchase orders, check, money order, cash and precious gems that are appropriately mounted. As always the MSA Education Committee welcomes new contributions to the video collection and encourages all members to consider presenting tutorials at our annual meeting on new topics or on topics previously covered, that are in need of updating.
Greg Erdos MSA Video Wrangler
Gregory W. Erdos Ph.D. Assistant Director, Biotechnology Program P.O. Box 118525 217 Carr Hall University of Florida Gainesville, FL 32611 gwe-at-ufl.edu 352-392-1295
Dear Alessandro, I also use a JEOL Evaporator. I use 5mm diameter carbon or graphite rods, one just flattened on the end and the other sharpened to a 1mm diameter tip, about 4mm long. These rods just touch, the sharpened point in the centre of the flat one. I stick the rods out of the metal holders about 10 mm each, with the springs on one about one-half stretched, so that the two stay in contact as the sharpened tip burns down. I pump to at least a 5 X 10-3 torr vacuum, then turn the heater current up to 35A, not too quickly. Leave it there and the current will increase by itself. I go until it reaches 45 to 50 A, but do not leave the heat on for more than 30 seconds. If the vacuum degrades above 10-4 torr, turn the current off, wait for the vacuum to improve, then turn up the current again. The best way to decide on how much to coat is by coating a polished brass disc at the same time, beside your samples. The brass goes from yellow to gold to orange to red and then has a sudden transition from red to blue at 25 nm thickness. For TEM replicas I would use about the orange or red thickness, but experience will help you to know what works best. Good luck, 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: "by way of Ask-A-Microscopist" {alessandro-at-polymer.kth.se} To: {Microscopy-at-sparc5.microscopy.com} Sent: Wednesday, July 16, 2003 5:37 AM
Dear Alessandro Mattozzi,
We can share our optimized conditions for making carbon.
The carbon rod which we used is about 1/4" in diameter. We sharp one end (about 1/4" length) of rod to 1/8" in diameter.
According to our experiences, the vacuum in the evaporating chamber is the most important parameter for making the high quality carbon films. The higher is better. Normally, we use 1.5 X 10 -6 torr.
The best current value is really depended on the vocuum which you use. Lower vocuume needs low current to get sufficient speed of growthing carbon film, higher vocuum needs higher current value. However, the carbon film made under low vocuum is easy to be oxidized and the film is very soft and easy to be broken.
One more tip for you is that, the speed for growthing carbon film is related with the current value. However, maxima current for growing a high qulity carbon film is limited by if you could see the spark away from the filament. The spark is large group of carbon moleculars which will mass up your carbon film.
We are a manufactory of making TEM grids for more than 5 years in United States. We provide the high quality TEM grids and the grids pre-coated with carbon films. You are welcome to try our product. For more information about product, please visit our website as below,
http://www.grid-tech.com/
shall you have any questions, I will be happy to answer.
with our best regards
Wendy Zhang ===== Pacific GridTech A high quality EM grid provider 3505 Caminito Carmel Landing San Diego, CA 92130, USA Tel: (858) 336 8938; Fax: (858) 259 5511 Email: info-at-grid-tech.com Web: http://www.Grid-Tech.com/
} } Dear readers } I need some advice in order to optimize he carbon } coating process of my } replicas for TEM. In particular I need some tips } about: } -geometry of carbon rods, sharpening, distance } between the tips, etc; } -current density; } -use of carbon ropes: how do they work and how has } to be changed the set up } I would be really grateful if You can help me. } Regards } } Alessandro Mattozzi } Dept. of Fibre and Polymer Technology } }
===== Pacific GridTech A high quality EM grid provider 3505 Caminito Carmel Landing San Diego, CA 92130, USA Tel: (858) 336 8938; Fax: (858) 259 5511 Email: info-at-grid-tech.com Web: http://www.Grid-Tech.com/
In many cases you can simply connect the output of the SEM to a video card. However, those signals need to follow the various video standards (NTSC, RS-170, PAL, etc.) which limits their resolution (for example: 640x480 for RS-170). Typically the signals are transmitted as composite signals on a coax cable.
If you want to digitize higher resolution modes, you need some special hardware (for an alternative to the Quartz system, see the ADDA II on our web site). These systems are more expensive, as they use special hardware.
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: Kevin Mackenzie [mailto:k.s.mackenzie-at-abdn.ac.uk] Sent: Wednesday, July 16, 2003 7:00 AM To: Microscopy-at-sparc5.microscopy.com
I have the possibility of obtaining an Cambridge S90 that includes the factory installed Digital Image Store, that will allow Frame Averaged, Frame Integrated or Line Integrated video at tv rate.
Does anyone have any experience of connecting the tv output of the SEM to a PC TV card to allow the image to be saved on the computer?
Or do I need to spend more money in buying a digital imaging system.
I welcome any suggestions or recommendations
many thanks
Kevin
Electron Microscope unit School of Biological Sciences University of Aberdeen Aberdeen AB24 2TZ
Tel 01224-272847 Fax 01224-272396 ------------ Kevin Mackenzie k.s.mackenzie-at-abdn.ac.uk
Stu- Xylene is pretty strong stuff. Won't it hurt the coatings? I just use 1) a Q-tip with optical lens cleaner and then lens tissue paper with lens cleaner. Is that okay? Rgds, Mike Shaw Roselle, NJ
} The accepted method of cleaning oil from immersion } lenses is with lens tissue paper wetted with xylene. } The lens tissue paper should be available from any } camera store. } } Stu Smalinskas } SKF USA } Plymouth, Michigan
Dear Wendy Zhang Yes, you right: better vacuum, better carbon film quality. But, using Electron Gun evaporation in combination with good vacuum delivered even better results. Thermal evaporation is not effective to produce mono atomic vapors. It's more like clouds of 10-15+ atom's clusters. Electron beam evaporation utilized bombardment of carbon by accelerated electrons. This way is more effective in formation nearly mono atomic carbon "clouds" and therefore better carbon film homogeneity/uniformity. This process is also may be controlled better than thermal evaporation. Finally, you may produce very thin, uniform and stable films. Personally, I am using 1.2-1.5 nm thick films for routine work in shadowing and negative staining. Best wishes, Sergey.
At 02:24 PM 7/17/2003, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
Things are changing rapidly. Five years ago only the ESEM could offer SE imaging and water on/in the specimen. Now many of the rivals can offer SE imaging in VPSEM. At a recent meeting one competitor claimed, in converstion, to be able to image water. I have not got round to asking for some images from them. I would check out the suppliers for current performance information.
Dave
On Thu, 17 Jul 2003 15:05:48 -0400 Paula Allan-Wojtas {AllanWojtasP-at-agr.gc.ca} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Hi, all, } } I am trying to explain and compare the similarities and differences of } ESEM equipment and VPSEM equipment in plain language to our } Administrators. I know that the 2 types of equipment share some } characteristics but others are unique. I also understand that the 2 } types of equipment are unique in that individual companies produce one } or the other, but not both. } } I know that there have been some discussions about this which are } archived, but with the pace that technology changes, I wanted to ask } these questions again to be sure that I could provide the most up to } date information possible to my Administrators. } } Could you please help? } } Thank you in advance. } } Kind regards, } } Paula. } } } } Paula M. Allan-Wojtas } Research Scientist - Food Microstructure / Chercheur scientique - } microstructure des aliments } Food Safety and Quality Team / Salubrité et qualité des aliments } Agriculture and Agri-Food Canada /Agriculture et Agroalimentaire } Canada } Telephone / Téléphone: 902-679-5566 } Facsimile / Télécopieur: 902-679-2311 } 32 Main Street / 32 rue Main } Kentville, Nova Scotia / Kentville (Nouvelle-Écosse) } B4N 1J5 } allanwojtasp-at-agr.gc.ca } } } } } } } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
We'll soon be getting a Nikon SMZ 1000 stereoscopic zoom microscope with a beam splitter for a digital camera. This stereomicroscope will be used to help with the making and cleaning of TEM specimens. We'd like to attach a digital camera to catalog the specimen, but really don't need a high end digital camera. Just something to take a quick image of the sample to place in the notebook. So my questions are:
Does anyone know where I can find an adapter to attach a consumer grade digital camera (one bought at Target or Best Buy) to a Nikon SMZ 1000 stereoscopic zoom microscope?
Any recommendations on which consumer grade camera to get?
Thanks for your help everyone!
William Stratton
------------------- William G. Stratton Research Assistant University of Wisconsin - Madison
1509 University Avenue Madison, WI 53706 Office: 608-265-6391 Fax: 608-262-8353 wgstratton-at-wisc.edu
I enclose details of our latest microscopy report.
The report will include market segment size, growth rates, ten year projections, major players and competitive strategies for the microscopy market segment. The report will include an analysis of technology platforms, descriptions of companies in the field, and trends in technology and business impacting this market segment.
It will discuss technology platforms in various forms of microscopy including light, confocal, electron, and scanning probe. The report will examine products in development, perceived medical and market needs, the market outlook, economic considerations, and pricing for the microscopy market segment.
Each corporate profile includes the following elements:
History
Unique Company Strengths Corporate Research and Development Strategy Intellectual Property Assessment Corporate Collaborations Products and Developmental Pipeline Discovery-Stage Projects Summary Assessment of Market Prospect Direct Pipeline Competition Potential Improvements Over Existing Products Financial Details Summary
The information in this report is based upon interviews with sales and marketing professionals of companies in the biotechnology laboratory instrument market. Professionals at laboratories and dealers around the U.S. were queried, some several times, about their institution's products and marketing strategies as well as their overall thoughts about their industry segment.
Other sources of information for the report were trade association publications and meetings, product brochures and catalogs, and company literature. An examination was made of a number of company internet sites, and telephone interviews were conducted with a number of industry marketing executives. Where possible, an examination of the annual reports, 10k filings, and financial reports were used as the basis of the data reported.
Some of the information obtained for the report was taken from Biotechnology Associates databases and from the private data stores of the author. The information set forth in this study was obtained from sources that we believe to be reliable, but we do not guarantee the accuracy, adequacy or completeness of any information or the results obtained by the use of such information.
For a complete index of this report click on http://www.researchandmarkets.com/reports/5531
Report Index:
1. Executive Summary
2. Introduction
3. Light Microscopy 3.1 Products and Technologies 3.2 Market Analysis
4. Confocal Microscopy 4.1 Technologies and Products 4.2 Market Analysis
5. Electron Microscopy 5.1 Technologies and Products 5.2 Market Analysis
7. Semiconductor Processing Systems 7.1 Technologies and Products 7.2 Market Analysis
8. Automated Imaging Systems 8.1 Technologies and Products 8.2 Market Analysis
9. Summary and Conclusions
10. Company Profiles 10.1 Thermo Spectra Corporation 10.2 Applied Imaging Corporation 10.3 Bio-Rad Laboratories, Incorporated 10.4 Digital Instruments 10.5 Applied Precision 10.6 International Remote Imaging Systems (IRIS) 10.7 KLA-Tencor Instruments Corporation 10.8 Leica Incorporated 10.9 Autocyte/Neopath/Cytyc/Neuromedical Systems, Inc. 10.10 Nikon, Inc. 10.11 Olympus Optical Company Limited, Tokyo, Japan 10.12 Carl Zeiss, Inc.
11. Company Directory
List of Tables
Table 1 U.S. Sales of Light Microscopes 1996-2005 (in Dollars) Table 2 U.S. Sales of Light Microscopes 1996-2005 (in Units) Table 3 Breakdown of U.S. Sales of Light Microscopes by Price Range 1999 (in Dollars) Table 4 Breakdown of U.S. Sales of Light Microscopes by Price Range 1999 (in Units) Table 5 Breakdown of U.S. Sales of Light Microscopes by User Segment 1999 (in Dollars) Table 6 Breakdown of U.S. Sales of Light Microscopes by User Segment 1999 (in Units) Table 7 Breakdown of U.S. Sales of Light Microscopes by Manufacturer 1999 (in Dollars) Table 8 U.S. Sales of Confocal Microscopes 1996-2005 (in Dollars) Table 9 U.S. Sales of Confocal Microscopes 1996-2005 (in Units) Table 10 Breakdown of U.S. Sales of Confocal Microscopes by User Segment 1999 (in Dollars) Table 11 Breakdown of U.S. Sales of Confocal Microscopes by User Segment 1999 (in Units) Table 12 Breakdown of U.S. Sales of Confocal Microscopes by Manufacturer 1999 (in Dollars) Table 13 US. Sales of Electron Microscopes 1996-2005 (in Dollars) Table 14 U.S. Sales of Electron Microscopes 1996-2005 (in Units) Table 15 Breakdown of U.S. Sales of Electron Microscopes by Technology 1999 (in Dollars) Table 16 Breakdown of U.S. Sales of Electron Microscopes by Technology 1999 (in Units) Table 17 Breakdown of U.S. Sales of Electron Microscopes by User Segment 1999 (in Dollars) Table 18 Breakdown of U.S. Sales of Electron Microscopes by User Segment 1999 (in Units) Table 19 Breakdown of U.S. Sales of Electron Microscopes by Manufacturer 1999 (in Dollars) Table 20 U.S. Sales of Scanning Probe Microscopes 1996-2005 (in Dollars) Table 21 U.S. Sales of Scanning Probe Microscopes 1996-2005 (in Units) Table 22 U.S. Sales of Scanning Probe Microscopes by Type 1999 (in Dollars) Table 23 U.S. Sales of Scanning Probe Microscopes by Type 1999 (in Units) Table 24 U.S. Sales of Scanning Probe Microscopes by Application 1999 (in Dollars) Table 25 U.S. Sales of Scanning Probe Microscopes by Application 1999 (in Units) Table 26 U.S. Sales of Scanning Probe Microscopes by Manufacturer 1999 (in Dollars) Table 27 U.S. Sales of Semiconductor Processing Equipment 1996-2005 (in Dollars) Table 28 U.S. Sales of Semiconductor Processing Equipment 1996-2005 (in Units) Table 29 Breakdown of U.S. Sales of Semiconductor Processing Equipment by Manufacturer 1999 (in Dollars) Table 30 U.S. Sales of Automated Imaging Systems 1996-2005 (in Dollars) Table 31 U.S. Sales of Automated Imaging Systems 1996-2005 (in Units) Table 32 Breakdown of U.S. Sales of Automated Imaging Systems by Application 1999 (in Dollars) Table 33 Breakdown of U.S. Sales of Automated Imaging Systems by Application 1999 (in Units) Table 34 Breakdown of U.S. Sales of Automated Imaging Systems by Manufacturer 1999 (in Dollars) Table 35 U.S. Sales of Light, Confocal, Electron, And Scanning Probe Microscopes 1996-2005 (in Dollars) Table 36 U.S. Sales of Confocal, Electron And Scanning Probe Microscopes 1996-2005 (in Dollars) Table 37 U.S. Sales of Confocal, Electron And Scanning Probe Microscopes 1996-2005 (in Units) Table 38 1996 and 2005 Sales By Category Of Light, Confocal Electron, and Scanning Probe Microscopes (in Dollars) Table 39 1996 and 2005 Sales By Category of Confocal, Electron, And Scanning Probe Microscopes (in Dollars) Table 40 1996 and 2005 Sales By Category of Confocal, Electron And Scanning Probe Microscopes (in Units) Table 41 Breakdown of U.S. Sales of Light, Confocal, Electron, and Scanning Probe Microscopes by Manufacturer 1999 (in Dollars)
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The method we use is to use a solution of Windex (the original blue one) and water 1:1 and lens tissue. Moisten the lens tissue, DO NOT CRUMPLE it up, and gently sweep it across the front element of the lens. Repeat as needed. Never apply pressure to the lens. You should check with the manufacturer of your lens to see what they recommend, since different companies may use different coatings on their lenses and different coatings may need different handling (a lot of differences!). A build-up of old, dried oil may need more aggressive cleaning, but again, check with the manufacturer or distributor of your microscope. Lee -- Lee Cohen-Gould Electron & Optical Microscopy Facilities Weill Medical College of Cornell U. (212)746-6146 Rms A-105, LC-207
as far as 5302 is concerned, good luck. according to kodak canada it is no longer produced. i have not talked to kodak in new york, but your only available source may be supply houses which have stockpiled it.
Ev; I can certainly understand your disappointment at the lack of registration for the spouses program, but it doesn't appear that you have given much consideration to the people who purchased non-refundable airplane tickets to come to San Antonio, expecting to participate in the spouses program for which they registered.
John Mardinly Intel
-----Original Message----- } From: "efosten-at-mmm.com"-at-sparc5.microscopy.com [mailto:"efosten-at-mmm.com"-at-sparc5.microscopy.com] Sent: Thursday, July 17, 2003 1:06 PM To: Microscopy-at-sparc5.microscopy.com
To M&M 2003 attendees,
The Spouses Program offered as a separate social event at the M&M 2003 meeting in San Antonio has been canceled. The number of people registering for the event fell far short of the 35 person minimum we needed. Refunds can be arranged at the meeting in the registration area by contacting Monica Eggers at the registration desk, or Ev Osten or Dwight Erickson at the Local Arrangements Committee booth.
At the Local Arrangements Committee booth we can give you information on some of the venues that were part of the program and some of the things you can do on your own.
Ev Osten
Local Arrangements Committee Chair Microscopy & Microanalysis 2003 August 3 - 7, San Antonio
efosten-at-mmm.com 651-736-0104 fax: 651-733-0648
3M Company Corporate Analytical Technology Center 3M Center, 201-BE-16 St. Paul, MN 55144-1000 USA
Dear Paula, ESEM is a brand name of the FEI company and their term "Environmental SEM" refers to higher pressures. VPSEM is a generic term that includes ESEM and the other SEMs that use this form of imaging. Each manufacturer will give the range of pressures that they will operate at in their specifications. Good luck, 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: "Paula Allan-Wojtas" {AllanWojtasP-at-agr.gc.ca} To: {microscopy-at-sparc5.microscopy.com} Sent: Thursday, July 17, 2003 12:05 PM
Hi, all,
I am trying to explain and compare the similarities and differences of ESEM equipment and VPSEM equipment in plain language to our Administrators. I know that the 2 types of equipment share some characteristics but others are unique. I also understand that the 2 types of equipment are unique in that individual companies produce one or the other, but not both.
I know that there have been some discussions about this which are archived, but with the pace that technology changes, I wanted to ask these questions again to be sure that I could provide the most up to date information possible to my Administrators.
Could you please help?
Thank you in advance.
Kind regards,
Paula.
Paula M. Allan-Wojtas Research Scientist - Food Microstructure / Chercheur scientique - microstructure des aliments Food Safety and Quality Team / Salubrité et qualité des aliments Agriculture and Agri-Food Canada /Agriculture et Agroalimentaire Canada Telephone / Téléphone: 902-679-5566 Facsimile / Télécopieur: 902-679-2311 32 Main Street / 32 rue Main Kentville, Nova Scotia / Kentville (Nouvelle-Écosse) B4N 1J5 allanwojtasp-at-agr.gc.ca
I've worked in many labs where xylene was set up as the solvent to clean immersion lenses, so I figure this solvent is most widely accepted. I know it smells strong, but I don't consider it a powerful solvent. I'm open to other ideas. Perhaps we should look at this the other way and ask which chemicals we should avoid. Harm may come to the optical coating and glue used to set the lenses in place if the wrong solvent is used. I believe we can cross off acetone from the accepted solvent list.
Stu- Xylene is pretty strong stuff. Won't it hurt the coatings? I just use 1) a Q-tip with optical lens cleaner and then lens tissue paper with lens cleaner.
Is that okay? Rgds, Mike Shaw Roselle, NJ
} The accepted method of cleaning oil from immersion } lenses is with lens tissue paper wetted with xylene.
} The lens tissue paper should be available from any } camera store. } } Stu Smalinskas } SKF USA } Plymouth, Michigan
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To all, The literature that accompanies each box of 4489 film, including the New Formulation still indicates processing "with nitrogen-burst agitation (1-second burst every 8 seconds)." Has there been an (un)official statement from Kodak suggesting otherwise?
Paul
Paul J. Gerroir Microscopy Materials Characterization Xerox Research Centre of Canada 2660 Speakman Drive Mississauga, Ontario L5K 2L1
Thanks, Stu- Will not use Acetone. Am also checking with manufacturer of my lenses for additoinal info. Rgds, Mike Shaw
In a message dated 7/18/2003 1:31:29 PM Eastern Standard Time, smalinskas-at-yahoo.com writes:
} Perhaps we should } look at this the other way and ask which chemicals we } should avoid. Harm may come to the optical coating } and glue used to set the lenses in place if the wrong } solvent is used. I believe we can cross off acetone } from the accepted solvent list.
William- I have a Canon A40, and I hook it up to a TV monitor for viewing and focusing. The nice thing about it is that you can buy a ready-made Canon bayonette adapter-to-52mm right from Canon at camera store. This pops onto the front of the camera, and it is a simple matter to hook up to a T-mount or any other ring from 52mm to whatever you need. You then have easy-on and easy-off of the camera from the microscope without having to unscrew the camera and remove cords... That's why I chose the Canon A-40. If you want more info- I have pictures as well as the set up on my website, so let me know if you are intersted. Rgds, Mike Shaw Roselle, NJ
In a message dated 7/18/2003 8:13:29 AM Eastern Standard Time, wgstratton-at-wisc.edu writes:
} Does anyone know where I can find an adapter to attach a consumer grade } digital camera (one bought at Target or Best Buy) to a } Nikon SMZ 1000 } stereoscopic zoom microscope? } } Any recommendations on which consumer grade camera to get? } } } Thanks for your help everyone! } } William Stratton
The Nikon Coolpix cameras have threaded lenses for various adapters that can be used to attach these consumer grade cameras to your microscope. Nikon and some aftermarket suppliers provide an adapter from the threads on the lens to a C-mount that should be on your micoscope. The adapter will also fit right into some eyepiece tubes for use on microscopes without a C-mount.
We have used the Coolpix 990 and 4500 models in this way. Obviously, the images are not of the same quality as the dedicated digital acquisition systems, but are pretty good quality for the cost. The cameras use a Compact Flash card for image storage, which is not as convenient as direct saves to disk but is relatively convenient with an inexpensive USB card reader. I am not so found of the direct USB attachment to the camera and Nikon softward for retrieving images.
Your Nikon rep should be able to provide information on the adapter, or do a quick search on google for "coolpix microscope adapter" to find information on Nikon and aftermarker alternatives.
-- Larry D. Hanke, P.E. Materials Evaluation and Engineering, Inc. Practical Solutions Through Technology and Innovation http://www.mee-inc.com (763) 449-8870
A company rep reads this mailing list and he will, I hope let you know that he has what is needed to connect the Coolpix to microscopes. We bought their generic adapter and the pieces needed to attach it to an inverted, a metallurgical and to a sophisticated trinocular. As it turns out, with just the generic, we can put it on our dissecting scope, and into the port on our hardness tester. We even used it on a trinocular scope in another lab. It is quite versatile.
Ron L
-----Original Message----- } From: Larry Hanke [mailto:hanke-at-mee-inc.com] Sent: Sunday, July 20, 2003 5:12 PM To: William Stratton Cc: Microscopy-at-sparc5.microscopy.com
The Nikon Coolpix cameras have threaded lenses for various adapters that
can be used to attach these consumer grade cameras to your microscope. Nikon and some aftermarket suppliers provide an adapter from the threads
on the lens to a C-mount that should be on your micoscope. The adapter will also fit right into some eyepiece tubes for use on microscopes without a C-mount.
We have used the Coolpix 990 and 4500 models in this way. Obviously, the
images are not of the same quality as the dedicated digital acquisition systems, but are pretty good quality for the cost. The cameras use a Compact Flash card for image storage, which is not as convenient as direct saves to disk but is relatively convenient with an inexpensive USB card reader. I am not so found of the direct USB attachment to the camera and Nikon softward for retrieving images.
Your Nikon rep should be able to provide information on the adapter, or do a quick search on google for "coolpix microscope adapter" to find information on Nikon and aftermarker alternatives.
-- Larry D. Hanke, P.E. Materials Evaluation and Engineering, Inc. Practical Solutions Through Technology and Innovation http://www.mee-inc.com (763) 449-8870
I'm having some trouble aligning the imaging system of an Hitachi H-7000. When it comes to changing the spot sizes from 1-7, and then moving it back to position, I find that even if I move just from 1 spot size to the next, the spot moves right off the screen, and it takes a very long period of time to find it. If I tried to directly move to spot 7 from say spot 5, when I finally do find spot 7 with condensor shift controls, I find that the beam is not perfectly round anymore, but half of the beam is obscured somehow, even though all of the apertures are out, and the specimen rod is out of the beam.
I tried for a long time working on this problem with no success. It seemed to take forever to find the beam after just changing from one spot size to the adjacent spot size. IT would just FLY off the screen, and it didnt' help to try "reset" button of the microscope or even just resetting the condensor shift controls.
I haven't used this H-7000 for quite some time, and now I'm only used to the JEOL, where we have a "bright tilt" button to turn on first. Is there a corresponding "bright tilt" button somewhere on the H-7000 that I have forgotten about?
On most TEMs (I don't know specifically about the H-7000) this alignment is done with the gun translation control. Most TEMs have 2 sets of deflectors between the gun and the sample. You are controlling only one set with the beam shifts. This set of deflectors is between the C2 lens and the sample. The other set of deflectors is between the anode and the C1 lens. These are the gun tilt and gun translate coils.
The gun translate is usually set by varying the C1 lens between a strong and weak setting. Think about the case where the beam (coming out of the gun) is entering the C1 lens off-axis.
Consider a strong C1 - the focal length is short and the cross-over will be close to the optic axis. So as a 1st approximation we consider it to be on the axis and center the beam with the normal beam translation knobs.
Now weaken the C1 lens - the focal length gets longer and the cross-over is proportionately farther away from the axis. Now use the gun translate (or shift) control to align the beam entering the C1 lens.
Repeat the process until the shift is minimal. Check your owners book to see how Hitachi recommends doing this alignment.
Cheers, Henk
At 09:30 AM 7/21/2003 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hendrik O. Colijn colijn.1-at-osu.edu Campus Electron Optics Facility Ohio State University (614) 292-0674 http://www.ceof.ohio-state.edu Time is that quality of nature which keeps events from happening all at once. Lately it doesn't seem to be working.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Xiao.ming.wang-at-mail.mcgill.ca) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, July 21, 2003 at 08:55:22 ---------------------------------------------------------------------------
Email: Xiao.ming.wang-at-mail.mcgill.ca Name: Xiaoming Wang
Organization: McGill Univesity
Education: Graduate College
Location: Montreal Canada
Question: Dear Sir/Madam,
I am looking for a software on trace analysis on TEM. Do you know any body who has the software or who is working on it?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jaideepp-at-rci.rutgers.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, July 21, 2003 at 10:07:26 ---------------------------------------------------------------------------
Question: Are there any softwares available to calculate the edge orientation polarograms and and/or anisotropy index from SEM images of oriented particles
In a message dated 7/21/03 2:09:28 PM, jaideepp-at-rci.rutgers.edu writes:
} Question: Are there any softwares available to calculate the edge orientation } polarograms and and/or anisotropy index from SEM images of oriented particles
That function is included in the Image Processing Tool Kit (www.ReindeerGraphics.com)
As previously announced, the spouses/companions program had to be canceled because we fell far short of the minimum number required by the tour company. As an alternative, I suggest that the people who are still interested in such activities get together at the Local Arrangements Committee booth in the registration area of the Convention Center at 8:30 on Monday morning, August 4. There you can meet the other people, and we (the LAC) can provide you with suggestions and information on things that an unstructured group can do.
Ev Osten
Local Arrangements Committee Chair Microscopy & Microanalysis 2003 August 3 - 7, San Antonio
efosten-at-mmm.com 651-736-0104 fax: 651-733-0648
3M Company Corporate Analytical Technology Center 3M Center, 201-BE-16 St. Paul, MN 55144-1000
Garry, The following is an alignment procedure we use for our Hitachi H-7000. You might try following it to see if it will help you. This method always works for us, so if you still have problems you might need to call service. Good luck. Mary Gail Engle
ALIGNMENT OF HITACHI H 7000
Center and Saturate Filament Turn acc voltage on- press 75, located above ready/on button Set magnification to 5000X Turn filament knob clockwise until screen is just illuminated. Condense beam with brightness knob to see wehnelt image Center wehnelt image with gun tilt and gun horizontal pods (lower left panel). Saturate filament by turning filament knob until image barely disappears. (Do not oversaturate) Spread beam with brightness knob
Center Condenser Aperture (two methods for this) Take objective aperture (center pod on column) out by moving lever to right. Condenser aperture (top pod on column) should be in #2 position. Expand beam with brightness knob to edge of focusing screen. Center beam with condenser aperture X/Y knobs Reverse brightness knob and repeat above (expand beam and center with X/Y knobs) If beam spot is elliptical correct with condenser stigmator knobs (lower right panel) until beam spot is circular.
OR:
Center Condenser Aperture Take objective aperture (center pod on column) out by moving lever to right. Condenser aperture (top pod on column) should be in #2 position. Press Index, press enter, press 11, press Index Move cursor to C3 lens using arrows Press C3/OBJ Modulation Button (upper right panel) Adjust X/Y knobs of the movable condenser aperture until beam spreads concentrically. Press C3/OBJ; Press Index If beam spot is elliptical correct with condenser stigmator knobs (lower right panel).
Center Beam Condense and center beam (with brightness knob) Set spot size to 2 (lower left panel). Center spot with brightness centering knobs. Set spot size to 7. Center spot with gun horizontal knobs (lower left panel). Repeat until beam is centered and doesn't move off center. Set spot size to 5. Center beam with brightness centering and spread beam Insert objective aperture (move lever to left)
FOR THE FOLLOWING PROCEDURES INSERT HOLEY GRID OR SAMPLE
Center Objective Aperture Make sure beam is on specimen and not a grid bar or empty space Condense beam with brightness knob. Press DIFF button. Set camera length to 0.4 with magnification knob. Adjust bright spot to very small size with diffraction spot knob Center aperture with X/Y knobs on the objective aperture pod (on the column). **DO NOT LEAVE IN DIFFRACTION MODE FOR MORE THAN 30 SECONDS** Press zoom button and spread beam with brightness knob
Align Imaging System Turn magnification to 20,000X and focus image with focus pods Turn on HV modulation (top right panel) Center beam with brightness centering pods. Adjust beam tilt knobs (lower right panel) until imaged moves in and out and not side to side. Turn off HV modulator button
Check Objective Aperture Astigmation Turn magnification up to 80,000. Press objective stigmatior reset switch (lower left panel) Look for a small hole. Under focus (counterclockwise) so white edge appears inside the hole evenly. If the white rim is not even around the hole, then use the objective stigmator X/Y knobs (lower left panel) to make it even around the entire hole
Mary Gail Engle Sr. Research Laboratory Manager Electron Microscopy & Imaging Facility Health Sciences Research Bldg. 001 University of Kentucky Lexington, KY 40536-0305
This is a common problem which arises due to alignment conflicts between the electron gun and the condenser system. The alignment procedures is set out below, it is a generic alignment for any TEM.
1. Large spot size centre the gun alignment shift (use some if the spot and halo moves out of alignment) 2. Small(er) spot size centre the illumination alignment shift. 3 Repeat for a constant centre.
Note 1 - most experienced operators simply align for the positions either side of the spot size they use most. Note 2 - to reduce this misalignment, train ALL operators that if they are looking at a filament image in order to align the gun, they must ONLY use gun tilt and shift, never use illumination shift as this causes the misalignment.
Good luck
Steve Chapman Senior Consultant Protrain Electron Microscopy Training and Consultancy World Wide Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 www.emcourses.com
----- Original Message ----- } From: "Garry Burgess" {GBurgess-at-exchange.hsc.mb.ca} To: {Microscopy-at-sparc5.microscopy.com} Sent: Monday, July 21, 2003 3:30 PM
Dear readers I am using TEM in order to look at my polymer samples. I replicated it with the double stage system. Do I need some kind of support films (Formvar-Carbon) on the grids or the carbon-metal replicas can be observed with a normal copper grid? I would be grateful if You can help me. Regards Alessandro Mattozzi
Many moons ago I was given the task of embedding and sectioning grey hair (human hair). I had a heck of a time getting the epoxy to infiltrate the cuticle, hence the sections tore and the images were less than stellar. The project ended before I could figure out what the best solution to the problem would be. Many moons later the problem has raised its ugly head and a solution must be found. The infiltration times have been expanded, different epoxies have been tried, and several different protocols have been enacted. The samples must not be subject to harsh chemicals because the integrity of the sample must not be compromised.
Any suggestions would help; I am being vague because the samples are not mine and you all know how companies get when it comes to their hair care products, just send some ideas.
thanks,
John Grazul TEM Facility Manager Cornell University Ithaca, NY 607 255 6421
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Thanks for the great responses to my question regarding attaching a digital camera to a Nikon SMZ 1000 stereomicroscope. To make sure this info is placed in the archives, I'm sending out this email with the responses I received. Hopefully this info can help some of you as well.
Thanks again,
William Stratton
------------------- William G. Stratton Research Assistant University of Wisconsin - Madison
1509 University Avenue Madison, WI 53706 Office: 608-265-6391 Fax: 608-262-8353 wgstratton-at-wisc.edu
The Nikon Coolpix cameras have threaded lenses for various adapters that can be used to attach these consumer grade cameras to your microscope. Nikon and some aftermarket suppliers provide an adapter from the threads on the lens to a C-mount that should be on your micoscope. The adapter will also fit right into some eyepiece tubes for use on microscopes without a C-mount.
We have used the Coolpix 990 and 4500 models in this way. Obviously, the images are not of the same quality as the dedicated digital acquisition systems, but are pretty good quality for the cost. The cameras use a Compact Flash card for image storage, which is not as convenient as direct saves to disk but is relatively convenient with an inexpensive USB card reader. I am not so found of the direct USB attachment to the camera and Nikon softward for retrieving images.
Your Nikon rep should be able to provide information on the adapter, or do a quick search on google for "coolpix microscope adapter" to find information on Nikon and aftermarker alternatives.
-- Larry D. Hanke, P.E. Materials Evaluation and Engineering, Inc. Practical Solutions Through Technology and Innovation http://www.mee-inc.com (763) 449-8870
*************************************************
Hello William,
Correct me if I am wrong, but I thought that in many cases the problem is much less one of resolution than it is one of available light and the sensitivity of the camera and the ability to view specimens at low light levels.
The so-called "high end" digital cameras have also dropped way down in price , so that there is not as much difference as previously between the "high end" vs. "consumer" cameras, but with the better camera, you also get better sensitivity.
The Pixera line of cameras is one of those so called "high end" cameras, some of which sell for a low price, which also includes a lot of useful software that does not come with a consumer camera. Now I am not exactly a disinterested thirty party since we now offer the Pixera line of cameras, see URL http://www.2spi.com/catalog/photo/pixera/pixera.html
But for lower available light levels, you might not get what you need with the best of consumer cameras, that is why I make this point to you. Note that I not posted this message through the listserver outof concern that someone might think it "too commercial".
Chuck
PS: Remember that we are striving to be 100% paperless, therefore there are no paper copies kept of this correspondence. Please be sure to always reply by way of "reply" on your software so that the entire string of correspondence can be kept in one place. ============================================
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William- I have a Canon A40, and I hook it up to a TV monitor for viewing and focusing. The nice thing about it is that you can buy a ready-made Canon bayonette adapter-to-52mm right from Canon at camera store. This pops onto the front of the camera, and it is a simple matter to hook up to a T-mount or any other ring from 52mm to whatever you need. You then have easy-on and easy-off of the camera from the microscope without having to unscrew the camera and remove cords... That's why I chose the Canon A-40. If you want more info- I have pictures as well as the set up on my website, so let me know if you are intersted. Rgds, Mike Shaw Roselle, NJ
see ig you can get a some sort of consumer nikon digital camera is my advise. They have less sexpensive modles. They have an incredible macro mode (2CM away). Optem sells and adaptor for about 135 there are many others. good luck.
I've run across some commercially made adapters for consumer cameras, but don't have the info at home. I'll try and dig it out and send it to you on Monday if I can find it (or post to the list).
As far as what grade camera to get, I'd offer the following thoughts:
It sounds like your use on the stereo won't require much, so just about anything over the $100 threshold should work for that.
You may want to consider something a little upscale from that for other uses. If you don't already have one, these things are great for documenting procedures, getting photo's of staff for the web or bulletin board, and just about anything where "a picture is worth a thousand words". If you haven't gotten into the habit already, it can happen quick. I also use mine to take pictures of my white board before cleaning it off. It sure beats manual transcription, and lets me get back to work faster.
For reference, at about 3 Gigapixels you can take a picture (assuming a decent lens system) and print it with a decent inkjet on photo paper and most people would have a hard time telling an 8 x 10 from a film based 8 x 10. (If you know what to look for, you can tell the difference.)
Just food for thought from a non-microscopist.
GO RED!
John W. Raffensperger, Jr. IS Manager Helwig Carbon Products, Inc. Milwaukee, WI
These three pictures were taken with a SONY DSC-P31 held up against a small maginfying glass up against the eyepiece of a stereodissection 'scope from Fisher Scientific. I would recoemmend something better, but, as you can see, even a crappy setup can yield decent pics.
____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
Greetings William. We use a Nikon Coolpix (990 - I think the latest model is 999) with our Olympus stereomicroscope. Coupler options seem to be best for this series, and it takes good pictures for us. Optem International, www.OptemIntl.com, has lots of couplers and can help you choose the right components (coupler and C-mount). You might also check with your Nikon microscope rep; ours (Mager Scientific in Michigan) sells the Coolpix and adapters for customers who don't have 4-5k$ for their fancy digital camera options. I don't think you'll have problems finding what you need, but if you do just let me know and I will put you in touch with a rep at Mager.
Sincerely, Matt
Matthew Stephenson Analytical Associate Impact Analytical/MMI 1910 West Saint Andrews Road Midland, MI 48640 (989) 832-5555 X506 stephenson-at-impactanalytical.com
Alessandro metal-shadowed carbon replicas can be supported on normal copper grids without a support film. Chris
Dr. Chris Jeffree Inveresk Cottage 26, Carberry Road Inveresk Musselburgh Midlothian EH21 8PR Tel: +44 131 665 6062 FAX +44 131 653 6248 Mobile 07710 585 401 ----- Original Message ----- } From: "Alessandro Mattozzi" {alessandro-at-polymer.kth.se} To: {Microscopy-at-sparc5.microscopy.com} Sent: Tuesday, July 22, 2003 3:25 PM
Good Day. On behalf of the Local Arrangements Committee of the 2003 Microscopy and Microanalysis Meeting in San Antonio, Texas, I would like to invite you (again) to the 2003 Golf Outing to be held Sunday, August 3 at the Pecan Valley Golf Club. Pecan Valley is a majestic golf course, located only six miles from downtown San Antonio and the Riverwalk. Pecan Valley was the site of the 50th Anniversary PGA Championship in 1968 when Julius Boros edged Arnold Palmer on the 18th hole and has also hosted three Texas Opens. It has been rated in Golf Digest¹s Top 50 Public Courses and #1 Public Golf Course in the State of Texas for 2002 (http://www.golftexas.com/pecan1.htm Rating: 74.5 - Slope: 136 - Yards: 7,071). The cost will be $70.00 and will include greens fees, cart, transportation to and from the Convention Center, driving range, lunch buffet and awards banquet. The bus will pick up people at three hotels: Marriott at 6:15, Hyatt at6:45, Hilton at 6:50. Leave the Hilton at 7:00 and arrive at the golf course at 7:15. Callaway club rental is available onsite at $35.00 per set.
People who want to play golf but don't go through online registration should get me a check for $70 made out to "Microscopy & Microanalysis" or pay onsite by check the M&M representative.
I would appreciate you contacting me by e-mail if you will be joining us for the golf outing. Please reply by Friday, July 25, 2003 (even if you have already registered) to Mark Sanders (msanders-at-cbs.umn.edu) with the following information:
Name:
Address:
Phone number:
E-mail:
Hotel in San Antonio:
Have you registered and paid: yes or no
Do you need rental clubs:
Do you have special dietary needs?
Do you want to play in a particular foursome?
Another addition to this year¹s outing is the inclusion of various types of sponsorships. We are also looking for additional raffle prizes and / or giveaways for the event. Any company or individual that becomes a sponsor will be promoted in the following ways: * On the hole of their choice (for hole sponsorships) * On the lunch placemats * In each ³goodie² bag that every golfer will receive at the course If you are interested in playing in the outing, please contact me directly (msanders-at-cbs.umn.edu). If you have a foursome, please let me know and include all four names. If you do not have a foursome, that is fine as well we can pair you into a foursome at the course. If you are interested in becoming a sponsor, please contact me directly and I can get further details to you.
I would also like to announce that there would be 2 ³free² slots for students (preferably). These will be in the name of a former member, golfer and a good friend of mine who passed away a few years ago, Joe Polak.
Contact Mark Sanders with any questions (msanders-at-cbs.umn.edu)
Thanks in advance for your consideration in this matter!
At Pecan Valley you can truly ³Play where Champions Have Left Their Footprints.²
My Best Regards,
Mark Sanders
MSA/MAS Golf Tournament at the Pecan Valley Golf Club
Alessandro Mattozzi wrote: ========================================================= I am using TEM in order to look at my polymer samples. I replicated it with the double stage system. Do I need some kind of support films (Formvar-Carbon) on the grids or the carbon-metal replicas can be observed with a normal copper grid? I would be grateful if You can help me. ========================================================== A lot will depend on the nature of the surface itself. Generally speaking, if you are talking about Pt/C replicas, we normally apply, after the Pt/C a carbon "backing" film, evaporated more or less vertically over the sample (without shadowing). That usually is enough to keep surface tension forces from pulling apart the replica film on the floating water surface when the plastic is dissolved. I have assumed you used polyacrylic acid (PAA) as the replicating polymer. A relatively flat surface results in an inherently more robust replica film and a rough (e.g. fracture) surface results in a replica film that is more likely to need some kind of backing film (because cracks tend to form out of any long straight "shadows").
You can of course always use a support film but any support film can take away some of the otherwise nice contrast in the replica film.
Of course, everyone has their own personal variation on this theme, and I have cited the one we normally use in our own laboratory for making replicas of polymer surfaces.
Chuck
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I would like to use an IEEE 1394 or FireWire camera as a replacement for a PAL video camera on a microscope for regular image capture, but also for time-lapse and real-time recordings. The camera is to be used mainly for widefield microscopy (brightfield and fluorescence), but later on Nipkow disc based confocal microscopy is an option.
I am interested in finding a digital (IEEE 1394 or FireWire) color camera in combination with a FireWire interface (PCI) which can deliver frames at least at a rate of 25 fps. The size of the frames should be at least PAL video size (PAL = 760 x 576 pixels) or more if possible.
To observe polymer sample in TEM, you do need the grids pre-coated with carbon film.
The carbon-metal film is stable, but it could create a strong background of metals on your polymer images, that result the image resolution reducing. In contrast, formvar-carbon film or pure carbon film could provide a clean, weak and amorphous background.
On 200 mesh grid, many carbon films on the holes could be broken autmatically by the time going or by the huminity changing, which is because the holes size is too big to support the pure carbon film strongly. Normally, people use the formvar-carbon film on 200 mesh grids. However the formvar-carbon film has its own weakness, such as the formvar film is easy to be burn by electron beam under higher manification and the formvar could have the chemical reaction with the organic matter in your buffer.
The pure carbon film on 300 mesh and 400 mesh grids is much more stable than that on 200 mesh. I suggest you could try the 300 mesh or 400 mesh grids pre-coated with pure carbon film. I hope the hole size didn't block too much interest area under the magnification which you used.
Our company product any kinds of above carbon coated grids. If you are interested, you could get more information from our website,
http://www.grid-tech.com/
Shall you have more question, I will be happy to answer.
with my best regards,
Wendy Zhang ===== Pacific GridTech A high quality EM grid provider 3505 Caminito Carmel Landing San Diego, CA 92130, USA Tel: (858) 336 8938; Fax: (858) 259 5511 Email: info-at-grid-tech.com Web: http://www.Grid-Tech.com/
} } Dear readers } I am using TEM in order to look at my polymer } samples. I replicated it with } the double stage system. Do I need some kind of } support films } (Formvar-Carbon) on the grids or the carbon-metal } replicas can be observed } with a normal copper grid? } I would be grateful if You can help me. } Regards } Alessandro Mattozzi } }
===== Pacific GridTech A high quality EM grid provider 3505 Caminito Carmel Landing San Diego, CA 92130, USA Tel: (858) 336 8938; Fax: (858) 259 5511 Email: info-at-grid-tech.com Web: http://www.Grid-Tech.com/
This sounds like one more case of the proverbial "hair of the dog that bit you."
Seriously, in case you haven't already tried this: embed in Spurr or LR White. Both are very low viscosity and may offer a better infiltration. Alternatively, look at different infiltration solvents (i.e. ethanol vs acetone vs ...).
Good luck.
"The statements and opinions expressed here by Gary M. Brown represent neither those of ExxonMobil Corporation nor its affiliates."
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
john grazul {grazul-at-ccmr.corne To: microscopy-at-sparc5.microscopy.com ll.edu} cc: Subject: embedding grey hair
07/22/03 04:01 PM
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All,
Many moons ago I was given the task of embedding and sectioning grey hair (human hair). I had a heck of a time getting the epoxy to infiltrate the cuticle, hence the sections tore and the images were less than stellar. The
project ended before I could figure out what the best solution to the problem would be. Many moons later the problem has raised its ugly head and
a solution must be found. The infiltration times have been expanded, different epoxies have been tried, and several different protocols have been enacted. The samples must not be subject to harsh chemicals because the integrity of the sample must not be compromised.
Any suggestions would help; I am being vague because the samples are not mine and you all know how companies get when it comes to their hair care products, just send some ideas.
thanks,
John Grazul TEM Facility Manager Cornell University Ithaca, NY 607 255 6421
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (manton-at-biol.uoa.gr) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 23, 2003 at 09:45:09 ---------------------------------------------------------------------------
Email: manton-at-biol.uoa.gr Name: Dr. Marianna Antonelou
Organization: Univ. Athens, Fac. Biology, Dpt. Cell Biology and Biophysics
Education: Graduate College
Location: Athens, Greece
Question: My goal is to do TEM-immunolabelling on human cells in culture using polyclonal and monoclonal primary antibodies, gold-labeled secondary ntibodies and the post-embedding method. What kind of acrylic embedding resin (compatible with agar) and what protocol (fixatives, time, buffers) would you suggest?
Probably not since Hepes is an organic buffer and KMnO4 is a strong oxidizer that would strongly react with it. Phosphate or an inorganic buffer would be better. Anyway, KMnO4 is such a "destroyer" of fine detail (good for membranes but not much else) that buffers are probably not very useful.
JB
} Hello, } Quick question, Is Hepes compatible with potassium permanganate for } yeast fix? } Mike D
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NO! I did this experiment recently and the mix formed a sludge rapidly and then over the course of hours it turned into a clear solution with a black precipitate. I didn't have sodium cacodylate around so after paraformaldehyde/glut fix, i rinsed in buffer, did 3 quick rinses in water and then fixed in KMnO4 in water and got great results with plant seeds. You risk some osmotic effects with the water rinse but I got away with it. if you are worried, stick with cacodylate (my least favorite buffer). good luck.
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} } } Dear Micronetters, } } } } } } Does anyone know what does the average Image Analysis technician makes } in } } } salary or hourly wage? } } } Is there an image analysis salary survey out there somewhere? } } } } } } I know this is a broad question, but there are so few IA technicians out } } } there. } } } } } } I perform DNA ploidy on breast carcinomas. } } } } } } Thanks for any advance info. } } } } } } Donald G. Awbrey, HT(ASCP) QIHC } } } Electron Microscopy / Image Analysis } } } 817-878-5647 } } } donaldawbrey-at-texashealth.org } } }
Oh! Thanks Nester......
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I don't think you'd beed to buffer the KMnO4 to use as a fixative.
The really good prep method for TEM of yeast that I have tried with pretty good results, uses 2% aqueous potassium permanganate (NOT buffered) as a 2nd fixative, after washing out all traces of the primary glutaraldehyde fix buffered with the PIPES buffer (also containing other ingredients).
The reference is: Robin Wright, Transmission Electron Microscopy of Yeast, Microscopoy Research and Technique 51:496-510 (2000).
Hope this helps,
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)625-5754 FAX, ahlst007-at-tc.umn.edu http://www.cbs.umn.edu/ic/
"You can learn a lot by observation - just by lookin'!" - Yogi Bera
} ------------------------------------------------------------------------ } 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, } Quick question, Is Hepes compatible with potassium permanganate for } yeast fix? } Mike D
I am using fresh aqueous 4% solution of KMnO4, 2 hour on ice with good results. For some unclear to me reason, permanganate fixation always better on yeasts than osmium. Sergey
At 08:10 AM 7/23/2003, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
I'm looking for some kind of mesh, or screen-like material that I can put on double sided tape such that small, hand-picked mineral grains can be dropped and nicely organized in the rectangular openings. After mounting, the minerals and grid will be covered in epoxy, and polished for WDS analyis.
I'm looking for 0.5-1mm -sized openings (that's millimeter, not micron). I've had a wire mesh company send me some samples but as the openings increase in size, so does the diameter of the wire. WIth the right sized holes the wire diameter is way too fat, and also the metal mesh doesn't stay flat on the tape, it curls up. I have a piece of no-see-um tent screen, which has an amazing, yet unacceptable weave under the microscope. The best I have so far is screen-door material, but this has 2mm squares. I've tried fabric shops, and the web. Any other ideas would be appreciated. Scott
************************************************ ....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 ************************************************
I just measured the magnification of the Zeiss 25X plan Neofluar N.A. 0.8 multi immersion objective and found that it has an apparent magnification of 26.88X. Independently, one of our technicians measured it and came out with 27X. (I was checking because I expected to measure 400 um and she had measured 370 um.)
Has anybody else noticed the discrepancy between the advertised value and the measured value? Is this one of those "known" things that we just didn't know?
The Zeiss 10, 40 and 100X objectives gave the expected results.
____________________________________________________________________________ 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/
} Many moons ago I was given the task of embedding and sectioning grey } hair (human hair). I had a heck of a time getting the epoxy to } infiltrate the cuticle, hence the sections tore and the images were } less than stellar. The project ended before I could figure out what } the best solution to the problem would be. Many moons later the } problem has raised its ugly head and a solution must be found. The } infiltration times have been expanded, different epoxies have been } tried, and several different protocols have been enacted. The } samples must not be subject to harsh chemicals because the integrity } of the sample must not be compromised. } } Any suggestions would help; I am being vague because the samples are } not mine and you all know how companies get when it comes to their } hair care products, just send some ideas. } } thanks, } } John Grazul } TEM Facility Manager } Cornell University } Ithaca, NY } 607 255 6421
John, I have a similar problem with the cuticle of drosophila pupae and have found that the best that I can do is to face the block and trim with a glass knife. Orient the cuticle perpendicular to the knife edge. The hair should be exposed at the bottom of the block face to avoid chatter in the sections. Most likely the hair will still break out of the section so a supported grid (formvar or collodion) will be needed to photograph the edges of the sample. Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 91904-6018 215-898-7145 psconnel-at-sas.upenn.edu
In a previous E-mail I sent information on the facility manager¹s session at M&M2003. The information is repeated below.
While arranging to get the Core Facility management session included in the annual meeting, it was suggested that we might consider forming a Focused Interest Group (FIG) for facility managers. The idea would be to have a group to act as a unit for inclusion in the annual meetings and to interact in other ways (web, newsletter, etc) to disseminate information of interest to this group and discuss common problems and concerns.
I hope to assemble a group of volunteers to discuss this idea of whether the core facility managers should petition the MSA Board to form a Focused Interest Group (MSA) within MSA. We would have to meet during the meetings, Tuesday or Wednesday with room and time to be arranged, to discuss this possibility and draft a request document. The summary of this discussion on forming a FIG and the draft document will be presented at the beginning of the Core Facility Management session for brief discussion and a vote.
Please contact me ASAP (dsherman-at-purdue.edu) if you would be willing to attend the initial meeting or would like to weigh in with your opinions. I will contact you either before the meeting or via the message board at the meeting with time and place. Feel free to comment on this possibility even though you may not be going to the meeting this year. All E-mails will be presented as part of the discussions. Thanks
Debby
============================== Original E-mail as follows:
Please note that the Core Facility Management session at M&M 2003 has been rescheduled. It originally was scheduled for Thursday August 7 from 1:00pm to 3:00pm. NEW TIME is Thursday August 7 from 8:00am to 10:00am in room 202B. Please change the time in your meeting schedules.
Topics for this year are:
How to Increase the Use of the Core Facility...with a Corresponding Increase in Revenue. Facilitator: Elaine Humphrey, U. of British Columbia
Defining the Roles of the Lab manager/Director and the Advisory Committee of a major Core Facility. Facilitator: Debby Sherman, Purdue University.
The goal of this session to to promote discussion of topics of interest to facility managers/directors. Therefore attendees are encouraged to bring information relative to the topics that they would like to share. In this case it may include examples of ways to advertise facility services (brochures, web sites, etc). Or perhaps you have a mission statement that helps clarify roles of staff and advisory committee members. Please contact me if you would like to share information and what sort of audio-visual equipment you would like to have available. Although it helps to have advanced notice to better organize the session, last minute inclusions are always welcome.
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
The two main types of color cameras used in microscopy I know of are the ones with 3 separate CCDs for each of the three primary colors and the ones with a Bayer filter design. In the 3CCD camera the spatial sampling is the same for each of the three colors, but not in the Bayer filter camera design ? How do you deal with this difference when doing quantitative spatial measurements of colored samples in microscopy ?
What is the relative actual resolution seen with a 3CCD color camera and a camera with Bayer filter reconstruction? How to take into account the reduced sensitivity of the Bayer filter type for each color, compared with the 3CCD camera if any ? When are Bayer filter type cameras a valid alternative for a 3CCD camera in microscopy, besides the price ?
What about pixel shifts in a 3CCD camera due to misalignment of the color filters/splitter and the CCDs ?
Scott- You may find what you are looking for at the following site:
http://www.internetplastic.com/intromesh.html
They sell electroformed (flat) grid material with a wide range of pitches out of Ni, Cu, and Au. It is more expensive than what you have been using though. I don't know what thickness you need, this may be too thin, but they have all sorts of other grid materials as well. Good Luck.
Sincerely, Matthew Ervin, Ph.D. (301)394-0017 phone, (301)394-1559 fax MErvin-at-ARL.Army.mil
M/S: AMSRL-SE-RL US Army Research Laboratory 2800 Powder Mill Road Adelphi, MD 20783-1197
Disclaimer: The opinions and views expressed above are those of the author and do not necessarily represent those of the U.S. Army Research Laboratory or any other government agency
-----Original Message----- } From: S. Kuehner [mailto:kuehner-at-u.washington.edu] Sent: Wednesday, July 23, 2003 4:14 PM To: MSA
to the list-
I'm looking for some kind of mesh, or screen-like material that I can put on double sided tape such that small, hand-picked mineral grains can be dropped and nicely organized in the rectangular openings. After mounting, the minerals and grid will be covered in epoxy, and polished for WDS analyis.
I'm looking for 0.5-1mm -sized openings (that's millimeter, not micron). I've had a wire mesh company send me some samples but as the openings increase in size, so does the diameter of the wire. WIth the right sized holes the wire diameter is way too fat, and also the metal mesh doesn't stay flat on the tape, it curls up. I have a piece of no-see-um tent screen, which has an amazing, yet unacceptable weave under the microscope. The best I have so far is screen-door material, but this has 2mm squares. I've tried fabric shops, and the web. Any other ideas would be appreciated. Scott
************************************************ ....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 ************************************************
It is not unusual for objectives to vary by several percent. That is why,when measurements are critical and/or you are using video systems, we recommend actually calibrating the image using a stage micrometer and the full set of optics used for the experiment.
If you have earlier measurements, all is not lost. You can determine an "adjustment" ratio between the old measurements and your new, calibrated values.
Good hunting!
Barbara Foster Microscopy/Microscopy Education 125 Paridon Street, Suite 102 Springfield, MA 01118 PH: 413-746-6931 FX: 413-746-9311 Web: www.MicroscopyEducation.com
Will you be at M&M in San Antonio? If so, don't forget the Tuesday night seminar on Fluorescence Calibration. Also, join the tradition of over 10,000 microscopists: participate in our survey at any time during the meeting and receive a "sweet thank you". Booth #218 -at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at-
At 04:21 PM 7/23/03 -0400, Michael Cammer wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Contact the apps lab at Buehler (Waukegan, IL). They have a great little vacuum embedder which might help you to get better infiltration.
Good hunting, Barbara Foster Microscopy/Microscopy Education 125 Paridon Street, Suite 102 Springfield, MA 01118 PH: 413-746-6931 FX: 413-746-9311 Web: www.MicroscopyEducation.com
Will you be at M&M in San Antonio? If so, don't forget the Tuesday night seminar on Fluorescence Calibration. Also, join the tradition of over 10,000 microscopists: participate in our survey at any time during the meeting and receive a "sweet thank you". Booth #218 -at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at--at-
At 05:34 PM 7/23/03 -0400, Pat Connelly wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
A product is available from SPI, and probably the other EM supply companies as well, that consists of a substrate (glass, conductive material, plastic, .. ???) upon which is imprinted a matrix of tacky elastomeric dots. Particles can be placed on the tacky dots and held in place for analysis, etc. In your case, you might consider using the tacky dots to hold the isolated particles in place during segregation and subsequent embedment.
I'm sure that my "tacky dot" terminology is wrong but I have actually seen the product at the 2002 M&M exhibition. It looks like it could work for your application.
Good luck,
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
"S. Kuehner" {kuehner-at-u.washing To: MSA {microscopy-at-sparc5.microscopy.com} ton.edu} cc: Subject: mineral mount grids
07/23/03 03:13 PM
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
to the list-
I'm looking for some kind of mesh, or screen-like material that I can put on double sided tape such that small, hand-picked mineral grains can be dropped and nicely organized in the rectangular openings. After mounting, the minerals and grid will be covered in epoxy, and polished for WDS analyis.
I'm looking for 0.5-1mm -sized openings (that's millimeter, not micron). I've had a wire mesh company send me some samples but as the openings increase in size, so does the diameter of the wire. WIth the right sized holes the wire diameter is way too fat, and also the metal mesh doesn't stay flat on the tape, it curls up. I have a piece of no-see-um tent screen, which has an amazing, yet unacceptable weave under the microscope. The best I have so far is screen-door material, but this has 2mm squares. I've tried fabric shops, and the web. Any other ideas would be appreciated. Scott
************************************************ ....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 ************************************************
I'm not aware of any Zeiss specs for objective accuracy. There must certainly be some amount of allowed error. +/- 0% would be a really expensive objective!!
If you measured 26.88X, that would be about 7.5% error. If she measured 27X, that is 8% error. If you measured 400u and she got 370u, then the 30u difference is 7.5% error relative to your reading. So this jives with the mag readings. It seems that your method of checking magnification may not be as perfect as the objective. But if so, why do the other objectives check out OK? Perhaps there is an issue with two different pairs of eyes looking through the same oculars? Were the oculars tweaked for diopter correction?
Try a calibrated stage micrometer and see what you get. For critical measurement work, the micrometer is used for each objective. Then, that difference in mag is cranked into subsequent measurement readings.
gary g.
At 01:21 PM 7/23/2003, you wrote:
} I just measured the magnification of the Zeiss 25X plan Neofluar N.A. 0.8 } multi immersion objective and found that it has an apparent magnification } of 26.88X. Independently, one of our technicians measured it and came out } with 27X. (I was checking because I expected to measure 400 um and she } had measured 370 um.) } } Has anybody else noticed the discrepancy between the advertised value and } the measured value? Is this one of those "known" things that we just } didn't know? } } The Zeiss 10, 40 and 100X objectives gave the expected results. } } ____________________________________________________________________________ } 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/ } }
Good Afternoon, We are starting some eye projects in my laboratory, and I have done very little with eyes except diagnostic biopsies, etc. Is there a good reference or tips on processing rat eyes for ultrastructural studies. Since there is interest in finding specific areas within the eye (therefore the need to know lateral, medial, rostral, and cranial zones of the eye), how usually is the eye cut (minced) for ultrastructural studies? Is perfusion superior to immersion for fixation? Thank you for any help you can give me. Stanley E. Hansen
I am going to buy an achromatic objective 20x n.a. 0.45 (spring loaded) from the Lambda Praha Ltd. and manufactured as DIN standard i.e. for 160 mm. tube length. I wonder if someone knows that kind of objective and he could tell me something about its quality.
If the measurement is done on a digital image acquired by a digital camera and not directly, this could cause the problem. The variation in the measurement could be caused by inaccuracy of spatial sampling. Calibrating a microscope equipped with a digital camera at "low" magnification with a lens with a relative high N.A. is prone to spatial sampling errors.
If the the Zeiss 25X plan Neofluar N.A. 0.8 objective is not matched to your camera, you get an error, caused by misalignment of the sample relative to the CCD-grid of the camera.
There is an excellent article on this issue by Prof. T. Young from the T.U.Delft in the Netherlands, scroll down a bit and have a look at the section on "Sampling Density":
} } I just measured the magnification of the Zeiss 25X plan Neofluar N.A. 0.8 } } multi immersion objective and found that it has an apparent magnification } } of 26.88X. Independently, one of our technicians measured it and came out } } with 27X. (I was checking because I expected to measure 400 um and she } } had measured 370 um.) } } } } Has anybody else noticed the discrepancy between the advertised value and } } the measured value? Is this one of those "known" things that we just } } didn't know? } } } } The Zeiss 10, 40 and 100X objectives gave the expected results. } } } } ____________________________________________________________________________ } } 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/ } } } }
A variety of polymer or metal screens are available from Small Parts, Inc. www.smallparts.com. They sell plain weave polyester with 1mm openings and the open area is 57.5%. With Nylon the openings can be 2 mm with an open area of 53%. If you can tolerate metal then Buckbee-Mears Company, (612) 228-6400, makes metal masks by electropatterning. They have a free sample sheet that contains a flat (no weave) stainless steel screen with 1mm X 0.5 mm openings. They can also make custom parts.
Edison Analytical Laboratories, Inc. is an analytical services laboratory that has no commercial interest in either of the cited sources other than as a user of their products.
Dr. James Carnahan
Edison Analytical Laboratories, Inc. 301 Nott Street Schenectady, NY 12305 (518) 393-2112
In our lab we calibrated every combination of lenses and magnifications we use on our metallograph microscope. We did this with a calibrated stage micrometer and keep this calibration chart nearby for when we need to present accurate measurements.
Stu Smalinskas SKF USA Plymouth, Michigan
At 01:21 PM 7/23/2003, you wrote:
} I just measured the magnification of the Zeiss 25X plan Neofluar N.A. 0.8 } multi immersion objective and found that it has an apparent magnification } of 26.88X. Independently, one of our technicians measured it and came out } with 27X. (I was checking because I expected to measure 400 um and she } had measured 370 um.) } } Has anybody else noticed the discrepancy between the advertised value and } the measured value? Is this one of those "known" things that we just } didn't know? } } The Zeiss 10, 40 and 100X objectives gave the expected results. } } ____________________________________________________________________________ } 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/ }
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Dear Fellow Listers, I am interested in purchasing a Cryo-Ultramicrotome for our user facility; we deal mainly with materials science applications. Would any vendors specializing in this type of product please contact me off-line with information about your products.
Also, if anyone out there has strong opinions, pros or cons, about different systems please contact me. Your input would be greatly appreciated.
Best regards, Ray Twesten
Ray D. Twesten, PhD. Center for Microanalysis of Materials Seitz Materials Research Laboratory 104 S. Goodwin Ave., Urbana, IL 61801 +1 217 244-6177 (Fax -2278)
} From:Dr.ALHAJI USMAN ALIEGO Nigerian National Pet.Corp. Lagos-Nigeria. Attn: President/Ceo.
STRICTLY CONFIDENTIAL BUSINESS PROPOSAL RE: TRANSFER OF US$58.5 MILLION (FIFTY EIGHT MILLION, FIVE HUNDRED THOUSAND US DOLLARS ONLY).
I know this email will reach you as a surprise,but need not to worry as we are using the only secured and confidential medium available to seek for foreign assistance/partnership in a business transaction which is of mutual benefit.
I am a member of the Government of Nigeria Contract Award and Monitoring Committee in the Nigerian National Petroleum Corporation(NNPC)Sometime ago,a contract was awarded to a foreign firm in NNPC by my Committee.This contract was over invoiced to the tune of US$58.5Million .U.S.Dollars. This was done deliberately,the over-invoicing was a deal by my committee to benefit from the project. We now want to transfer this money which is in a suspense Account with NNPC into any Overseas Account which we expect you to provide for us. SHARE: - For providing the account where we shall remit the money into,you will be entitled to 20% of the money,70% will be for me and my partners while 10% has been mapped out from the total sum to cover any upfront or out of pocket expenses that may be incurred by all parties during the course of this transfer,both locally and international expenses.In order for us to commence the transaction i would require the following: -
1.Your company's name,address,telephone and fax numbers.
2.Your bank name,address and account details.(May an account with zero balance)
The above information would be use to make formal applications as a matter of procedure for the release of the money and onward transfer to your account.It does not matter whether or not your company does contract projects of this nature described here. The assumption is that your company won the major contract and subcontracted it out to other companies.More often than not,big trading companies or firms of unrelated fields win major contracts and subcontracts to more specialized firms for execution of such contracts. We have strong reliable connections and contacts at the Central Bank Of Nigeria,as well as the Ministry of Finance and we have no doubt that all the money will be released and transferred if we get the necessary foreign partner to assist us in this deal.Therefore, when the business is successfully concluded we shall through our same connections withdraw all documents used from all the concerned Government Ministeries for 100% security.
We are ordinary Government workers and we will not want to miss this once in a lifetime opportunity to get rich. We want this money to be transferred to the overseas Accounts for us,before the present Government start Auditing all Government owned Parastatals accounts. Please contact me immediately through my email addresses whether or not you are interested in this deal.If you are not,it will enable me scout for another foreign partner to carry out this deal.But where you are interested, send the required documents aforementioned herein through my CONFIDENTIAL email address {usman009-at-freesurf.fr} as time is of the essence in this business. I wait in anticipation of your fullest co-operation.
Yours faithfully,
Dr.ALHAJI USMAN ALIEGO
PLEASE IF YOU ARE NOT INTERESTED ON THIS, KINDLY DESTROY THIS MESSAGE AND MENTAIN THE ADEQUATE SECRET FOR THE SAFETY OF ME AND MY COLEAGUES.
____________________________________________________________ Charle con sus amigos online usando CHAT 123 http://www.123.com/sp/chat/section.php?id_section=329
Scott Kuehner wrote and Gary M. Brown commented on the following request: ========================================================== I'm looking for some kind of mesh, or screen-like material that I can put on double sided tape such that small, hand-picked mineral grains can be dropped and nicely organized in the rectangular openings. After mounting, the minerals and grid will be covered in epoxy, and polished for WDS analysis.
I'm looking for 0.5-1mm -sized openings (that's millimeter, not micron). I've had a wire mesh company send me some samples but as the openings increase in size, so does the diameter of the wire. WIth the right sized holes the wire diameter is way too fat, and also the metal mesh doesn't stay flat on the tape, it curls up. I have a piece of no-see-um tent screen, which has an amazing, yet unacceptable weave under the microscope. The best I have so far is screen-door material, but this has 2mm squares. I've tried fabric shops, and the web. Any other ideas would be appreciated. ============================================================ Gary is right, the SPI Tacky Dot Slides (a proprietary product of SPI Supplies) would probably work, the product being described on URL http://www.2spi.com/catalog/new/tacky.shtml
The standard slide product with the largest dots is the one of 300 µm dots on 2,000 µm centers. So the dots might be a bit too small and the center to center spacing might be too small as well for this particular application... .unless one was to custom fabricate a slide with a larger center to center spacing (not a problem to do).
But the technique will work, and as evidence of that statement, see URL http://www.2spi.com/catalog/new/tackdot_array.html They were prepared exactly as proposed, except with the use of a Tacky Dot Slide instead of a screen mesh.
The cross-sectioned particles are all arranged in an orthogonal array, ready for automated analysis if one has stage automation on their system.
If you still wanted a mesh, you might want to see URL http://www.2spi.com/catalog/standards/stndcal6.shtml
A 30 mesh electroformed screen is available and it has a hole size of 785 µm or 0.785 mm which is in the range of your requirement. The "wire" is 61.5 µm , so I don't know if it will work for you, but that seems to be what you requested.
Charles A. Garber, Ph. D. Ph: 1-610-436-5400 President 1-800-2424-SPI SPI SUPPLIES FAX: 1-610-436-5755 PO BOX 656 e-mail:cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust.Service: spi2spi-at-2spi.com
Look for us! ######################## WWW: http://www.2spi.com ######################## ============================================
I am Dr.Ezenwa Chukwu. Chairman of the Tender Committee of the Nigerian National Petroleum Corporation (NNPC). My Committee is principally concerned with payment of all contract awarded from 1998 to date, in order of priority as regard capital projects of the NNPC. The information we gathered from the Foreign Office of the Nigeria Chambers of Commerce and Industries is so positive as to convince us that you would provide us with solution to a money transfer deal valued at Thirty One Million United States Dollars and subsequently a joint business venture. In the course of our duties as values, and project inspectors for the on-going liquefied Natural Gas (LNG) project, we have over-invoiced the value of some jobs done by foreign contractors for the NNPC to the tune of US$31M. As follows: - Computer optimization and Installation $16,000.000.00 Installation of 250,000.00 Monax Turbine$10,000.000.00 Turn Around Maintenance $5,000,000.00 Our aim of over-invoicing this payment is to divert the excess amount to a discrete account abroad. This fund is now floating in suspense account at the Central Bank of Nigeria (CBN). This is the fund my colleagues and I have decided to transfer into your account since we, as civil servants are not allowed to operate or own foreign account. The money will be shared as follows after transfer:30% for you (Account Owner) 60% for me and my colleagues10% to off-set both local and international expenses that Would be incurred in the course of this transaction. To be able to claim the funds, we will be purporting your company to be the original contractor / beneficiary of the funds so all procedures for international transfer shall be strictly followed, as we have worked out all modalities for a swift and riskfree transfer.
If this proposal satisfies you, please contact me Through ezenwachukwu009-at-go.com with the following important information. Bank Name / Address Account Name ,Account NumberTel/Fax Telex of Bank Personal Phone / Fax Numbers for easy communication. This transaction will last for 14 working days from the time we submit the required information, as all modalities concerning this transaction have been worked out and it is completely risk free.
Please be informed that this subject is classified sensitive. Therefore treat the transaction with utmost confidentiality and urgency.
Yours Faithfully. Dr.Ezenwa Chukwu URGENT REPLY:Please reply to: ezenwachukwu009-at-netzero.com
I am Dr.Ezenwa Chukwu. Chairman of the Tender Committee of the Nigerian National Petroleum Corporation (NNPC). My Committee is principally concerned with payment of all contract awarded from 1998 to date, in order of priority as regard capital projects of the NNPC. The information we gathered from the Foreign Office of the Nigeria Chambers of Commerce and Industries is so positive as to convince us that you would provide us with solution to a money transfer deal valued at Thirty One Million United States Dollars and subsequently a joint business venture. In the course of our duties as values, and project inspectors for the on-going liquefied Natural Gas (LNG) project, we have over-invoiced the value of some jobs done by foreign contractors for the NNPC to the tune of US$31M. As follows: - Computer optimization and Installation $16,000.000.00 Installation of 250,000.00 Monax Turbine$10,000.000.00 Turn Around Maintenance $5,000,000.00 Our aim of over-invoicing this payment is to divert the excess amount to a discrete account abroad. This fund is now floating in suspense account at the Central Bank of Nigeria (CBN). This is the fund my colleagues and I have decided to transfer into your account since we, as civil servants are not allowed to operate or own foreign account. The money will be shared as follows after transfer:30% for you (Account Owner) 60% for me and my colleagues10% to off-set both local and international expenses that Would be incurred in the course of this transaction. To be able to claim the funds, we will be purporting your company to be the original contractor / beneficiary of the funds so all procedures for international transfer shall be strictly followed, as we have worked out all modalities for a swift and riskfree transfer.
If this proposal satisfies you, please contact me Through ezenwachukwu009-at-go.com with the following important information. Bank Name / Address Account Name ,Account NumberTel/Fax Telex of Bank Personal Phone / Fax Numbers for easy communication. This transaction will last for 14 working days from the time we submit the required information, as all modalities concerning this transaction have been worked out and it is completely risk free.
Please be informed that this subject is classified sensitive. Therefore treat the transaction with utmost confidentiality and urgency.
Yours Faithfully. Dr.Ezenwa Chukwu URGENT REPLY:Please reply to: ezenwachukwu009-at-netzero.com
I mostly have to deal with digital microscopy in extreme low-light conditions (fluorescence microscopy) and the quality of the images is very important for the subsequent quantitative analysis.
I would like to know if the following formula for calculating the maximum Signal to Noise Ratio (SNR) for a CCD camera still holds (in general). Photon noise or photon shot noise, refers to the inherent natural variation of the incident photon flux and a such this form of noise limits the maximum SNR for a given CCD-element and the CCD camera ?
If we consider the capacity of one CCD element of a (Silicon-based) CCD, the capacity for photoelectrons "C" relates to the "maximum SNR" of a CCD camera:
SNRmax = 10 log10(C)
In general a CCD camera seems to have a photoelectron capacity per square micron of about 700 photoelectrons/µm^2. So, the maximum SNR of a CCD camera is defined by the "well" capacity of each individual CCD-element (pixel) and cannot be surpassed. So, there is no substitute for square microns per CCD-element to get a higher maximum SNR ? Increasing the SNR in this way however relates inversely to the spatial sampling ?
The total well capacity divided by the noise of the camera system, gives the dynamic range of the camera. The higher the dynamic range, the less noisy the camera becomes. So, there is a direct relation between individual CCD-element surface area and dynamic range ?
There is a lot more to be said on this in digital micrscopy, but I want to know wether this relation is (still) valid and that I did not miss some recent developments. What about the varying quantum efficiency depending on the wavelength of the incident photons ? What about coating CCD-cameras to improve the quantum efficiency in the shorter wavelengths or blocking near-infrared for which CCD's seem to be very sensitive in order to improve the SNR over the entire visible spectrum ?
We are using multiwell plates, mostly according to SBS standards on our automated microscope systems, ranging from 6 wells up to 1536 wells (and beyond for arrays if necessary). I am looking for something which makes life easier to calibrate the XY (and maybe Z) positioning of the sytem.
Are there plates or molds available which contain pinholes or other marks which make it easier to locate the center of the wells for SBS-standard plates (or others formats) ? We can position in XYZ with submicron precision, but XY-positioning in the sub-mm. range should do to start with. This would allow for an automated calibration procedure on the system.
Are there glass plates available in SBS-size format, which can be used for scanning arrays beyond 6144 format density ?
Dear List, We probably have a leak in the body of the Diffstak oil pump in our Philips SEM 515 microscope. Did anybody try to cure such a leak using a leak sealent? Is there any chance to succeed? Another question is on possibility of replacement of the pump (Edwards Diffstak PH 63 58 03, 230 V). Is it still produced or maybe is available as "second hand" item? Thank you for any suggestions,
Leszek Kepinski
Institute of Low Temperature and Structure Research, Polish Academy of Sciences, P.O.Box 1410, 50-950 Wroclaw, Poland kepinski-at-int.pan.wroc.pl
Has anyone been experiencing any section contamination recently that is extremely hard to tract down? Something resembling pepper, but is not due to staining artifacts? Especially in sections with an Epon or EmBed resin component?
??????
Curiously, Randy
Randy Tindall EM Specialist Electron Microscopy Core---We do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
Just a reminder that we are hosting the 8th Annual Materials Microtomy Course in Tucson, Arizona October 28 - 31, 2003.
The course is filling up rapidly so if you wish to participate please drop me an email, or stop by our booth in San Antonio.
Full details, including a printable version of the course brochure, can be found at our website: www.rmcproducts.com
Dave Roberts Director - RMC EM Products Boeckeler Instruments, Inc 4650 South Butterfield Drive Tucson, AZ 85714 Tel: (520) 745-0001 Fax: (520) 745-0004 website: www.rmcproducts.com
A researcher on campus is working with photoreceptor cells, that have been dissociated from the retina. This lab would like to know if anyone on the list serve has a procedure for fixing the cells for light microscopy? They are also interested in a procedure for lectin binding? They would be grateful for any help. Thank you. Nancy Cherim University of New Hampshire
On Monday, July 28, 2003, at 02:21 AM, Peter Van Osta wrote:
} Hi, } } I mostly have to deal with digital microscopy in extreme low-light } conditions (fluorescence microscopy) and the quality of the images is } very important for the subsequent quantitative analysis. } } I would like to know if the following formula for calculating the } maximum Signal to Noise Ratio (SNR) for a CCD camera still holds (in } general). Photon noise or photon shot noise, refers to the inherent } natural variation of the incident photon flux and a such this form of } noise limits the maximum SNR for a given CCD-element and the CCD camera } ? } } If we consider the capacity of one CCD element of a (Silicon-based) } CCD, } the capacity for photoelectrons "C" relates to the "maximum SNR" of a } CCD camera: } } SNRmax = 10 log10(C) } } In general a CCD camera seems to have a photoelectron capacity per } square micron of about 700 photoelectrons/µm^2. So, the maximum SNR of } a } CCD camera is defined by the "well" capacity of each individual } CCD-element (pixel) and cannot be surpassed. So, there is no substitute } for square microns per CCD-element to get a higher maximum SNR ? } Increasing the SNR in this way however relates inversely to the spatial } sampling ? } } The total well capacity divided by the noise of the camera system, } gives } the dynamic range of the camera. The higher the dynamic range, the less } noisy the camera becomes. So, there is a direct relation between } individual CCD-element surface area and dynamic range ? } } There is a lot more to be said on this in digital micrscopy, but I want } to know wether this relation is (still) valid and that I did not miss } some recent developments. What about the varying quantum efficiency } depending on the wavelength of the incident photons ? What about } coating } CCD-cameras to improve the quantum efficiency in the shorter } wavelengths } or blocking near-infrared for which CCD's seem to be very sensitive in } order to improve the SNR over the entire visible spectrum ? } } Best regards, } } Peter Van Osta } Dear Peter, Please post the replies you get to the list. We have a similar problem, since we are doing low-dose EM. The questions asked in your post are the kinds of things we have been considering. We have also considered such problems as readout noise and whether the electronics gain can be optimized to give better S/N. We also need to get high spatial frequencies, so small pixel size and narrow point-spread function are important to us as well. 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
Hello. If you see pepper precipitate just over the tissue it might be osmium. Often when sample is not washed properly after fixation with glut osmium will precipitate. I do not think there is a cure for that when sample if already embedded. Dorota
Still looking for something to do Sunday morning in San Antonio? There is still space available for the 2003 golf outing at Pecan Valley Golf Club! On behalf of the Local Arrangements Committee of the 2003 Microscopy and Microanalysis Meeting in San Antonio, Texas, I would like to invite you (again) to the 2003 Golf Outing to be held Sunday, August 3 at the Pecan Valley Golf Club. Pecan Valley is a majestic golf course, located only six miles from downtown San Antonio and the Riverwalk. Pecan Valley was the site of the 50th Anniversary PGA Championship in 1968 when Julius Boros edged Arnold Palmer on the 18th hole and has also hosted three Texas Opens. It has been rated in Golf Digest¹s Top 50 Public Courses and #1 Public Golf Course in the State of Texas for 2002 (http://www.golftexas.com/pecan1.htm Rating: 74.5 - Slope: 136 - Yards: 7,071). The cost will be $70.00 and will include greens fees, cart, transportation to and from the Convention Center, driving range, lunch buffet and awards banquet. The bus will pick up people at three hotels: Marriott at 6:15, Hyatt at6:45, Hilton at 6:50. Leave the Hilton at 7:00 and arrive at the golf course at 7:15. Callaway club rental is available onsite at $35.00 per set.
People who want to play golf but don't go through online registration should get can pay at the registration booth on Saturday or pay onsite at the course by check to the M&M representative. The cost is $70 made out to "Microscopy & Microanalysis"
I would appreciate you contacting me by e-mail if you will be joining us for the golf outing. Please reply by Wed July 30, 2003 (even if you have already registered) to Mark Sanders (msanders-at-cbs.umn.edu) with the following information:
Name:
Address:
Phone number:
E-mail:
Hotel in San Antonio:
Have you registered and paid: yes or no
Do you need rental clubs:
Do you have special dietary needs?
Do you want to play in a particular foursome?
Shirt size: M L XL XXL
Another addition to this year¹s outing is the inclusion of various types of sponsorships. We are also looking for additional raffle prizes and / or giveaways for the event. Any company or individual that becomes a sponsor will be promoted in the following ways: * On the hole of their choice (for hole sponsorships) * On the lunch placemats * In each ³goodie² bag that every golfer will receive at the course If you are interested in playing in the outing, please contact me directly (msanders-at-cbs.umn.edu). If you have a foursome, please let me know and include all four names. If you do not have a foursome, that is fine as well we can pair you into a foursome at the course. If you are interested in becoming a sponsor, please contact me directly and I can get further details to you.
Contact Mark Sanders with any questions (msanders-at-cbs.umn.edu)
Thanks in advance for your consideration in this matter!
At Pecan Valley you can truly ³Play where Champions Have Left Their Footprints.²
My Best Regards,
Mark "keep your head down" Sanders
MSA/MAS Golf Tournament at the Pecan Valley Golf Club
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (hoganbecky-at-hotmail.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, July 28, 2003 at 20:07:39 ---------------------------------------------------------------------------
Question: I will be teaching gifted 5th graders in a new math/science program this year and need to know what type of microscope would be best for my program. Also, where can I buy a microscope for a reasonable price? Do colleges ever donate used microscopes to schools? I am working with a shoestring budget of $0.00 so I will have to make a personal purchase in order for my students to use this wonderful teaching tool. Thank you for asssisting me with this matter. I want to offer the best science program possible! Becky Hogan
} On Monday, July 28, 2003, at 02:21 AM, Peter Van Osta wrote: } } } Hi, } } } } I mostly have to deal with digital microscopy in extreme low-light } } conditions (fluorescence microscopy) and the quality of the images is } } very important for the subsequent quantitative analysis. } } } } I would like to know if the following formula for calculating the } } maximum Signal to Noise Ratio (SNR) for a CCD camera still holds (in } } general). Photon noise or photon shot noise, refers to the inherent } } natural variation of the incident photon flux and a such this form of } } noise limits the maximum SNR for a given CCD-element and the CCD camera } } ?
There are two excellent articles by J. M. Zuo in Ultramicroscopy,
which detail how to measure the gain, modulation transfer function, and detector quantum efficiency of a CCD imaging system. These articles deal with TEM, but I think the methods should be applicable to light microscopy as well.
For imaging in low signal conditions, I think the DQE is the most relevant parameter. It's defined as
SNR_OUT^2 DQE = ------------------ SNR_IN^2
The electronics that read the CCD chip and convert it to a digital image also add some noise, so you want to characterize the entire system, not just the chip.
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
The performance of a CCD camera depends on a lot of components - the characteristics of the detector on the chip (materials, dimensions, front or rear illumination) as well as the readout electronics (analog) and the subsequent digitization, all of which contribute noise (some additive, some multiplicative). It is a very complicated subject not easily reduced to a simple measurement, unfortunately. The best reference I have seen to the various sources of noise and other characteristics of performance is a very thick but comprehensive book: J. R. Janesick "Scientific Charge-Coupled Devices" SPIE Press, 2001 (isbn 0-8194-3698-4)
First try to determine the exact location of the leak. If it is in the steel/aluminum body it probably could be welded. Are you convinced it is not an O-ring? Torr seal will hold up to 120 degrees C. E-mail our evaporator manufacturing department directly at mb-at-laddresearch.com if you have any further questions and they can probably help.
John Arnott
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: sales-at-laddresearch.com ----- Original Message ----- } From: "Leszek Kêpiñski" {kepinski-at-int.pan.wroc.pl} To: {Microscopy-at-sparc5.microscopy.com} Sent: Monday, July 28, 2003 10:01 AM
Randy,
We can't tell exactly the cause of your problem, but a number of years ago we experienced some problems with raw DDSA. We would think it could be dust particles, but the source could be anywhere. If it is Ladd LX112 Epon or our accelerators or hardeners please let us know and we'll investigate. There have been no other such reports.
John Arnott
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: sales-at-laddresearch.com ----- Original Message ----- } From: "Tindall, Randy D." {TindallR-at-missouri.edu} To: {microscopy-at-sparc5.microscopy.com} Sent: Monday, July 28, 2003 10:56 AM
Bill,
As I see it, the question should be answered differently for the light and electron microscopy cases. I have nothing to add to Peter Van Osta's comments on the light microscopy case. However, I think I can comment productively on the electron microscopy case. Note that it is beyond me to express the following without resorting to some mathematical formalism, so here goes:
The problem with a straightforward examination of pixel capacity and readout noise in the case of electron image capture is that the sequential image conversion (EM electron to light to CCD electron to DN) constitutes a stochastic chain of neighboring and hence non-independent detectors. The non-independence means that it is not possible to do a meaningful noise budget for the pixel in the CCD and it's mapped effective pixels at the scintillator and in the optical chain. As an example, early excitement over the possibility of measuring the gain of a CCD camera by measuring the shot noise in a uniform image met with the discovery that the noise was much less than could be accounted for by shot noise. The explanation was in the mixing of the light signal between neighboring pixels. Several attempts ensued to manage the mixing by means of a mixing factor. These attempts all failed due to the fact that noise and mixing occurred interspersed throughout the stochastic chain of image conversion and transfer - and were therefore inseparable.
It turns out that the non-independence of physical pixels can be dealt with through a conversion to Fourier space due to the convolution theorem. Pixel mixing in real space transforms to independent scaled channels in Fourier space and noise budgeting becomes possible once again. The formalism for this is stated (exactly as in Paul Voyles' email) but with an additional variable: spatial frequency.
After painful efforts to clarify these issues for electron microscopy it was discovered (by Hans DeRuijter) that an extensive literature existed already in the Medical Imaging field on exactly this concept (DQE (N, S) where S is the spatial frequency). Medical Imaging references can be found by searching on the names: Ian Cunningham and Rodney Shaw. Electron microscopy references can be found by searching on the names: Hans DeRuijter and Ruediger Meyer. Rudy has published a series of both theoretical and experimental analyses of electron cameras in terms of DQE in Ultramicroscopy.
In short, the SNR of an image (transformed into Fourier space) is the signal power spectrum divided by the noise power spectrum. The DQE is defined as the square of the output SNR over the input SNR. A simple regrouping of terms gives:
Which is fairly straightforward to measure since for low-contrast, low-dose imaging, the noise power spectrum "in" is just the Poisson noise in the incident beam, and since that noise distributes evenly in Fourier space (it is "white"), the NPSin = N. The NPSout is just the power spectrum of the output image provided the image is recalibrated in units of EM electrons before transformation. (The Fourier transform must be normalized to conserve sum of squares.) The MTF can be measured by the "tilted edge method" using a relatively clean beam-stop edge.
It can also be seen that the SNR out of a detector is just sqrt (DQE * specimen contrast * dose). Thus DQE can be seen as the "dose-efficiency" of a detector as a function of dose and spatial frequency. Twice the DQE means you need half the dose at any given spatial frequency of interest.
Hope this helped,
Paul
Paul Mooney
Program Manager, Imaging Product Development Gatan, Inc. 6678 Owens Dr. Pleasanton, CA 94588
tel: 925 224-7335 FAX: 925 463-0204
-----Original Message----- } From: Bill Tivol [mailto:tivol-at-caltech.edu] Sent: Monday, July 28, 2003 4:40 PM To: microscopy-at-sparc5.microscopy.com
On Monday, July 28, 2003, at 02:21 AM, Peter Van Osta wrote:
} Hi, } } I mostly have to deal with digital microscopy in extreme low-light } conditions (fluorescence microscopy) and the quality of the images is } very important for the subsequent quantitative analysis. } } I would like to know if the following formula for calculating the } maximum Signal to Noise Ratio (SNR) for a CCD camera still holds (in } general). Photon noise or photon shot noise, refers to the inherent } natural variation of the incident photon flux and a such this form of } noise limits the maximum SNR for a given CCD-element and the CCD camera } ? } } If we consider the capacity of one CCD element of a (Silicon-based) } CCD, } the capacity for photoelectrons "C" relates to the "maximum SNR" of a } CCD camera: } } SNRmax = 10 log10(C) } } In general a CCD camera seems to have a photoelectron capacity per } square micron of about 700 photoelectrons/µm^2. So, the maximum SNR of } a } CCD camera is defined by the "well" capacity of each individual } CCD-element (pixel) and cannot be surpassed. So, there is no substitute } for square microns per CCD-element to get a higher maximum SNR ? } Increasing the SNR in this way however relates inversely to the spatial } sampling ? } } The total well capacity divided by the noise of the camera system, } gives } the dynamic range of the camera. The higher the dynamic range, the less } noisy the camera becomes. So, there is a direct relation between } individual CCD-element surface area and dynamic range ? } } There is a lot more to be said on this in digital micrscopy, but I want } to know wether this relation is (still) valid and that I did not miss } some recent developments. What about the varying quantum efficiency } depending on the wavelength of the incident photons ? What about } coating } CCD-cameras to improve the quantum efficiency in the shorter } wavelengths } or blocking near-infrared for which CCD's seem to be very sensitive in } order to improve the SNR over the entire visible spectrum ? } } Best regards, } } Peter Van Osta } Dear Peter, Please post the replies you get to the list. We have a similar problem, since we are doing low-dose EM. The questions asked in your post are the kinds of things we have been considering. We have also considered such problems as readout noise and whether the electronics gain can be optimized to give better S/N. We also need to get high spatial frequencies, so small pixel size and narrow point-spread function are important to us as well. 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
I am looking for a new/used EM tissue processor. Any suggestions/recommendations/info are highly appreciated.
TIA,
GN
Gang (Greg) Ning 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
Caroline Schooley, the project MICRO coordinator, gave me this advice when I turned to her with a similar question some time ago:
First, I went to the Project MICRO webpage http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html {http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html} .
You’ll find very helpful the section “Buying microscopes†in the Microscopic Explorations part, as well as great projects ideas for the class.
For the microscope purchase, you can go to http://www.microscopeworld.com/
There are several options according to your needs and budget. I bought a few #185 monocular dissecting scopes for $75 each (what a deal!) for the class of second graders, both the kids and the teachers love them. The parents of the kids in the class decided to sponsor this, and contributed about $30 per family - that might be an option in a class with no funding for new equipment.
Good luck,
Alice.
Alice Dohnalkova Pacific Northwest National Laboratory Richland, WA 99352 (509) 372-0692
-----Original Message----- From: by way of Ask-A-Microscopist [mailto:hoganbecky-at-hotmail.com] Sent: Tue 29/07/2003 07:26 To: Microscopy-at-sparc5.microscopy.com Cc: Subject: Ask-A-Microscopist: teaching gifted 5th graders
------------------------------------------------------------------------ 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 (hoganbecky-at-hotmail.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, July 28, 2003 at 20:07:39 ---------------------------------------------------------------------------
Question: I will be teaching gifted 5th graders in a new math/science program this year and need to know what type of microscope would be best for my program. Also, where can I buy a microscope for a reasonable price? Do colleges ever donate used microscopes to schools? I am working with a shoestring budget of $0.00 so I will have to make a personal purchase in order for my students to use this wonderful teaching tool. Thank you for asssisting me with this matter. I want to offer the best science program possible! Becky Hogan
The Department of Molecular, Cellular and Developmental Biology and the Neuroscience Research Institute of the University of California at Santa Barbara are sponsoring a Microscopy and Digital Imaging workshop on Sept. 15-19, 2003. Participants will get hands-on instruction and experience with advanced light, fluorescence, and confocal microscopy and imaging techniques that are essential tools for research. For further information and registration, please check our web site:
'Once again the MSA Education Committee is organizing Exhibitor Demonstrations and Tutorials on Tuesday, August 5 from 6:00 to 8:00pm in the Exhibit Hall. These mini-seminars or tutorial demonstrations are held in the booths of the participating companies after the Hall is closed to non-participants at 5:00 pm.
Signup sheets with titles and descriptions are at the MSA Education table in the MSA Mega Booth. When you sign up you will be issued a ticket, which you will need to renter the Hall after it is closed. You need to sign up no later than noon on Tuesday. The number of attendees is limited, so visit the MSA Education table soon, since the demonstrations get filled up quickly!
Here's a list of participating Exhibitors and titles:
Accurion Scientific Instruments "Atomic Force Microscopy meets Optical Microscopy"
Asylum Research "Simultaneous AFM and Fluorescence Imaging with the MFP-3D AFM"
Bitplane, Inc. "4D Particle Tracking"
Delong America "LVEM5: Applications and Operations"
EDAX "Stereo Imaging - how to collect and view images in 3D" "What you can do with an Orientation Imaging Microscopy EBSD System"
Emispec Systems, Inc. "A new software architecture for SPM imaging and spectroscopy; a joint development between Emispec System, Inc. and Molecular Imaging Corporation"
Extec Corp./A4I America "SIMPLICITYŠIntegrated Image Analysis commencing with Sample Preparation"
FEI Company "Overcoming Hurdles in High-resolution TEM Tomography"
Gatan, Inc. "The Gatan User Tutorial"
HACKER Instruments & Industries Inc. "Advances in Microwave Specimen Preparation for TEM"
Hitachi High Technologies America, Inc. "FE-SEM: SE/BSE Mixing- Pros and Cons"
Horiba, Inc. "Transmission X-Ray Analysis on your Desktop"
Imago Scientific Instruments Corporation "Introducing Imago's LEAPTM Atom Probe Microscope: innovative microscopy for rapid 3-D imaging and analysis at the atomic scale"
Microscopy/Microscopy Education, Inc. "Calibrating Fluorescence"
Molecular Imaging "Simultaneous Sample Topography and BioMolecular Force Recognition Mapping with a Versatile New Single Molecule Force Recognition Tool Kit"
Nanoptek Corporation "Photon Tunneling Microscopy: Sub-nanometer profiling in real time"
Nanotech-America "Breaking the bounds of AFM: New Hybrid Techniques"
Quantum Dot Corp "Lighting Up Cells and Tissues with Qdot Labels"
RMC-Boeckeler Instruments Inc. "Cryofixation & Freeze Substitution - A Complete Solution"
SensIR Technologies "The IlluminatIR - Combining light microscopy with infrared spectral analysis"
Soft Imaging System (SIS) "Laboratory Image Management"
South Bay Technology, Inc. "Ion Beam Sputtering: Applications to Electron Microscopy"
To All Those who are attending, or may be interested in attending, the Annual MSA "Microscopy & Microanalysis" meeting in San Antonio, TX next week (August 3-8), there will be:
A SPECIAL SESSION ON
LEGAL ISSUES
ASSOCIATED WITH MICROSCOPY AND RELATED TECHNOLOGIES
IN A
PUBLIC POLICY FORUM
"PATENT AND INTELLECTUAL PROPERTY ISSUES FOR MICROSCOPISTS"
Henry M. Schaffer Esq., MSA Counsel
A POTPOURRI OF PATENT TOPICS OF INTEREST TO MICROSCOPISTS INCLUDING:
* WHAT CAN AND CANNOT BE PATENTED
* CAN ANYONE USE YOUR PATENT - FOR FREE?
* WHAT ARE THE DIFFERENCES BETWEEN PATENTS AND COPYRIGHTS?
* HOW LONG DOES A PATENT LAST? CAN IT BE RENEWED?
* PATENTS AND STANDARDS - A RISKY MIX
} WEDNESDAY AUGUST 6, ROOM 206 A&B, 2.00 PM, CONVENTION CENTER, SAN ANTONIO
FOR MORE INFORMATION: CONTACT - www.msa.microscopy.com/MMHomePage.html
-- Peter Ingram Chair, MSA Public Policy Committee Duke University Medical Center Box 90319 LaSalle Street Extension DURHAM NC USA 27708-0319
Hi Everyone, I have a bunch of apertures for a Zeiss TEM (the TEM has been decommissioned). Most are from Ted Pella, Inc., cat #69020-30. I don't know if they are new or not, but they are all the same price (free). I will be in San Antonio for M&M, and if anyone who will also be in attendance wants them, drop me an email and I'll bring them along. Otherwise, they get pitched. Randy
Randy Nessler University of Iowa Central Microscopy Research Facility Phone 319-335-8142 Fax 319-384-4469
} Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (hoganbecky-at-hotmail.com) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, } July 28, 2003 at 20:07:39 } --------------------------------------------------------------------------- } } Email: hoganbecky-at-hotmail.com } Name: Becky Hogan } } Organization: Summerville Elementary } } Education: K-8 Grade Grammar School } } Location: Summerville, South Carolina } } Question: I will be teaching gifted 5th graders in a new } math/science program this year and need to know what type of } microscope would be best for my program. Also, where can I buy a } microscope for a reasonable price? Do colleges ever donate used } microscopes to schools? I am working with a shoestring budget of } $0.00 so I will have to make a personal purchase in order for my } students to use this wonderful teaching tool. } Thank you for asssisting me with this matter. I want to offer the } best science program possible! } Becky Hogan } } --------------------------------------------------------------------------- Becky -
In general, used college microscopes won't work well for you at the 5th grade level. Please look at the detailed advice on the Project MICRO website (URL below). If you have questions after reading that advice, I'll be happy to answer them directly. Thank YOU for your enthusiasm!
Caroline
-- Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jwirth4-at-juno.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 30, 2003 at 10:30:25 ---------------------------------------------------------------------------
Email: jwirth4-at-juno.com Name: Jim Wirth
Organization: Piedmont Community College
Education: Undergraduate College
Location: Charlottesville, VA 22903
Question: Hello, When using an inverted microscope, is it typical to focus on the bottom or top plane of the stage? Is it necessary to drop the specimen to the lower plane?
Is there a typical stage opening and insert dimension across the major manufacturers?
John Hoffpauir has sent me his comments. We believe his comments are quite plausible.
John Arnott
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: sales-at-laddresearch.com ----- Original Message ----- } From: JHoffpa464-at-aol.com To: sales-at-laddresearch.com Sent: Wednesday, July 30, 2003 1:40 PM
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (asmus-at-centre.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 30, 2003 at 13:41:25 ---------------------------------------------------------------------------
Email: asmus-at-centre.edu Name: Steve Asmus
Organization: Centre College
Education: Graduate College
Location: Danville, Kentucky, US
Question: I'm comparing 2 fluorescence microscopes and would like feedback on how these scopes compare or any info on rankings of these scopes. I'm choosing between a Nikon E600 and an Olympus BX51. They have identical capabilites and similar prices. I am also interested in adding confocal capabilies eventually, and both appear to be upgradeable. Any advice from someone familiar with these scopes would be appreciated.
It is a very good and practical question! There are more than one source of the Cu characteristic X-rays your described. The TEM column is not "clean", since there are bremsstrahlung X-rays and uncollimated electrons, as a result of the electrons interacting with column components such as the diaphragms and polepieces. The bremsstrahlung X-rays and uncollimated electrons can strike the Cu grid and your specimen, producing Cu x-rays which are not distinguishable in your EDS spectra.
Some of the transmitted electrons are scattered at sufficiently high angles and strike other TEM column components such as the objective lens polepieces and objective diaphragm if it is not removed. They would also give rise to spurious X-rays. You can also check the probe size by directly imaging the probe, and check if there is any long tail. To get a feel of the background X-rays, you can place the beam through a hole in the sample, and collect a spectrum, which would show some peaks.
I hope this is somewhat helpful to you.
Kind Regards.
Wentao
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Dear Fellow Listers, There is quite a good book -Artifacts in Biological Electron Microscopy edit Richard Crang and Karen Klomparens (Plenum Press 1988) (dont know if still in print though) which has pictures of 'pepper' caused by embedding, fixation and staining as well as a host of other oddities. I have a few questions though as I was surprised to see such a question posted, Where do electron microscopists train and get experience these days, many labs seem to have one routine method and they stick to it? Is there a good web site that maybe images (descriptions just don't do it,) such as those found in the above book, could be posted to?
Cheers
Gillian Brown Histology Section, Asthma Biology RI CEDD http://ukdiscovery.gsk.com/histopathology/ ----- Forwarded by Gillian 2 Brown/PharmRD/GSK on 31-Jul-2003 09:44 -----
"Ladd Research" {ladres-at-worldnet.att.net}
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To: "Microscopy Listserver"
cc: Subject: Re: Contamination in resin sections---TEM
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John Hoffpauir has sent me his comments. We believe his comments are quite plausible.
John Arnott
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: sales-at-laddresearch.com ----- Original Message ----- } From: JHoffpa464-at-aol.com To: sales-at-laddresearch.com Sent: Wednesday, July 30, 2003 1:40 PM
Jim;
Is this inverted microscope for potted cross-sections? I generally use an inverted microscope for encapsulated cross-sections so that the polished face is perpendicular to the objective lens and the plane of focus would be that face or the top of the stage.
Peter Tomic Agere Systems Allentown, PA
-----Original Message----- } From: jwirth4-at-juno.com [mailto:jwirth4-at-juno.com] Sent: Wednesday, July 30, 2003 2:06 PM To: Microscopy-at-sparc5.microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jwirth4-at-juno.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, July 30, 2003 at 10:30:25 ---------------------------------------------------------------------------
Email: jwirth4-at-juno.com Name: Jim Wirth
Organization: Piedmont Community College
Education: Undergraduate College
Location: Charlottesville, VA 22903
Question: Hello, When using an inverted microscope, is it typical to focus on the bottom or top plane of the stage? Is it necessary to drop the specimen to the lower plane?
Is there a typical stage opening and insert dimension across the major manufacturers?
When I use TEM sections in SEM, I see Cu peaks also. I place grids on a carbon mounts, so backscattered electrons from this substrate generate the peaks very similar to ones we see with TEM.
Vladimir M. Dusevich, Ph.D. Electron Microscope Lab Manager 3127 School of Dentistry 650 E. 25th Street Kansas City, MO 64108-2784
} -----Original Message----- } From: Wentao Qin [mailto:wentao_qin-at-yahoo.com] } Sent: Thursday, July 31, 2003 1:08 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: Re: Cu peaks in STEM/EDS } } Hi Dr. Pan, } } It is a very good and practical question! There are } more than one source of the Cu characteristic X-rays } your described. The TEM column is not "clean", since } there are bremsstrahlung X-rays and uncollimated } electrons, as a result of the electrons interacting } with column components such as the diaphragms and } polepieces. The bremsstrahlung X-rays and } uncollimated electrons can strike the Cu grid and your } specimen, producing Cu x-rays which are not } distinguishable in your EDS spectra. } } Some of the transmitted electrons are scattered at } sufficiently high angles and strike other TEM column } components such as the objective lens polepieces and } objective diaphragm if it is not removed. They would } also give rise to spurious X-rays. You can also check } the probe size by directly imaging the probe, and } check if there is any long tail. To get a feel of the } background X-rays, you can place the beam through a } hole in the sample, and collect a spectrum, which } would show some peaks. } } I hope this is somewhat helpful to you. } } Kind Regards. } } Wentao } } __________________________________ } Do you Yahoo!? } The New Yahoo! Search - Faster. Easier. Bingo. } http://search.yahoo.com } }
"Biomedical Electron Microscopy" by Maunsbach and Afzelius (Academic Press, 1999) has lots of examples of artifacts. It is an excellent book. The Crang and Klomparens book mentioned below is also excellent. Glutaraldehyde in tissues can react with osmium to form artifacts, as John mentioned earlier. I did not know that it was buffer dependent.
} Dear Fellow Listers, } There is quite a good book -Artifacts in Biological Electron Microscopy } edit Richard Crang and Karen Klomparens (Plenum Press 1988) (dont know if } still in print though) which has pictures of 'pepper' caused by embedding, } fixation and staining as well as a host of other oddities. } I have a few questions though as I was surprised to see such a question } posted, Where do electron microscopists train and get experience these days, many labs seem to have one routine method and they stick to } it? Is there a good web site that maybe images (descriptions just don't } do it,) such as those found in the above book, could be posted to? } } Cheers } } Gillian Brown } Histology Section, Asthma Biology } RI CEDD } http://ukdiscovery.gsk.com/histopathology/ } ----- Forwarded by Gillian 2 Brown/PharmRD/GSK on 31-Jul-2003 09:44 ----- } } } "Ladd Research" {ladres-at-worldnet.att.net} } } 30-Jul-2003 20:01 } Please respond to "Ladd Research" {sales-at-laddresearch.com} } } } } } To: "Microscopy Listserver" } } cc: } Subject: Re: Contamination in resin sections---TEM } } ------------------------------------------------------------------------ } 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 **********************************************
Many thanks to all of you who responded to my first query on this topic. I was deliberately vague, hoping to get the widest range of ideas. We got some very good thoughts and we are trying all of them, in one form or another.
To be more specific, we are getting pepper in nearly every tissue we look at, but not uniformly everywhere in the tissue. I will email images to anyone requesting them. For example, in retinal tissue it most consistently shows up in the rods. Our standard processing involves 2% gluteraldehyed/2% paraformaldehyde in 0.1M cacodylate buffer, but we often use 0.17M cac buffer, Sorenson's phosphate buffer, and varying concentrations of fixatives, depending upon the project. Immunocytochemical processing is generally done in 4% paraformaldehyde/0.1% glut in cacodylate or phosphate buffers, usually with a small amount of CaCl2.
Here is what we have tried:
1) Redoing our water purification system and trying water from other labs (we're now on our 4th type of water, including purchased distilled water, reverse osmosis feed water into a Millipore Simplicity polishing system; two-tank-in-series deionized water system feeding into the Millipore; and house deionized water run through another lab's wall-mounted Millipore polishing system. No other lab using the latter water supply is reporting any problems; 2) Rewashing ALL of lab glassware; 3) Remixing ALL of our solutions, using fixatives from different lot numbers; 4) Remixing ALL of our resins; 5) Processing with and without secondary and tertiary fixatives (osmium and uranyl acetate)in all combinations; 6) Looking at both stained and unstained sections (Pb and UA); 7) Looking at different tissue types; 8) Embedding in different resins, including Epon/Araldite, Spurr's, pure Epon (actually Embed 812), LR White, and Araldite. 9) Cutting on different diamond knives, with different types of syringe-filtered water in the boats.
So far the cleanest sections have been in LR White, while the same tissue processed for Epon/Araldite showed contamination, but the use of several other resins seems to indicate that resin is probably not the issue. BUT, the LRW material was processed with a much lower percentage of gluteraldehyde in the fixative. But wouldn't the use of different lots of gluteraldehyde argue against this as the culprit?
We have been able to successfully remove or minimize the contamination by post-treating sections with 2% periodic acid, which also chews up our copper grids and adds its own yuck to the process. However, we are now cutting on gold and nickel grids to give us this option until we figure out what's going on here.
A genuine puzzler and a major nuisance. I guarantee that when we get this solved, I will post a detailed cure in hopes of saving anyone else this nightmare.
Thanks much for any further advice!
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
The "red-gren-red" indicator light for density adjust always seems to remain lit as red on the left side, making me believe that there is a problem with the light meter.
Changes in brightness, and exposure time don't seem to have any effect on the density readings.
Is there anyone else who has had problems with their light meter, and knows any simple remedies to this problem?
The camera does advance, and the beams seems to go on and off making an exposure for the time set by the exposure time, so it can take pictures. It's just that we seem to making the pictures in the dark.
A course on Quantitative Image Analysis will be taught in San Francisco, November 6-7, 2003 by John Russ and the Reindeer Graphics staff. The course emphasizes practical solutions to imaging problems and includes 2 days of intensive training in current techniques, including an evening open laboratory where participants are encouraged to bring and work with their own images. The course is taught using the Fovea Pro software {http://www.reindeergraphics.com/foveapro} , and participants receive both a copy of Fovea Pro and a copy of "The Image Processing Handbook" 4th Edition (CRC Press).
The course syllabus and registration information are available at the Reindeer Graphics website. {http://www.reindeergraphics.com/courses} Further information may be obtained from the website or by contacting Reindeer Graphics directly at courses-at-reindeergraphics.com or by telephone at 828.252.7515
Dear Jim, Our inverted microscopes are meant to focus on the face of the sample that is face-down on the stage, so we focus on the top surface of the stage. These microscopes are used for large, reflected-light samples, because the design of the stage does not restrict the size of the sample. As for openings, our microscopes come with a variety of openings available. These are discs with an outside diameter of about two inches, that fit into an opening in the stage, and a selection of inside diameters, to accommodate different sized samples. The top of these discs is flat, while the bottom of them is concave, to allow the objectives to swing by. I hope this helps. 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: {jwirth4-at-juno.com} To: {Microscopy-at-sparc5.microscopy.com} Sent: Wednesday, July 30, 2003 11:06 AM
Garry
The H7000 that I use has another exposure meter display inside the column just behind the screen. this consists of two red LEDs (ie no green middle. If the two come on together that is a correct exposure but increasing or decreasing the beam density on the screen should make one or the other come on. I apologise if you have checked this but if the column meter works then it sounds like it is the other meter. Also have you checked that no one has 'mucked about' with the meter calibration settings.
If everything else checks out then maybe it's just the link from the central screen that has broken - remember this is a current density rather than light meter.
I hope this helps.
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: Garry Burgess {GBurgess-at-exchange.hsc.mb.ca}
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Hi Randy Do you have the same problem when you use gold or nickel grids? Some time ago I worked on immunogold technique in tissue and I had problem with pepper/dirt film over the sections. I used nickel grids that were bought ages ago. I bought a new batch, cleaned it and sections were clean, no pepper/dirt film. Dorota
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