On a related subject, has anyone else noticed that Si samples get darker when you start to ion mill them? I usually get TEM cross-sections down to ~8-10 um, and when the sample goes in the mill it's an amber/red colour (from the rather poor light shining through it). Ten minutes later it's almost opaque. Maybe it's just the surface roughness increasing; or perhaps the effect of an amorphous layer? And while I'm on the subject of looking through samples in the ion mill, has anyone come up with a way of seeing how thick GaAs or InP are while milling? GaAs seems to become transparent at about 1 um (ish), and InP always looks dark no matter what. A real pain when setting the timer!
Cheers,
Richard Beanland GMMT Ltd., Caswell, Towcester, Northants NN12 8EQ UK
} Recently we were asked to analyze the uniformity of hydrogel coating on the } surface of the 100 micron particles. The hydrogel consists of 10% bifunctional } polyacrylate and some additives. The rest, of course, is water. We froze the } samples in liquid N2, and sublimed frozen water using freeze dryer. After SEM } observation, teh coatings looked collapsed anyway. This is not what our } customer wants. AFM analysis of the obtained samples yields similar } results. In } addition, it is compounded by the fact that the particles are spherical } and the } curvature really throws off the small magnificAtion images. We recommended the } customer a liquid Tapping Mode AFM (we are not eqiupped with it). Is there any } other way we can prepare and analyze the surface morphology of these samples? } This is my first time addressing the ListServer, but there is always time for } first. } Thanks for possible help, } Alex Mejiritski } Ph. D. Student } Center for Photochemical Sciences } Bowling Green State University } Bowling Green , Ohio 43402 } (419) 372-7830 }
How about an Environmental SEM? You could look at the sample in a fully hydrated state, then slowly dehydrate, to reveal sub-surface structure.
Ed is quite right, one space plus one line. If possible, the accuracy is improved if many spaces/lines and are included. A stage micrometer could be used to establish the magnification and for a given lens combination that figure would be "permanent". For some uses it could be most useful to then establish the length of the negative in micrometers and use that as a standard, but some people would prefer large latex spheres incorporated with the specimens. For low power SEM and reflected LM there is a 0.01mm graduated scale available for calibration of those instruments (See Pelco or ProSciTech). Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 350+ Links, MSDS ************************ http://www.proscitech.com.au } } } Can anyone suggest what to use as a length standard with a 100x } } } objective? I intend to print out an image of this standard when I print } } } images of samples, in order to determine final magnification. I am a little } } } concerned with the use of my micrometer slide (10 microns between } } } graduations) because the width of the individual lines is significant } } } compared to the distance being measured between lines. Also there is } } } no tolerance stated for the separation between the lines. I can think of } } } using calibrated latex beads suspended in water, but the refractive index } } } difference between latex and the aqueous medium (or air) causes a lot of } } } problems. Are there other standards I can use? } } } } } } Thanks } } } Richard } } } Richard_Thrift-at-Depotech.com } } } } } } Richard, } } } } I was under the impression that micrometer slides were meant to measure } } from one side of a line to the corresponding side on the adjacent line, } } not between the lines. Have I been mis-led? } } I thought the line thickness is accounted for in this manner. } } } } } | } | not | { } | } } } } Cheers } } ed } } } } Edward J. Basgall, PhD } } The Pennsylvania State University } } Surface Chemistry Group ejb11-at-psu.edu } } Materials Research Institute Building Ph: 814-865-0493 } } University Park, PA 16802-7003 FAX: 814-863-0618 } } } } } }
What about a piece of black and and white photographic negative film for a cheap test specimen? On a good microscope, the silver grains appear with sharp edges. I don't have a toy microscope around to compare. But film is widely available and cheap, dry, and you don't even need a microscope slide or coverslip. No regular pattern to detect barrel or pincushion distortion, but these probably wouldn't matter to most users anyway.
BTW, someone around here made up a bunch of cheap stage micrometers by photographing a metric ruler from a distance with a 35 mm camera. With the reduction, the one mm divisions on the scale are 100 micrometers apart on the negative. These were then just cut up and glued to microscope slides. They are a little fuzzy but if you measure center to center of each line it is close enough. Perhaps one could photograph a grid to make a test specimen for distortion.
Gary Radice, Associate Professor gradice-at-richmond.edu Department of Biology 804-289-8107 (voice) University of Richmond VA 23173 804-289-8233 (FAX)
David - use the find function on your browser and search for "Protist" on our link page. That link takes you a sites at the Uni of Montreol which has the very things you are looking for. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 350+ Links, MSDS ************************ http://www.proscitech.com.au
---------- } From: David Webb {davehawaiiedu-at-msn.com} } To: Microscopy-at-sparc5.microscopy.com } Subject: Images of Algae and Sea Weeds } Date: Monday, 30 June 1997 5:12 } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } I am looking for all types of images (light & EM) which deal with algae, } especially sea weeds. I plan to teach a course in which I will compare and } contrast vegetative and reproductive adaptations of algae with land plants. } The emphasis will be at the light microscope level, but SEM and TEM images } which deal with major aspects of form and function would be welcome.
Cleaning grids? Go your hardest if you have nothing better to do: Collect grids in a 20 or 30ml glass vial. Soak them in chloroform, give one change in chloroform, pour off solvent, replace with ethanol or acetone, replace with water, add drops of detergent, ultra sonicate for a short time. Rinse with water, etch for a moment in weak acid (10% acetic or 0.2N HCl) rinse in distilled water. Pour water off. Use washbottle with water (or for faster drying ethanol) to flush grids into a Petrie dish lined with filterpaper; pipette off excess fluid. Sit dish in an incubator until dry. Thousands of grids can be treated in one batch but the joy is in sorting the grids and throwing the bad once out. We sell standard copper grids at A$10/vial/100; which is a little over US$7. In a labour-costly country, with the odd exception, cleaning grids is a waste of time. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
} I don't know about the first, I've never had really good results from } cleaning off coated grids, but as to the second; in the absence of a Glow } Discharge apparatus you might get some result out of using a Zerostat } Antistatic Pistol. It is not so reliable as Glow Discharge and takes some } practice to obtain neutral charge grids. } } They are still available as far as I know. We bought one recently for } zapping our Balance after frustrating sessions of attempting to weigh out } powders which were flying all over the bench. It cured that. } } If you want to see if it works before buying, find yourself a Black Vinyl } Record collector and borrow one. They often use them for discharging static } on Discs before playing. I used one myself before going over to CD's many } years ago. } Regards } Stephen Griffiths } } {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} } Stephen Griffiths e-mail:- s.griffiths-at-ucl.ac.uk } Visual Science Department Phone:- 0171 608 6914 } Institute of Ophthalmology Fax:- 0171 608 6850 } Bath Street, London. EC1V 9EL } {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} } } } Just wondering about a couple of things. } } First the different ways there are to clean grids that have a } } film (2% parlodion), and a carbon coat, but no sample. } } Second the best way to reduce or eliminate static on coated grids, } } without using a glow discharge apparatus. } } Thanks, } } Maya Moody }
We still have a "bunch" available. See our on-line catalogue. Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au ---------- } From: joyce craig {bafpjec-at-csu.edu} } To: Listserver {Microscopy-at-sparc5.microscopy.com} } Subject: Cleaning grids (Zerostat) } Date: Tuesday, 1 July 1997 15:18 } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Does anyone know where I can purchase a Zerostat antistatic pistol? My } old source no longer carries them, and I find them really useful in dry } sectioning in low humidity (not a problem in Chicago in the summer, but } in the winter...) and in keeping grids from bouncing onto the lid of the } Petrie dish. } Cheers from } Joyce Craig } Chicago State University
From Mark Lund: } } --- } } Being the token optical engineer on this list makes me the token lens } designer, too :) Although I have never designed an immersion type } microscope objective the general trend is that as you go up in index } of refraction the job gets easier. A high index "glass" that won't } etch should make the lens design practical. Of course the high index } of diamond should make it possible to get higher NA's
But diamond has a high dispersion relative to quartz (0.044 vs 0.013). Wouldn't this make correcting for chromatic aberration difficult?
How about corundum? RI = 1.762 -1.77, dispersion = 0.018
On a separate note: as the "token optical engineer", have you been following the thread about microscopes for kids and lower-cost-but-still-quality ones for amateurs?
Phil
} Sic Hoc Legere Scis Nimium Eruditionis Habes { Philip Oshel Station A PO Box 5037 Champaign, IL 61825-5037 (217) 355-1143 oshel-at-ux1.cso.uiuc.edu *** looking for a job again ******************
The index of refraction of diamond is 2.4173. I don't know as much about the theory of microscope optics as I should but I do know that this plays a role.
Joe Tabeling Delaware Diamond Knives 3825 Lancaster Pike Wilmington, DE 19805 302-999-7476
The server was down for nearly a full day while I replaced the main drive. Expect a hic-cup or two while things settle in.
The subscribe/unsubscribe functions should (?) be functioning tomorrow AM. For those of you that have tried to unsubscribe you should be "off" the list tomorrow.
I apologize for the inconvenience...
Nestor Your Friendly and Blurry Eyed Neighborhood SysOp
We want to do immunohistochemistry on human heart muscle tissue cryosections. Which embedding medium should be used for cryosectioning on a Leica cm 3000 kryostat for light microscopy? Are there simple recipies and procedures (cryoprotection...)? Which freezing techniques should be used? We get the tissue in the operating theatre which freezing technique can be used directly there? I sthere a freezing technique for both, cryosectioning and RNA-preparation of the same tissue-block.
Thank you all for your answers
Christoph Guenther Klinikum der Charite der Humboldt Universitaet Ziegelstrasse 5-7 10117 Berlin /Germany Tel.: +49 30 2802 6327 Fax + 49 30 2802 6608
} There's a new CD-ROM (Windows & Mac) out } from a reputable source (Center for Bioengineering, U. of Washington, } Seattle) that may do some good (tho I haven't had a chance to review it yet } for the Project MICRO bibliography). Microscopy-Tutor, Lippincott-Raven, } ISBN 0-7817-1217-3, $195.00. http://www/lrpub.com, 800-777-2295.
The URL for Microscopy-Tutor is "http://www.lrpub.com/media/m1208.htm".
Here's an advertising blurb from the web site.
============================================== Microscopy-Tutor, CD-ROM for PC and Macintosh, with Two-Color Insert, by The Department of Laboratory Medicine and the Center for Bioengineering, University of Washington, Seattle, WA.
This interactive computer program guides students of biology and the health professions through the basic concepts of bright field light microscopy, developing and refining the user's knowledge by providing a more active role in learning. Simple, approachable, and largely qualitative, Microscopy-Tutor uses three-dimensional animation to simulate a microscope with an integrated illuminator and adjustable field diaphragm. The CD-ROM's animated diagrams are more accurate than those found in most university-level texts, but are also easy-to-understand because key concepts and equations are presented visually rather than symbolically. Since using Microscopy-Tutor feels more like operating a microscope than a computer it's successful in presenting changes in dynamic processes that occur during alignment and use of the microscope. Rather than explaining what happens through the use of words and pictures, the user -- who learns by doing -- can see what happens and better understand the implications. In some sections two animated perspectives are synchronized, allowing the viewer to change microscope settings and simultaneously see the resulting image. Interactive quizzes test the user's knowledge and accent areas for improvement. ==============================================
The information below is slightly wrong. Not a big deal, but in the interest of science I thought I would set things slightly straighter :)
Azriel Gorski wrote tht McCrone and McCrone wrote:
} Lenses have several types of aberrations which will cause the loss of } detail unless corrected for. Toy microscopes are not. } } Spherical aberration - "is especially apparent in lenses having sperical } surfaces. Light paths near the center of the lens focus at different } points compared to light paths near the periphery."
The name "spherical aberration" has historical roots in astronomy where a spherical mirror has this aberration but a parabolic mirror does not. In some instances a spherical mirror will have no spherical aberration wheras a parabolic mirror will have maximum spherical aberration. The aberration really has nothing to do with the spherical surfaces of lenses. The confusion comes because making a surface "aspheric" can cure spherical aberration in many instances.
} Field curvature - "is a natural result of using lenses with curved } surfaces. The image plane produced by such lenses will be curved. This } kind of image occurs in microscopy unless plano (flat-field) objectives } are used."
Actually, field curvature is a natural result of the geometry of the real world. Since an object at the edge of the field of view is farther from the lens "center" it will tend to be focussed closer to the lens than an object on axis. This naturally leads to field curvature unless the designer makes the lens weaker for off axis points. It has nothing to do with the lenses being curved. The confusion comes from the formula for Petzval curvature, which has lens powers in it.
To the amateurs - welcome! We need you to keep us fresh and excited in what we do.
The alk phos is more sensitive because it will continue to reduce substrate long after peroxidase has quit.
Bob Morphology Core
On Fri, 27 Jun 1997, Tom Phillips wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } We are starting some in situ work with digoxigenin labeled probes. A lot } of the kits and studies seem to use Alkaline Phosphatase labeled antibodies } to detect the digoxigenin. I am curious why they seem to prefer Alk Phos } over peroxidase coupled antibodies which are the more standard } immunocytochemical choice. Anybody have any thoughts on this? } } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } 3 Tucker Hall } University of Missouri } Columbia, MO 65211 } (573)-882-4712 (voice) } (573)-882-0123 (fax) } } }
Talk about a step backwards... Historically, some objectives were made with different gems in the 1800's (I believe, I don't have my history references at work), but they suffered from several problems. First there are problems with the optical clarity of the gem, color in the case of rubys and garnets, polishing the surfaces to the optimum shape and of course, cost. In the case of polarized light, the double refractive index stones were disastrous. This development did not lasted very long as it was too impractical. Also remember, an immersion system requires more than oiling the front lens of the objective. The condenser should be oiled to the slide as well as the objective. In these systems the resolution is limited by the lowest refractive index, which for a diamond condenser and objective could be the immersion oil.
I think Aldrich still sell them, catalogue number: Z10,881-2, I've only got an old catalogue, and Aldrich might be your old source, so this might not be too useful.
Ray
At 10:18 pm -0700 30/6/97, joyce craig wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Ray Hicks ________________________________________________________________________ |University of Cambridge |Tel 01223 330149 | |Department of Medicine |Fax 01223 336846 | |Level 5, Addenbrookes Hospital |e-mail {rh208-at-cus.cam.ac.uk} | |Hills Road Cambridge |Web http://facsmac.med.cam.ac.uk | |CB2 |ftp server ftp://131.111.80.78 | |UK | | |_________________________________|_____________________________________|
I have looked at the discussion on recrystallization with interest. As I am an agreeable fellow, I would say that you are both correct!
It is true that amorphously vitrified pure water will devtrify at about -130 C or lower, depending partly on the rates of cooling and rewarming. The other extreme was described by MacKemzie AP (1981) Modellling the ultra-rapid freezing of cells and tissues. In: Microprobe analysis of Biological Specimens (eds) Hutchinson TE & Somlyo AP. Academic Press, NewYork, London, 397-421. In this work, which used starches and gels, some specimens could be rewarmed to about -6 C before crystal growth took place (i.e. in the higher molecular weight substances).
I published one micrograph in Scanning Microsc. 6, 715-743 (KP Ryan, Cryofixation of tissues for electron microscopy: a review....) in 1992 which showed some fish spleen tissue elements stored at -60 for 48 hopurs which showed no observable signs of crystal growth. Other results (some published only in my thesis) showed that the samwe tissue could be stored at -40 for some days but after 8 days there were signs of crystal growth in the extracellular fluid (more 'watery'?).
This is an interesting aspect of 'cryo' and deserves more attention by someone. I only did a piece of this to validate the 48 h at -80 C freeze substitution that I used. It should not induce crystal growth. The same f/s has been done at -50 C without apparent crystal development (but not by myself). In tthat case I presume the substitution got to the frozebnwater before it had time to show growth in the crystals
Dear colleagues: All of our scopes are JEOL made and we have been using filaments made by JEOL. The engineers from JEOL also recommend us to use their parts. As far as I know, however, there are companies which also sell filaments. Could anyone of you give me some information as to which source is better than the others? Thank you very much. Regards, Yuhui
} } From Mark Lund: } } } } --- } } } } Being the token optical engineer on this list makes me the token lens } } designer, too :) Although I have never designed an immersion type } } microscope objective the general trend is that as you go up in index } } of refraction the job gets easier. A high index "glass" that won't } } etch should make the lens design practical. Of course the high index } } of diamond should make it possible to get higher NA's } .But diamond has a high dispersion relative to quartz (0.044 vs 0.013). } Wouldn't this make correcting for chromatic aberration difficult? } } How about corundum? RI = 1.762 -1.77, dispersion = 0.018 } } On a separate note: as the "token optical engineer", have you been } following the thread about microscopes for kids and } lower-cost-but-still-quality ones for amateurs?
First: sorry about my first post. My computer decided that it had more important things to think about and while I was trying to get its attention the message posted before I could finish it.
The important thing is not the dispersion--many glasses have higher dispersion than diamond, but that the dispersion is much higher than you would expect for a refractive index of 2.4. Not only is the dispersion anomalous, but the partial dispersions are all well off the normal "glass line." This means that making a lens where one of the elements is diamond would be much easier than otherwise. Not only would correcting the color be easier but correcting residual color (i.e. making the lens apochromatic) would be easier. Also, for an immersion lens (with the right index matching oil) you could probably get an NA greater than 2.0, maybe even as high as 2.3. Of course the condensor and slide would also need to be high index for this to be useful :(
It looks like a fun idea, let me know if you need someone to design one!
Sapphire would be the perfect first element except for its natural birefringence--it would come out astigmatic on axis.
best regards mark
Mark W. Lund, PhD Director } } Soft X-ray Web page http://www.moxtek.com { { MOXTEK, Inc. 452 West 1260 North Orem UT 84057 801-225-0930 FAX 801-221-1121 lundm-at-xray.byu.edu
"The state is good at simple tasks, like killing people and seizing their wealth. It has far more trouble reaching inside individuals and making them good." Doug Bandow
With regard to cutting undecalcified bone samples (with and without implants), a lot of people have found the best method of achieving high straightness, low damage, thin cuts is with an annular diamond blade.
This blade involved in this process is held in a precision chuck and mounted at high tension, to give very little lateral movement, thus producing the high quality of cut. There are many exponents of the method in the hard tissue & dental research fields.
I'd be pleased to send you a relevant technical article on the use of this system for this application and also further information on the commercial version of the annular cutting machine, the 'Microslice 2'. Please send me your address details so I may get the information through to you.
(Note to Group: This product is sold by us -- Please accept this as a notification of our commercial interest in this product)
I look forward to hearing from you.
Best Regards
Tim Hazeldine Product Manager, ULTRA TEC Mfg., Inc. ...............................................................the cutting and polishing specialists 1025 E. Chestnut Avenue Santa Ana CA 92701 - 6425 USA Tel. +1 714 542 0608 Fax. +1 714 542 0627 e-mail. info-at-ultratecusa.com or ultratecex-at-aol.com .............................................................................. .....................................
} The dispersion is 0.044. } In units of per nanometer? In any case, I still don't know what effect this might have on the practicality of using diamond lenses. If the illumination were single-wavelength, there would be no problem.
} Fabrication problems? } Yes, but a colleague suggests that one can use diamond powder as a grinding agent. You at DDK would know a lot more than I. On the plus side, diamond lenses would be very hard to scratch. Yours, Bill Tivol
Recently we did an SEM/EDX analysis that gave rather baffling results. After some thought we came up with what seemed to be a plausible explanation, however, the people for whom the analysis was done remained skeptical. Therefore I am writing the Listserver for a second opinion. The sample consisted of irregular shaped alumino-silicate particles roughly 2-5 microns is size. The specimen was uncoated resulting in some charging. Analysis (windowless) at 1000x mag. resulted in a carbon peak much larger than the Al or Si peaks. At 5000x the carbon peak was smaller; i.e., comparable to the Al and Si. Using a spot mode on individual particles, no carbon was detected. However, spot mode analysis after lowering the accelerating voltage from 20 to 10 kv, showed a small carbon peak. The only explanation we could envision to fit these results was a thin carbonaceous coating on the particle surface. Is this a plausible explanation or we missing something? Does anyone have an alternative suggestion? Any help would be appreciated.
Dan Schwab Center for Microscopy and Microanalysis Bowling Green State University Bowling Green, OH
Leica (and probably other manufacturers) sells NIST (US) or NPL (UK) traceable stage micrometers, certified to tenths of a micron. Measurements are from centers of the lines. They're not cheap.
If you want to do your own certification, start with some regular fine grid, perhaps a dime store holograph, measure it against some standard you trust, and figure out the spacing.
I have just today found a company that supplies the Zerostat antistatic pistol in the U.S.
Bellex International Voice: (302) 761-9885 FAX: (302) 761-9896
The company is located in Wilmington, DE. Ask for Deborah Hunt.
-=W.L. Steffens=- Department of Veterinary Pathology College of Veterinary Medicine University of Georgia Athens, GA 30602 USA http://www.vet.uga.edu/vpp/wls/steffens.html
There are at least two problems involved in storing objects (specimens, pole pieces, specimen holders, and other things commonly stored in dessicators) in a plastic vacuum dessicator under vacuum: (1) the dessicator may have a small leak, and over time come back up to atmospheric pressure, thus exposing the objects to ordinary air, (2) under the influence of the vacuum plasticizers may bleed out of the plastic from which the desiccator is made and find their way onto the stored objects, as noted by others.
Both of these problems can be minimized quite simply, however, by filling the dessicator to atmospheric pressure with a dry, inert gas, rather than leaving it evacuated. For most purposes dry nitrogen would be a satisfactory gas to use here, although helium or argon might be preferred for storing some reactive materials. If the gas is purchased in a high pressure tank it is necessary to be sure that it is oil-free, otherwise the fill-gas may carry oil vapors onto the stuff you are storing inside the dessicator, thereby defeating the purpose of the whole operation, a matter discussed on p. 64 of my book 'Vacuum Methods in Electron Microscopy'.
The vapor pressure of water at the temperature of liquid nitrogen is in the realm of 10-20 Torr, and the vapor pressure of most oils is even lower, and so the gas that is constantly boiling off each container of liquid nitrogen is about as clean and dry as any you can get. As described on p. 65 of Vac Meth in EM, this dry nitrogen can be collected and used to fill vacuum apparatus rather simply. Fit a one-hole stopper into the LN2 storage flask and connect it to the inlet valve of the vacuum container with non-collapsable, flexible tubing (ordinary polyethylene tubing works well). Attach a large, collapsable plastic beach ball to a Tee joint in this tubing with a length of soft, surgical-rubber tubing, and make a clean slit in this surgical tubing about 100 mm long with a sharp razor blade or scalpel. Ordinarily this slit will close thghtly enough so that the nitrogen gas evolved from the storage container will be directed into the beach ball; however, if the ball becomes full the slit will serve as a primative pressure-release valve by opening slightly and allowing the gas to escape so that there is no danger of over-pressurization. When the inlet valve to the evacuated chamber is opened, the nitrogen will flow into the chamber only under the influence of atmospheric pressure acting on the collapsable beach ball, and so there is no danger of exceeding atmospheric pressure in the chamber. An ordinary beach ball will hold enough gas to fill most dessicators several times.
If an oil-sealed rotary vane pump is used to evacuate the storage chamber then some care must be exercised to avoid having oil vapours backstream from the pump into the chamber. To avoid this, it is important not to pump the chamber down below the range of viscous flow (i.e. below about 0.1 Torr - see p.29 of Vac Meth in EM). If this is not considered to be a sufficient vacuum to remove as much atmospheric gas as desired, then the container can be filled with the dry gas, pumped out and filled again a couple of times. Each time the container is pumped out some 90% of the existing gas molecules are removed, and so two or three flushings should leave only an insignificant trace of the original atmospheric gases.
Wilbur C. Bigelow, Prof. Emeritus Materials Sci. & Engr., University of Michigan Ann Arbor, MI 48109-2136 e-mail: bigelow-at-umich.edu; Fx:313-763-4788; Ph:313-764-3321
On Jul 1, 4:09pm, Christoph Guenther wrote: } We want to do immunohistochemistry on human heart muscle tissue cryosections. } Which embedding medium should be used for cryosectioning on a Leica cm } 3000 kryostat for light microscopy? } Are there simple recipies and procedures (cryoprotection...)? } Which freezing techniques should be used? We get the tissue in the } operating theatre which freezing technique can be used directly there? I } sthere a freezing technique for both, cryosectioning and RNA-preparation of } the same tissue-block.
In skeletal muscle, we embed in OCT compound (Miles, Inc.) and plunge freeze in melting isopentane. To avoid freeze artifact, go with smaller pieces of tissue ( { 20-30 mg). Also, keep the specimen in the melting isopentane only for 8-12 sec otherwise sample cracking occurs. For more detail, see the "Color Atlas of Muscle Histochemistry" by R.A. Brumback and R.W. Leech (PSG Publishing Co., 1984).
-- Gordon L. Warren, Ph.D. Research Scientist Muscle Biology Laboratory 158 Read Building Texas A&M University College Station, TX 77843-4243 Fax (409) 862-4808 Phone (409) 862-4809 office e-mail root-at-rangers.tamu.edu
He remarked that freeze-drying (for SEM) produced collapsed coatings and that AFM (imaging in air) produced the same result.
} We recommended the } customer a liquid Tapping Mode AFM (we are not eqiupped with it). -- I am happy to say that ASM is equipped to do exactly this. We have had good success imaging wet samples.
Don Chernoff Analytical Services Division
Advanced Surface Microscopy, Inc. E-Mail: asm-at-indy.net 6009 KNYGHTON RD. Voice: 317-251-1364 INDIANAPOLIS IN 46220 Toll free: 800-374-8557 (in USA) web: http://www.a1.com/asm Fax: 317-254-8690 (If you experience difficulty in accessing our website, note that the web address uses numeral "1" in "a1")
We use a 1:2 mixture of OCT with our cryoprotectant buffer (PO4 + 20% sucrose) freezing in isopentane cooled in liquid nitrogen. This gives us a block consistency that allows for 3 um cryosections at -20 C, see Barthel & Raymond, J. Histochem Cytochem, 1990, 38:1383-1388 or our web site for a detailed protocol, http://www.personal.umich.edu/~praymond/ Others have used this combination with good success for both thin and thick (+10 um) sections.
To avoid sample cracking we try to be sure that ratio of block size to tissue size is large. It seems our tissue swells somewhat when it freezes. If the block is not large enough then it cracks. Sometimes we avoid the cracking by "quick" dipping the sample in the cooled isopentane until it is completely frozen.
If anyone has other suggestions to avoid cracking that would be great. It is a continuous battle with our samples. Seems some days are worse than others.
For using the samples for RNA (I presume you mean in situs?) we treat the samples no differently except that our reagents and labware (including poly-L-lysine slides) are RNase free.
When collecting your sample from the OR is it necessary to directly freeze there? Your best preservation would be if you placed your sample in a bottle of fix and returned to the lab to continue the processing. Linda Barthel Research Associate II Department of Anatomy and Cell Biology University of Michigan lab (313) 764-7476 fax (313) 763-1166 barthel-at-umich.edu
We use a 1:2 mixture of OCT with our cryoprotectant buffer (PO4 + 20% sucrose) freezing in isopentane cooled in liquid nitrogen. This gives us a block consistency that allows for 3 um cryosections at -20 C, see Barthel & Raymond, J. Histochem Cytochem, 1990, 38:1383-1388 or our web site for a detailed protocol, http://www.personal.umich.edu/~praymond/ Others have used this combination with good success for both thin and thick (+10 um) sections.
To avoid sample cracking we try to be sure that ratio of block size to tissue size is large. It seems our tissue swells somewhat when it freezes. If the block is not large enough then it cracks. Sometimes we avoid the cracking by "quick" dipping the sample in the cooled isopentane until it is completely frozen.
If anyone has other suggestions to avoid cracking that would be great. It is a continuous battle with our samples. Seems some days are worse than others.
For using the samples for RNA (I presume you mean in situs?) we treat the samples no differently except that our reagents and labware (including poly-L-lysine slides) are RNase free.
When collecting your sample from the OR is it necessary to directly freeze there? Your best preservation would be if you placed your sample in a bottle of fix and returned to the lab to continue the processing. Linda Barthel Research Associate II Department of Anatomy and Cell Biology University of Michigan lab (313) 764-7476 fax (313) 763-1166 barthel-at-umich.edu
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Not just an idea, but two GREAT ideas! I knew that the free spirits on the listserver would think of good stuff. Thanks. CS } } What about a piece of black and and white photographic negative film for a } cheap test specimen? On a good microscope, the silver grains appear with } sharp edges. I don't have a toy microscope around to compare. But film is } widely available and cheap, dry, and you don't even need a microscope slide } or coverslip. No regular pattern to detect barrel or pincushion distortion, } but these probably wouldn't matter to most users anyway. } } BTW, someone around here made up a bunch of cheap stage micrometers by } photographing a metric ruler from a distance with a 35 mm camera. With the } reduction, the one mm divisions on the scale are 100 micrometers apart on } the negative. These were then just cut up and glued to microscope slides. } They are a little fuzzy but if you measure center to center of each line it } is close enough. Perhaps one could photograph a grid to make a test } specimen for distortion. } } Gary Radice, Associate Professor gradice-at-richmond.edu } Department of Biology 804-289-8107 (voice) } University of Richmond VA 23173 804-289-8233 (FAX)
Caroline Schooley Educational Outreach Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/
Yuhui wrote: ================================================== All of our scopes are JEOL made and we have been using filaments made by JEOL. The engineers from JEOL also recommend us to use their parts. As far as I know, however, there are companies which also sell filaments. Could anyone of you give me some information as to which source is better than the others? =================================================== While new filaments are always an option, have you considered retipping your left over filament bases? While not everyone would agree, there is a good number of persons who would claim that a good retipped filament is indistinguishable in its performance from a brand new one. Of course, not all retipped filaments from different sources are created equal. But if they work for you, a considerable amount of money can be saved.
Disclaimer: SPI Supplies offers a service to retip filaments as do others such as some of our competitors such as Pella, EBS, PLANO, etc. so we would all have a vested interest in seeing more people using retipped filaments instead of new ones. You can find out more information about the retipping of filaments from our website given below.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
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I have a sample that I have labeled with gold tagged antibody and I want to sputter coat it with aluminum for viewing in the SEM. We have a sputter coater with a turbomolecular pump and an aluminum target. We tired to sputter coat the sample using essentially the same "settings" used for chromium, but could not see that any coating had occurred. Does anyone have any ideas? Thanks in advance for any help any of you can give this humble student.
Dan - Never trust the low energy peaks from a specimen that is charging. Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
} Recently we did an SEM/EDX analysis that gave rather baffling results. } After some thought we came up with what seemed to be a plausible } explanation, however, the people for whom the analysis was done remained } skeptical. Therefore I am writing the Listserver for a second opinion. } The sample consisted of irregular shaped alumino-silicate particles roughly } 2-5 microns is size. The specimen was uncoated resulting in some charging. } Analysis (windowless) at 1000x mag. resulted in a carbon peak much larger } than the Al or Si peaks. At 5000x the carbon peak was smaller; i.e., } comparable to the Al and Si. Using a spot mode on individual particles, no } carbon was detected. However, spot mode analysis after lowering the } accelerating voltage from 20 to 10 kv, showed a small carbon peak. } The only explanation we could envision to fit these results was a thin } carbonaceous coating on the particle surface. Is this a plausible } explanation or we missing something? Does anyone have an alternative } suggestion? Any help would be appreciated. } } Dan Schwab } Center for Microscopy and Microanalysis } Bowling Green State University } Bowling Green, OH }
} How about an Environmental SEM? You could look at the sample in a fully } hydrated state, then slowly dehydrate, to reveal sub-surface structure. } } Regards, } Larry Stoter
We have successfully images hydrogel microspheres in the wet state using ESEM. See Applications of the environmental scanning electron microscope to the analysis of pharmaceutical formulations. D'Emanuele A, Gilpin C Scanning 1996 Oct 18:7 522-7
Chris
Chris Gilpin Biological Sciences Electron Microscope Unit G452 Stopford Building Oxford Road Manchester M13 9PT phone +44 161 275 5170 fax +44 161 275 5171
Is there a method of restricting a protozoa such as paramecium from moving about? The protozoa solution I have is great for studying them, but not if I want to look at the internal structure. Also what stain is a good stain to bring out the details in paramecium such as the nucleus?
Finally how do you pronounce 'paramecium'?!!!
Conrad :)
------------------------------------------------------------------------ ----------- "Any sufficiently advanced technology is indistinguishable from magic" ----------------------------------------------------------- Arthur C Clarke ----
I am working with the human cornea and wish to find the optimal buffers for glutaraldehyde and osmium tetroxide dilutions in the preparation of specimens for TEM. Any suggestions or references would be greatly appreciated.
} Cleaning grids? Go your hardest if you have nothing better to do: } Collect grids in a 20 or 30ml glass vial. Soak them in chloroform, give one } change in chloroform, pour off solvent, replace with ethanol or acetone,
Ethanol is better, if you store acetone and chloroform together in a waste bottle there is a risk, as those two will react explosively under some cirecumstances.
Also, after a final wash in aqueous media, if they are such as to remove the natural oxide coating, the copper underneath can oxidize and go all 'orrible.
} In a labour-costly country, with the odd exception, cleaning grids } is a waste of time.
I don't know if this applies to the USA, but in the UK we find that non-standard washing up is too difficult to be left to untrained staff!
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
Yuhui Xu asked: } All of our scopes are JEOL made and we have been using filaments made by } JEOL. The engineers from JEOL also recommend us to use their parts. As far as } I know, however, there are companies which also sell filaments. Could anyone } of you give me some information as to which source is better than the others?
We (Energy Beam Sciences) have been manufacturing filaments for electron microscopes for more than 25 years. I know that some third party filaments are equal in quality to those purchased from the original equipment manufacturer, and some are not. In shopping for filaments, it is important to ask whether the filaments are vacuum-annealed (to minimize drift when heated) and precentered *after* annealing.
It is also important to know that, although the original equipment manufacturers all offer only one style of filament loop on their filament bases, usually a "V" hairpin filament (eg Philips) or a "curved" loop (eg JEOL), other filament loops can be mounted on the same bases to maximize performance in one way or another. For example, a pointed filament can dramatically increase brightness, while another loop configuration, such as our trademark "AR" loop, will provide longer life.
Anyone interested in the details of these special loop configurations can contact me directly back-channel.
Disclaimer: Energy Beam Sciences manufactures a wide range of tungsten filaments for electron microscopes.
Best regards, Steven E. Slap, Vice-President ******************************** Energy Beam Sciences, Inc. Adding Brilliance To Your Vision ebs-at-ebsciences.com http://www.ebsciences.com/ ********************************
I have had some similar samples generate noise in the spectrum which formed a peak in the area of carbon's. Usually the low energy side did not return toward baseline which was caused me to investigate a bit more. I found that if I used my Ultra Thin (metal/polymer) window the unusual response disappeared. If carbon was really there, the peak would be only slightly attenuated, but would head back "down hill" on the low side. I did no more experiments to determine the cause, but it was suggested to me by several that the sample was likely generating photons (~visible spectrum light) which were hitting the crystal - causing detector noise...
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Hello everybody,
Recently we did an SEM/EDX analysis that gave rather baffling results. After some thought we came up with what seemed to be a plausible explanation, however, the people for whom the analysis was done remained skeptical. Therefore I am writing the Listserver for a second opinion. The sample consisted of irregular shaped alumino-silicate particles roughly 2-5 microns is size. The specimen was uncoated resulting in some charging. Analysis (windowless) at 1000x mag. resulted in a carbon peak much larger than the Al or Si peaks. At 5000x the carbon peak was smaller; i.e., comparable to the Al and Si. Using a spot mode on individual particles, no carbon was detected. However, spot mode analysis after lowering the accelerating voltage from 20 to 10 kv, showed a small carbon peak. The only explanation we could envision to fit these results was a thin carbonaceous coating on the particle surface. Is this a plausible explanation or we missing something? Does anyone have an alternative suggestion? Any help would be appreciated.
Dan Schwab Center for Microscopy and Microanalysis Bowling Green State University Bowling Green, OH
Thats ok then, as that was how I did tend to pronounce it, I just thought it may have been pronounced:
PARA-MEK-EE-UM!!
I shall make a note of the useful info on the dyes! I currently have the standard methyl blue.
Conrad
} -----Original Message----- } From: Lou Ann Miller [SMTP:lamiller-at-uiuc.edu] } Sent: Wednesday, July 02, 1997 1:57 PM } To: Conrad Perfett } Subject: Re: keeping a protozoa still } } Pronouncing paramecium.... the US way: } } } pair a me see um } } } } I'm not an expert in the field, but I can tell you that ~ 0.5%( grams per } 100ml) Tolluidine Blue is often used for fungi. } } Crystal Violet along with iodine is often used for bacteria that's been } dried on a slide. } } Might try iodine, that would be inexpensive to try, maybe even a generic } Betadine at the drugstore / pharmacist's. } Would need a more dilute solution than what form it comes in, so as to not } overpower your object. } } } Also, speaking as one that has done about 10,000 hospital urine analysis, } try lowering the condensor by quite a bit, and it makes some things more } clear at about 40KX objective. ie floating blood cells, mucus strands, } casts , and a little protozoan called trichamonous. ( forgive my spelling } , it's been a while, and we just called them trich.) Which you hope you } never ever have! } } Also, 2 pieces of Xray film crossing at right angles ( polorized material) } will make the background darker, and make things like uric acid crystals } stand out. } } } Keep having fun! } } } } Lou Ann } } } } Finally how do you pronounce 'paramecium'?!!! } } } } Conrad :) } } Lou Ann Miller } Center for Microscopy & Imaging } College of Veterinary Medicine } Dept. of Veterinary Biosciences } University of Illinois } Rm 1108 Basic Sciences Bld } 2001 S Lincoln Ave. } Urbana, Illinois 61802 } } Phone: 217-244-1566 } email: lamiller-at-uiuc.edu } } Center for Microscopy & Imaging Home page: } http://www.cvm.uiuc.edu/MicImagLab/MicImagLab.html } } Central States Microscopy Society: } http://www.cvm.uiuc.edu/Homepages/LouAnnMiller/CSMS/csms.html } } Personal Home Pages: } http://www.cvm.uiuc.edu/Homepages/LouAnnMiller/LAM.html } }
How were the samples mounted during analysis? When we do EDX at our lab, we generally use clean, pure carbon pin-type stubs and carbon adhesive tape or discs. At a mag of 1000x while viewing particles 2-5 microns in size, it seems that your probe would be scanning a large area of whatever they were mounted on. If this is the case, going to higher mag and/or doing spot analysis, thereby eliminating more of the background, would be likely to reduce or eliminate the x-rays from the background, and this is what you observed.
Also, unless I'm mistaken, lowering the accelerating voltage would tend to emphasize the peaks of lower atomic weight elements, relative to heavier ones, so if you were getting x-rays from your mounting materials, they might tend to show up more at lower kv's.
Finally, if you're using a variable pressure scope, lower vacuum in the column can scatter the electron beam to a large degree. Again, this would tend to cause x-ray emission from areas other than the particles you are interested in.
Hope I'm not belaboring the obvious, but it really sounds to me like the carbon is from your mount.
Randy Tindall Center for Electron Microscopy Southern Illinois University at Carbondale
Dear microscopists, We will soon be purchasing an inverted metallograph for specimen prep. It appears all the major brands are quite good, but perhaps there are subtle advantages / disadvantages that are only evident after much use and experience. If anyone has any strong recommendations, I would appreciate your input. Please feel free to contact me off-line if you prefer. Thanks in advance for your time and help.
Sincerely, Mick Thomas
Materials Science Center Cornell University Ithaca, NY 14853 mgt3-at-msc.cornell.edu
At 6:59 PM 7/1/97, Linda Barthel wrote: see } Barthel & Raymond, J. Histochem Cytochem, 1990, 38:1383-1388 or our web } site for a detailed protocol, http://www.personal.umich.edu/~praymond/ } Others have used this combination with good success for both thin and } thick (+10 um) sections. } I tried to look up your web site and got an error message to the effect that the web site could not be found. Is this the right address?
thanks
Rob Homer
Robert Homer, MD, PhD Asst Prof, Pathology Yale University School of Medicine 310 Cedar St. PO Box 208023 New Haven, CT 06520-8023
Just a comment on a comment maninly on semantics...
} the cause, but it was suggested to me by several that the sample was likely } generating photons (~visible spectrum light) which were hitting the crystal } causing detector noise...
Visible light does not create noise, it is signal. Remember the SiLi detector is an ENERGY DISPERSIVE SPECTROMETER. The photons are energy packets just like X-rays both of which are creating electron/hole pairs. The energy is just such that is at the very low energy range of the detector and hence in correctly gets called noise. It may be signal you do not want to see, but it is signal. Preferential attenuation of the light by a very thin visible light absorbing "window" is a pretty standard trick to FILTER OUT the light (unwanted signal) from the low energy x-rays desired signal.
I guess that is one way! I did think of that, but I guess I wanted to keep it alive so I can see the processes going on! Although it may seem obvious, what is the best way to kill them? putting in something like a few drops of an alcohol solution?
Conrad
} -----Original Message----- } From: Robert Wieland [SMTP:wieland-at-me.udel.edu] } Sent: Wednesday, July 02, 1997 1:37 PM } To: Conrad Perfett } Subject: Re: keeping a protozoa still } } On Wed, 2 Jul 1997, Conrad Perfett wrote: } } [lots cut] } } Is there a method of restricting a protozoa such as paramecium from } } moving about? The protozoa solution I have is great for studying them, } } but not if I want to look at the internal structure. Also what stain is } } a good stain to bring out the details in paramecium such as the nucleus? } } There are any number of sources of methyl cellulose, which makes the } "water" they live in more viscous so they don't dart out of the field. To } make them absolutely still, I believe it's best to kill them. } } } } } Finally how do you pronounce 'paramecium'?!!! } } } par-ah-ME-see-um } } } } } Conrad :) } } } } ------------------------------------------------------------------------ } } ----------- } } "Any sufficiently advanced technology is indistinguishable from magic" } } ----------------------------------------------------------- Arthur C } } Clarke ---- } } } } } } Robert Wieland wieland-at-me.udel.edu } You can't go faster than light, you can't get colder than absolute } zero, and you can't help somebody by not telling them the truth. } }
Sorry about the web site address, there is one minor error, the correct address is: http://www-personal.umich.edu/~praymond/
On Wed, 2 Jul 1997, R Homer wrote:
} At 6:59 PM 7/1/97, Linda Barthel wrote: } see } } Barthel & Raymond, J. Histochem Cytochem, 1990, 38:1383-1388 or our web } } site for a detailed protocol, http://www.personal.umich.edu/~praymond/ } } Others have used this combination with good success for both thin and } } thick (+10 um) sections. } } } I tried to look up your web site and got an error message to the effect } that the web site could not be found. Is this the right address? } } thanks } } Rob Homer } } Robert Homer, MD, PhD } Asst Prof, Pathology } Yale University School of Medicine } 310 Cedar St. } PO Box 208023 } New Haven, CT 06520-8023 } } }
You don't mention what you mounted your sample on. If it is carbon based (carbon stub, conductive carbon tape, other adhesive using cargon), you will see the carbon peak. Going to smaller areas of excitation (i.e. increasing the relative proportions of sample to background scanned) will cause the carbon peak to be reduced. At spot mode on a particle, you are pretty much just exciting the particle and not the background so you do not see the carbon. At the lower accelerating voltage the emission volume may broden out closer to the surface of the background giving you the small carbon peak. It is important to remember that just because the initial point of excitation is small, it doesn't mean that the volume of exitation is small. With your sample, you are probably exciting several cubic microns of your sample. A good monte carlo program will demonstrate this to you.
Hope this helps
Michael D. Standing Electron Microscopist Brigham Young University e-mail: MDStandi-at-bioag.byu.edu
} I have a sample that I have labeled with gold tagged antibody and I want to } sputter coat it with aluminum for viewing in the SEM. We have a sputter } coater with a turbomolecular pump and an aluminum target. We tired to } sputter coat the sample using essentially the same "settings" used for } chromium, but could not see that any coating had occurred. Does anyone } have any ideas? Thanks in advance for any help any of you can give this } humble student. } } VTY, } Beth Bray } bbray-at-netside.com
Is it possible to successfull sputter Al? Al oxidises rapidly, so I would have thought you'd have a nice coating of Al oxide, an excellent insulator, on your specimen.
} I have a sample that I have labeled with gold tagged antibody and I want to } sputter coat it with aluminum for viewing in the SEM. We have a sputter } coater with a turbomolecular pump and an aluminum target. We tired to } sputter coat the sample using essentially the same "settings" used for } chromium, but could not see that any coating had occurred. Does anyone } have any ideas? Thanks in advance for any help any of you can give this } humble student.
I assume that you are coating with the lower atomic weight Al versus Cr in order to image the gold using backscattered imaging in the SEM. Lower atomic numbered metals are difficult to sputter coat properly - although it can be done using more energy. Also, Al is very reactive and will oxidize almost immediately after coating. Carbon would provide a better sputter coat if you have the capability. Actually, thermal evaporation is the preferred technique with both Al and C.
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing Southern Illinois University Carbondale, IL 62901-4402 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
In 1992, I did some SEM studies on corneal endothelial cells from rabbits. I found that a 0.1M phosphate buffered saline worked fine for that purpose. Osmolarity was maintained between 300 - 310. This work was published in the EMSA proceedings for that year; "Evaluation of the Biocompatibility of Polymer Surface Modifications with the Corneal Endothelium", p. 1106.
Regards,
Bob *************************** Bob Citron Chiron Vision Claremont, CA USA (909)399-1311 Bob_Citron-at-cc.chiron.com ***************************
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I am working with the human cornea and wish to find the optimal buffers for glutaraldehyde and osmium tetroxide dilutions in the preparation of specimens for TEM. Any suggestions or references would be greatly appreciated.
Sorry for the incomplete info in my original posting. Several replies questioned the mounting method. The specimen was packed into small grooves in a silica-lead glass -- no carbon containing mountant of any kind was involved.
Thank you all for your expert advice and interest.
How about cubic zirconia for microscope objectives? RI = 2.15 - 2.45, dispersion = 0.060, birefringence = 0, hardness = 8.5 (quartz = 7), no cleavage with good toughness. Diamond is very hard, but has perfect cleavage in 4 directions (which means it chips relatively easily). So, CZ gets into diamond's range for RI, and should therefore make high NA lenses. CZ is a synthetic, so it can be grown under known conditions, and is relatively cheap and easy to produce. The high dispersion could be a problem for correcting chromatic aberration, but how much of one?
Phil
} Sic Hoc Legere Scis Nimium Eruditionis Habes { Philip Oshel Station A PO Box 5037 Champaign, IL 61825-5037 (217) 355-1143 oshel-at-ux1.cso.uiuc.edu *** looking for a job again ******************
Beth, Al can be vey differcult to sputter off with DC process since it builds up a very thick, closely bonded native oxide layer on its surface. You might want to manually abrade the target to remove the oxide layer, pump down and sputter immeadiately. John Arnott
True... I was thinking in terms of undesired signal as noise. "Undesirable signal" is certainly more accurate!
Speaking of undesirable signal:
Some of my specimens have generated dead times as high as 90+ percent - with the beam OFF! Spectra (w/ beam on) have very wide peaks and a relatively huge background. I know the reason... Highly radioactive specimens wreak havoc with my EDS. Noise - or - signal???? {g}
BTW, I am using an (old) Kevex "Extra" detector. The massive turret snout does provide a bit of shielding. Would like to add more, but have no chamber room left (Etec).
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Just a comment on a comment maninly on semantics...
} the cause, but it was suggested to me by several that the sample was likely
} generating photons (~visible spectrum light) which were hitting the crystal
} causing detector noise...
Visible light does not create noise, it is signal. Remember the SiLi detector is an ENERGY DISPERSIVE SPECTROMETER. The photons are energy packets just like X-rays both of which are creating electron/hole pairs. The energy is just such that is at the very low energy range of the detector and hence in correctly gets called noise. It may be signal you do not want to see, but it is signal. Preferential attenuation of the light by a very thin visible light absorbing "window" is a pretty standard trick to FILTER OUT the light (unwanted signal) from the low energy x-rays desired signal.
HEEEEELP! I am a graduate student trying to get through my LAST expt and it is not working. I am coating copper grids with collodion (2% in amyl acetate) and trying to analyze mitochondrial dna on them. Initially this worked beautifully. The last 200 grids I've done have been a bust. Problems 1)the collodion keeps frying under the beam, it often seems to "wobble" under the beam. I have tried 3 different bottles of new collodion, cleaned the grids well with detergent-water-then acetone rinses all to no avail.
2)the dna seems to have stopped adhering to the collodion. I first heat nick the dna so it will be open circular (80 degrees, 30 min, in 0.5M NHAc) then I add cytochrome c to bind to the dna (I've tried concentrations from 2-100ug/ml) I then rotary shadow with platinum.
I do polarized light chemical microscopy in the style (or school) of Walter McCrone as part of doing general analytical chemistry for a pharmaceutical and chemical manufacturer. Examples of the types of problems I deal with are relating particle morphology of pure substances to properties and some rudimentary optical crystallography to distinguish crystal types. Can anyone provide experience or references on the possible usefulness of confocal or other "modern" light microscopic techniques in this area? Are there good journals covering such topics as well as classical PLM?
Dr. Leonard R. Corwin Principal Research Chemist Fort Dodge Animal Health Cyanamid Agricultural Research Center Quaker Bridge & Clarksville Roads PO Box 400 Princeton, NJ 08543-0400 609-716-2278 609-275-5239 fax corwinl-at-pt.cyanamid.com
Does anyone know where we can get our LKB 7800B knife breaker serviced? If possible, by someone in the Chicago area? It is making sporadically bad knives and we have adjusted all the knobs by the instruction booklet. If it needs to be replaced, can anyone recommend a good one? This one has been with us for over 20 years.
Thanks, Linda Fox, Loyola Univ. Medical Center, Chicago lfox1-at-wpo.it.luc.edu
To: Didier Le Thiec I.N.R.A. Centre de Recherches Forestieres Unite d'Ecophysiologie Forestiere Laboratoire de Pollution Atmospherique 54280 Champenoux - France
In response to request for information about WDX system with SEM or LVSEM.
Does anyone have a good SEM fixation method for planaria?
Nancy R. Smith Director of Operations Microscope And Graphic Imaging Center California State University, Hayward nsmith-at-csuhayward.edu http://www.csuhayward.edu/SCI/sem
} ... } } } I have a sample that I have labeled with gold tagged antibody and I } want to } } sputter coat it with aluminum ... } } ... } can be done using more energy. Also, Al is very reactive and will } oxidize } almost immediately after coating. ...
An Al oxidation layer tends to be very thin and impervious to further oxygen ... I might believe that under a vacuum the oxidization wouldn't occur until exposed to air, and would still be conductive (??)
cheerios, shAf -- {\/} /\ {\/} /\ {\/} /\ {\/} cogito, ergo zZOooOM {\/} /\ {\/} /\ {\/} /\ {\/} Michael Shaffer, R.A. - University of Oregon Electron Probe Facility mshaf-at-oregon.uoregon.edu -or- mshaf-at-darkwing.uoregon.edu http://darkwing.uoregon.edu/~mshaf/
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No, it's MEESE. And for a stain, go to a tropical fish store and get "ich" medecine. It's an aqueous copper salt & it works fine (see "Fun at 40 Power" in the MMMMMMProject MICRO bibliography).. Caroline
Caroline Schooley Educational Outreach Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/
Try "Protoslo" quieting solution from Carolina Science and Math, Phone number: 1-800-334-5551. You might consider requesting a catalog. I believe that you might find they sell a number of interesting living and dead organisms for observation by light microscopy.
Regards,
Kirk J. Czymmek, Ph.D. Biological Electron Microscopy Facility 111 Wolf Hall University of Delaware Newark, DE 19716
Dear microscopists, We will soon be purchasing an inverted metallograph for specimen prep. It appears all the major brands are quite good, but perhaps there are subtle advantages / disadvantages that are only evident after much use and experience. If anyone has any strong recommendations, I would appreciate your input. Please feel free to contact me off-line if you prefer. Thanks in advance for your time and help.
Sincerely, Mick Thomas
Materials Science Center Cornell University Ithaca, NY 14853 mgt3-at-msc.cornell.edu
Dear Mick:
What would you be using this inverted metallographic microscope for? If you would use it for mostly tripod-type TEM sample preparation, I would highly recommend Lieca's model. It has a nice design and has higher resolution than other models that we've tested. If it is for general all purpose sample preparation and inspection, others would be more qualified to answer your question.
Regards,
Michael Coviello EM Lab Manager The University of Texas -at- Arlington Arlington, TX 817-272-5496
A week before our service contract for our Hitachi-7000 was to be renewed by Thomas Technical we were informed that they did not have the staffing for our TEM. We currently are totally without a contract. We cannot afford Hitachi's offer for the limited service we need. Can anyone recommend an independent company which we might contact? Any ideas would be appreciated. Thanks, Hildy Crowley Dept of Biol Sciences University of Denver 2101 E Wesley Ave Denver, CO 80208 Tel. 303-871-3026 E-mail { hcrowley-at-DU.edu } FAX 303-871-3471
The classic way to quite protozoa was a dilute solution of cocaine HCl. (Try explaining that to your local DEA officer) A solution of methyl cellulose will increase the viscosity of the mount to slow activity down for a good look. Years ago I tried Novocaine, which I obtained from my dentist, with limited success.
Best wishes...Frank ---------------------------------------------------------------------------- -------
These opinions are mine alone and have no relationship to my employer. Thank you.
Frank Karl fskarl-at-goodyear.com Goodyear Tire & Rubber Co. Voice: 330-796-7818 Analytical Services Dept. 415B Fax: 330-796-3304 142 Goodyear Blvd. Akron, OH 44305 USA
They that give up essential liberty to obtain a little temporary safety deserve neither liberty or safety. Benjamin Franklin
Is there a method of restricting a protozoa such as paramecium from moving about? The protozoa solution I have is great for studying them, but not if I want to look at the internal structure. Also what stain is a good stain to bring out the details in paramecium such as the nucleus?
Finally how do you pronounce 'paramecium'?!!!
We are teaching a summer course to high school kids using protozoans as our subject. They will be doing a lot of staining to show various internal structures, and a list of these stains is available on the "under construction" website for the course (http://131.229.114.77/LAL) - look under the section on examining protozoa. Most of the solutions are about .01-.05% aqueous. Takes some fussing around to get it right, as different types of protozoans vary a lot. There is a great text called "Protocols in Protozoology" edited by Lee and Soldo and published by the Society of Protozoologists. It has a lot of info on slowing 'em down - ranging from compression under the coverslip to various exotic techniques. Methylcellulose (10% aqueous, heating (don't boil it) to dissolve) is good. I have used CO2 (club soda water) added to the drop, as well as .01% nickel chloride for ciliates. Also try MgCl2 and KCl. Good luck. Dick Briggs
To: Didier Le Thiec I.N.R.A. Centre de Recherches Forestieres Unite d'Ecophysiologie Forestiere Laboratoire de Pollution Atmospherique 54280 Champenoux - France
In response to request for information about WDX system with SEM or LVSEM.
WDX requires a large probe current which means a large spot size and inferior resolution. Depending upon the specimen, the large beam current can produce specimen damage - not a problem for most bulky physical specimens but probably a problem for organic specimens. Cold stages can reduce damage for some specimens, but because damage is radiation damage breaking down molecules anot heat damage, the reduction in damage is not universal.
Operating in the Low Vacuum of Variable Pressure mode does not do anything to prevent or enhance the beam damage. the residual gas molecules will scatter the beam, resultiong in x-rays being generated at positions away from the major beam incidence region. For this reason, care should be exercised in interpreting the results obtained from x-ray studies in the variable Pressure Sem mode. Shorter working distances minimize this effect. I hope shortly to report on another technique for further minimizing this effect.
I can't give you references on WDX in variable pressure SEM, but following are some references for the early work on the development of the variable pressure SEM capability.
References on Variable Pressure SEM
V.N.E. Robinson, A Wet Stage Modification to a Scanning Electron Microscope - Proceedings of the Eighth International Conference on Electron Microscopy, Ed. J.V. Sanders and D.J. Goodchild, Australian Academy of Science, Canberra, Vol. II, pp 50-51.
V.N.E. Robinson, A Wet Stage Modification to a Scanning Electron Microscope, J. Microscopy, 103, pp 71-77, 1975.
V.N.E. Robinson, The Elimination of charging artifacts in the Scanning Electron Microscope, J. Phys. E: Sci. Instrum. 8, pp 638-640, 1975.
V.N.E. Robinson, Scanning Electron Microscope Environmental Cells, Scanning Electron Microscopy 1976/I, Ed. O Johari and I Corvin, IITRI, Chicago pp 91-100.
V.N.E. Robinson, The Examination of Hydrated Biological Specimens in a Scanning Electron Microscope Environmental Cell, Electron Microscopy, 1976, Proc. 6th European Congress, Ed. Y. Ben-Shaul, Tal International, Jerusalem. Vol. II, pp 85-90.
D.A. Moncrieff, V.N.E. Robinson and L.B. Harris, Charge Neutralisation of Insulating Surfaces in the SEM by Gas Ionisation, J. Phys. D: Appl. Phys. Vol 11, pp 2315-2325, 1978.
D.A. Moncrieff, P.R. Barker and V.N.E. Robinson, Electron Scattering by Gas in the Scanning Electron Microscope, J. Phys. D: Appl. Phys. Vol. 12, pp 481-487, 1979.
G.D. Danilatos and V.N.E. Robinson, Principles of Scanning Electron Microscopy at High Specimen Chamber Pressures, Scanning, Vol 2, pp 72-82, 1979.
V.N.E. Robinson, the Examination of Hydrated Specimens in electron Microscopes, in Analysis of Organic Biological Surfaces, John Wiley and Sons, New York, 1984, Ed. P. Echlin, Ch. 8, pp 191-207.
Tom Ruscica ETP USA, Electron Detectors Inc. For V.N.E. Robinson
Thanks to all of you who sent replies in answer to my question about sputter coating with aluminum. The consensus appears to be that, in general, "you can't get there from here." Aluminum was chosen because, as John Bozzola correctly assumed, I want to image the gold using backscattered imaging in the SEM. I should have included that little fact in my initial query. My next move will be to check into using Zn, Cr, Cu, Ag, or Pd--which was another suggestion, and to research some of the suggested articles, as well as looking at using thermal evaporation. There are so many things to learn--whew!--but I am having more fun than I would ever have thought possible. I am a "mature student" who is fortunate enough to work for a company that is paying my way back to college to finish a degree that I started many years ago. I will graduate in December of this year (God willing and the creek don't rise) and I am having a blast in school! Any further ideas will very much welcomed. Again, many thanks!
Dear microscopists, We will soon be purchasing an inverted metallograph for specimen prep. It appears all the major brands are quite good, but perhaps there are subtle advantages / disadvantages that are only evident after much use and experience. If anyone has any strong recommendations, I would appreciate your input. Please feel free to contact me off-line if you prefer. Thanks in advance for your time and help.
Sincerely, Mick Thomas
Materials Science Center Cornell University Ithaca, NY 14853 mgt3-at-msc.cornell.edu
REPLY:
Dear Mick:
What would you be using this inverted metallographic microscope for? If you would use it for mostly tripod-type TEM sample preparation, I would highly recommend Lieca's model. It has a nice design and has higher resolution than other models that we've tested. If it is for general all purpose sample preparation and inspection, others would be more qualified to answer your question.
Regards,
Michael Coviello EM Lab Manager The University of Texas -at- Arlington Arlington, TX 817-272-5496
} The classic way to quite protozoa was a dilute solution of cocaine HCl.
As was mentioned use methyl cellulose. It is sold under the name Protoslo. Another trick is to chill the Protozoa medium and to use neutral density filters to keep the heat of the lamp off of the little buggers.
Blystone in Texas
Robert V. Blystone, Ph.D. {RBLYSTON-at-Trinity.edu} Professor of Biology Trinity University San Antonio, Texas 78212 210.736-7243 210.736-7229 FAX
Sorry to be so long in geting to this. In the rat race of life the rats are winning. Now to split a few hairs with you Mark, and hopefully we will not bore the list to death or scare away those amateurs. On Tue, 1 Jul 1997, Dr. Mark W. Lund wrote:
} } Lenses have several types of aberrations which will cause the loss of } } detail unless corrected for. Toy microscopes are not. } } } } Spherical aberration - "is especially apparent in lenses having sperical } } surfaces. Light paths near the center of the lens focus at different } } points compared to light paths near the periphery." } } } The name "spherical aberration" has historical roots in astronomy where } a spherical mirror has this aberration but a parabolic mirror does not. } In some instances a spherical mirror will have no spherical aberration } wheras a parabolic mirror will have maximum spherical aberration. } The aberration really has nothing to do with the spherical surfaces } of lenses. The confusion comes because making a surface "aspheric" } can cure spherical aberration in many instances.
What confusion? Spherical aberation (also sometimes called aperture aberation) is a recognized term. While the refraction of light remains constant, the "distance traveled" in the lens and the angles of the lens to the object change as a function of the curvature of the lens surface. That curvature resembles the surface of a sphere.
} } } } Field curvature - "is a natural result of using lenses with curved } } surfaces. The image plane produced by such lenses will be curved. This } } kind of image occurs in microscopy unless plano (flat-field) objectives } } are used." } } Actually, field curvatureis a natural result of the geometry of the } real world. Since an object at the edge of the field of view is } farther from the lens "center" it will tend to be focussed closer to the lens } than an object on axis. This naturally leads to field curvature unless } the designer makes the lens weaker for off axis points. It has nothing } to do with the lenses being curved. The confusion comes from the formula } for Petzval curvature, which has lens powers in it.
And the geometry you refer to is due to what? I would suggest that it is due to the curved surface/s of the lens. The curvature of the image does mimic the curvature of the lens.
As to "off axis points" we are not getting into Coma aberration are we?
To those one the list, hopefully this friendly discussion/hair spliting session does not bore you all to tears.
as an alternative to having your knife-maker serviced have you considered replacing the scoring tool part. I am familiar with the LKB 7801A and it's a simple task to replace the cutting wheel (it should be in the instructions). If you haven't got any spares then you may have to trace a supplier but a pack of several replacements would cost you less than the call-out for a service. My experience with these knife makers is that they virtually go on for ever.
Sorry if you've already done this.
Malcolm Haswell e.m. unit University of Sunderland UK ----------
Beth:
An additional thought, why not coat with a thin layer of evaporated carbon. You would only need to put on a few nm.
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Thanks to all of you who sent replies in answer to my question about } sputter coating with aluminum. The consensus appears to be that, in } general, "you can't get there from here." } Aluminum was chosen because, as John Bozzola correctly assumed, I want } to image the gold using backscattered imaging in the SEM. I should have } included that little fact in my initial query. My next move will be to } check into using Zn, Cr, Cu, Ag, or Pd--which was another suggestion, and } to research some of the suggested articles, as well as looking at using } thermal evaporation. } There are so many things to learn--whew!--but I am having more fun than } I would ever have thought possible. I am a "mature student" who is } fortunate enough to work for a company that is paying my way back to } college to finish a degree that I started many years ago. I will graduate } in December of this year (God willing and the creek don't rise) and I am } having a blast in school! } Any further ideas will very much welcomed. Again, many thanks! } } Beth Bray } bbray-at-netside.com } } }
As one fellow amateur to another, I'd be interested in the type of camera you are intending to use! I currently use a Praktica MTL50 but am thinking of getting an OM1 or 2. I have looked around and even secondhand they are still expensive! but it seems Olympus are the only ones that have the optionable clear screen (which seems to cost anywhere from 30 to 40 UK pounds for that alone!). I was thinking of following someones advice on here (from Australia if I remember correctly) and cementing a square coverslip with canada balsam to the focusing screen of another cheap camera, probably another Praktica as they are dirt cheap here secondhand. Anyone else tried this? Is it satisfactory? I realise it would make the camera only suitable for microphotography but that does not matter as it would be vastly cheaper.
Conrad
} -----Original Message----- } From: Frank J. Hogan [SMTP:jhogan-at-freenet.npiec.on.ca] } Sent: Sunday, June 29, 1997 4:16 AM } To: Conrad Perfett } Subject: RE: Amateur Microscopy } } Hi again Conrad, } Been away for a bit and just got caught up with the traffic on the } mic list....wow! you really started something good. It really is time } somebody looked into the toy microscope scene or should I say scam. I had } one when I was a little guy and it really put me off scoping until I got } my Bushnell lab model for med school. } I didn't graduate from med school but switched to journalism where I } earned my bread until recent early retirement. Fortunetely I kept my scope } after med school days. Now, I hope to make use of it as a hobby tool. } My scope has all the bells and whistles a lab scope should have I } guess. I am limited by a three objective turret, 4x,10x,40x and an 8x and } 10x eyepieces. I am equiped to do photomicroscopy and it is something I'm } looking forward too. } From the sound of things all you folks on the list are way ahead of me } equipment-wise, but I'm content to putt along with what i've got for now. } It also sounds like the more advanced people are providing all the info } you can handle. } } Very best } Frank }
I am a non-smoker! Also I *do* already have a protozoa solution to slow the critters down, but was looking more for a way to keep them still so I can look and photograph them
Thanks for the replies I now have a wealth of information!
Conrad :)
} -----Original Message----- } From: Keith Ryan [SMTP:KPR-at-wpo.nerc.ac.uk] } Sent: Thursday, July 03, 1997 9:07 AM } To: fskarl-at-goodyear.com; microscopy-at-Sparc5.Microscopy.Com } Subject: RE: keeping a protozoa still -Reply } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Conrad, You could try putting the little critters in the fridge for a while before you look at them. This works for fruit flies and slows them down, so it may be worth a try. Good luck Nikki
******************************** Nikki Bock EM Technician Dept.ME&MD University Of Nottingham Nottingham NG7 2RD Tel: (0115) 9513759 Email: emznjb-at-emn1.nott.ac.uk
I am re-posting my question, since I have not received any replies and our further trials are still unsuccessfull. Maybe this time, with a different Subject, somebody will notice and help us. My appologies to those not interested in immuno.
Dear immuno colleagues,
We would like to label F-actin at the EM level in growth plate chondrocytes and matrix vesicles. We have done some pilot experiments at the light microscope level using:
1) Rabbit anti-actin antibody (A 2066 from Sigma) on paraffin sections. The label was confined only to a subset of smooth muscle cells in control tissues.
2) Phalloidin-BODIPY FL worked very nicely on cryostat sections, therefore we tried anti-BODIPY antibody, followed by biotinylated goat-anti-rabbit, Streptavidin peroxidase and AEC substrate. However, we got no labelling.
We would appreciate any input on F-actin labelling at EM level in non-muscle cells or any experience dealing with rabbit anti-BODIPY FL antibody.
Thank you,
Sarka Lhotak EM Facility, McMaster University Hamilton, Ontario, Canada
I am re-posting my question, since I have not received any replies and our further trials are still unsuccessfull. Maybe this time, with a different Subject, somebody will notice and help us. My appologies to those not interested in immuno.
Dear immuno colleagues,
We would like to label F-actin at the EM level in growth plate chondrocytes and matrix vesicles. We have done some pilot experiments at the light microscope level using:
1) Rabbit anti-actin antibody (A 2066 from Sigma) on paraffin sections. The label was confined only to a subset of smooth muscle cells in control tissues.
2) Phalloidin-BODIPY FL worked very nicely on cryostat sections, therefore we tried anti-BODIPY antibody, followed by biotinylated goat-anti-rabbit, Streptavidin peroxidase and AEC substrate. However, we got no labelling.
We would appreciate any input on F-actin labelling at EM level in non-muscle cells or any experience dealing with rabbit anti-BODIPY FL antibody.
Thank you,
Sarka Lhotak EM Facility, McMaster University Hamilton, Ontario, Canada
I believe that Leica is now handeling (or has acquired) the LKB knife breaker. You may try to contact your local Leica sales rep. for more information.
Disclaimer: I have no personal or financial interest in Leica
Michael Standing Electron Microscopist Brigham Young University e-mail: MDStandi-at-bioag.byu.edu
---------- } From: NANCY SMITH {nsmith-at-gauss.sci.csuhayward.edu} } To: microscopy-at-Sparc5.Microscopy.Com } Subject: Planaria } Date: Mon, 30 Jun 1997 15:03:41 PSD8PDT } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I think this one works ok, it has been some years since I used it, but it was fine then! 200ml dist. H2O, 2 ml conc. HNO3, 4.5 ml formalin and 2.5 g MgSO4. Fix by dropping solution onto the planaria, done at room temp, a higher temp causes improper relaxation, a lower one, mucus secretion. Fixation complete within 24hrs, the specimens may be stored in 70% ethanol, or 5% formalin. Osmicate specimens and dehydrate as per usual. Good luck!
Alex Black Department of Anatomy University College Galway IRELAND
Further to my earlier note about tobacco smoke, I now have the reference: CFA OPantin. Notes on Microscopical Technique for Zoologists. Cambridge University Press. 1969., page 7.
Funnily enough, guess what it is recommended for - Paramecium !! Also flagellates, the cilia of Mytilus and Hydra. Not very politically correct (but in certain matters, I may not be!). Expedient, if not finally terminal for both the Paramecium and the microscopist!!
Also in this small book (of 76 pages) are recommended: 1. 10% alcohol - if you are an amateur, maybe whisky etc could be added to your water. 2. magnesium chloride - 7.5% MgCl2.6H2O or 20% MgCl2.7H20 are both isotonic with seawater, use half seawawater and half MgCl2 solution. This works atraet within minutes with baby cuttlefish and squid (in fact, we got a paper about it!)
Use 2.5% Mgcl2.6H2O fo freshwater organisms, it is slow e.g. 2 hours for Planaria.
3. Menthol. - scatter a few crystals on the surface of the water and leave overnight. Our specimen dept. used to do this with marine invertebrates.
4. Ether vapour - good luck (explosively flammable!)
A more modern item is polyethylene oxide with a Molecular Weight of 4,000,000. From the Aldrich catalogue, cat. no. 18,964-4, 5 grams costs #12.20 in UK before 17.5% tax. Used at 1% in your medium, this works very well. This is a tip from the person for whom I recently posted a fluorescence microscopy question.
Best wishes - Keith Ryan Plymouth Marine Lab., UK (Bonjour, Daniele!)
Does anyone know where we can get hydrophilic glass coverslips, please? We have seen a reference to them in a German paper, as supplied by ILMGLAS, but we cannot find this company on the web. Any other suppliers?
Thanks in advance,
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
POSITION AVAILABLE: Laboratory Manager, Interdepartmental Laboratories
The American Museum of Natural History is seeking a Laboratory Manager to manage the Museum's Interdepartmental Laboratories. Duties include maintenance and operation of analytical microscopy equipment, particularly a scanning electron microscope with energy dispersive spectrometer, confocal laser scanning microscope, and digitial imaging and printing facilities; day to day management of laboratory operations; training users on analytical instrumentation; and participation in biological, geological and anthropological research. Qualifications include an advanced degree in Museum science-related field, 5 years experience in analytical laboratory operation and with scanning electron microscopes and energy dispersive systems. Ability to work with research scientists and to manage day-to-day operation of a digital imaging facility also required Background in confocal laser scanning electron microscopy desirable. Full Benefits. Please send resume with salary history to:
Human Resources American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192.
You may also e-mail your application to: barnett-at-amnh.org
An Equal Opportunity Employer. No phone calls or faxes, please.
-------------------------------------------------------------------------- | William K. Barnett, Ph.D. | | Director, Interdepartmental Laboratories barnett-at-amnh.org | | American Museum of Natural History | | Central Park West at 79th Street | | New York, NY 10024-5192 | --------------------------------------------------------------------------
The recent discussion about inexpensive, high quality microscopes, reminded me that I have a couple of excess microscopes that I would be happy to sell. These were my primary instruments until I recently moved up to Zeiss. They are older, Bausch and Lomb/American Optical Spencer microscopes with illuminators, excellent objectives, lenses, and condensors. They are in excellent condition with available lenses ranging from 3.5x to 100x and include oil immersion. If I remember right, eyepieces are either 6x or 10x. These are very good for amateur, exploration, etc. at a low cost. No plastic lenses or cheap parts.
Price: 150 U.S. to $250 U.S. (depending on what you want in the say of objectives and lenses) plus shipping.
I won't be able to answer you for a few days, but let me know what you think. I'll get back to you shortly.
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The successful candidate will join a small team of experts involved in the structural examination of catalysts, polymers, biomaterials, and other nanophase materials. The position requires an M.S. or B.S. or equivalent in material science or a related field with 3+ years experience in transmission electron microscopy (TEM) associated with materials. A strong background in TEM and associated preparative techniques, including embedding, ultramicrotomy, and dark room procedures is essential.
The successful candidate will be highly motivated, self-directed, interested in learning new skills, and effective working independently or in a team-based research environment. Excellent interpersonal, verbal and written communication skills are necessary. Experience with SEM and other imaging techniques, as well as digital imaging and image analysis are desired but not required.
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} Aluminum was chosen because, as John Bozzola correctly assumed, I want } to image the gold using backscattered imaging in the SEM. I should have } included that little fact in my initial query. My next move will be to } check into using Zn, Cr, Cu, Ag, or Pd--which was another suggestion, and } to research some of the suggested articles, as well as looking at using } thermal evaporation.
Be careful in chosing the proper metal to coat your specimen because if you use a metal with too high atomic number (Pd, Ag, Au, Cr, etc) the backscatter emissions may mask completely the gold emission from the specimen - especially since it will be weak to begin with. In this case, thermally evaporated carbon is the coating of choice.
Isn't microscopy fun! Good luck in your career |8 { ))
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing Southern Illinois University Carbondale, IL 62901-4402 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
I have been asked to study copper cathode microstructure with LM. Can anyone direct me to any descriptions, micrographs or papers written about cathode copper microstructure. Addresses of Companies and/or organizations to contact with for help on this subject would also be very helpful to me.
} HEEEEELP! I am a graduate student trying to get through my LAST } expt and it is not working. I am coating copper grids with } collodion (2% in amyl acetate) and trying to analyze mitochondrial } dna on them. Initially this worked beautifully. The last 200 } grids I've done have been a bust. Problems } 1)the collodion keeps frying under the beam, it often seems to } "wobble" under the beam. I have tried 3 different bottles of new } collodion, cleaned the grids well with detergent-water-then acetone } rinses all to no avail.
Check that you are not over-irradiating the specimen in the TEM (emission too high, too large condenser spot size or aperture, too low kV). OR you may have overloaded the grid with specimen material.
} 2)the dna seems to have stopped adhering to the collodion. I } first heat nick the dna so it will be open circular (80 degrees, 30 } min, in 0.5M NHAc) then I add cytochrome c to bind to the dna (I've } tried concentrations from 2-100ug/ml) I then rotary shadow with } platinum.
The cytochrome c may have gone bad. This happened to me once and the dna was not coated properly. Same batch used as previously?
Is the rotary shadowing being done properly - i.e., adequate Pt being deposited? If you put down enough Pt, this should stabilize the collodion as well. Not enough Pt will lead to lack of contrast (no dna seen) as well as drifting/unstable films.
Did you try checking out plain collodion coated grids, minus specimen? If bad, try a different vendor.
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing Southern Illinois University Carbondale, IL 62901-4402 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
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An aqueous extract of tobacco leaf usually has enough nicotine to make them comatose. And if you're a classroom volunteer, it's a dramatic & definitely PC demonstration for the young folk... Caroline
Caroline Schooley Educational Outreach Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/
We have a similar setup though we use our TN 5500/5600 on a microprobe. There is a flex module that enable you to choose to send data out one of the four ports on the back of your 5500. It is the +} DV command. +} DV 0 is the default which sends data out the printer port. We send data to +} DV 4 and print out formated information to a PC via a 9-pin cable and into comm port 1 or 2. This is 1200 baud communications so images take a long time. On the PC side, we have basic programs that capture what is comming in. There are programs for both data string (analyses) and image transfer. All are very limited but being able to transfer data out of the PDP is an absolute nec.
E-mail if you want more info.
Ciao for now, Ken
Kenneth JT Livi Department of Earth and Planetary Sciences 34th and Charles Streets Johns Hopkins University Baltimore, Maryland 21218 USA Phone: (410) 516-8342 Fax: (410) 516-7933 e-mail: klivi-at-jhu.edu
Hi! Just a note on tobacco juice. I was told that stuffing tobacco leaves on your socks will prevent leaches from crawling up your legs as you cross rivers. Any scientific truth in it? Anything to do with the nicotine?
K.M. Khoo
On Thu, 3 Jul 1997, Caroline Schooley wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } -----------------------------------------------------------------------. } } } } Further to my earlier note about tobacco smoke, I now have the } } reference: CFA OPantin. Notes on Microscopical Technique for } } Zoologists. Cambridge University Press. 1969., page 7. } } } } Funnily enough, guess what it is recommended for - Paramecium !! Also } } flagellates, the cilia of Mytilus and Hydra. Not very politically correct (but } } in certain matters, I may not be!). Expedient, if not finally terminal for } } both } } the Paramecium and the microscopist!! } } An aqueous extract of tobacco leaf usually has enough nicotine to make them } comatose. And if you're a classroom volunteer, it's a dramatic & } definitely PC demonstration for the young folk... } Caroline } } } } Caroline Schooley } Educational Outreach Coordinator } Microscopy Society of America } Box 117, 45301 Caspar Point Road } Caspar, CA 95420 } Phone/FAX (707)964-9460 } Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html } Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/ } } }
I travel the world teaching practical electron microscopy and it worries = me the emphasis that people place on "Spot Mode". I do not teach the use of=
spot mode for the following reasons-
1. Just because the spot appears in a certain position on the screen=
is this its correct position. I have found machines many microns out of step in X and Y directions.
2. Specimens charge and switching out of spot mode does not give the=
operator an easy opportunity to see if the spot had moved during the analysis.
3. Spot mode gives a false sense of analytical volume, no matter how=
small that spot may be we are almost certainly evaluating microns of material.
Do others worry about spot mode accuracy, do others test the spot mode accuracy? Is it not better to simply increase the magnification watching=
the area of interest all the way up before the analysis and all the way down after the analysis?
I have made an experiment with mites and than an embedding with Spurr=B4s epoxy resin. After polymerisation the blocks are fragile and shatter. Is there a possiblity to resolve Spurr=B4s and than to repeat the embedding of the samples or other solutions?
Thanks in advance
Heike Buecking Dr. Heike Buecking University of Bremen UFT Physiological Plant Anatomy Leobener Str. D 28359 Bremen Germany TEL: +49-421-218-2954 or TEL: +49-421-218-7283 FAX: +49-421-218-3737 e-mail: heibueck-at-uft.uni-bremen.de
unsubscribe =================================================================Dr. Francois Goutenoire Laboratoire des Fluorures - UPRES A 6010 Faculte des Sciences-Universite du Maine Avenuue O. Messian, BP 535 Fax:(33).2.43.83.35.06 72000 Le Mans Cedex, France Tel:(33).2.43.83.33.53 =================================================================
} Date: Fri, 04 Jul 1997 09:41:38 +0200 } To: microscopy-at-msa.microscopy.com } From: Heike Buecking {heibueck-at-uft.uni-bremen.de} } } Dear all, } } I have made an experiment with mites and than an embedding with Spurr=B4s epoxy resin. After polymerisation the blocks are fragile and shatter. Is there a possiblity to resolve Spurr=B4s and than to repeat the embedding of the samples or other solutions? } } Thanks in advance } } Heike Buecking Dr. Heike Buecking University of Bremen UFT Physiological Plant Anatomy Leobener Str. D 28359 Bremen Germany TEL: +49-421-218-2954 or TEL: +49-421-218-7283 FAX: +49-421-218-3737 e-mail: heibueck-at-uft.uni-bremen.de
I would agree with Stephen that it is important to emphasise the possible inaccuracies with using spot mode. However I would not go as far as saying don't use spot mode.
Spot mode can be useful for both quick looks to check compositions of a variety of particles in a field of view as well as for accurate microanalysis. Obviously with the latter it is important to be aware of the possibility of drift and to be sure of the position of the spot. Certainly with thin sections it is often possible to see the burn marks where the beam hit the sample. I tend to emphasise to students that when using spot mode the analysis will be a volume of 1-2 microns diameter and depth depending on sample and conditions. Even when using general rastering the same problem will be there.
This is just my opinion.
} I travel the world teaching practical electron microscopy and it worries me } the emphasis that people place on "Spot Mode". I do not teach the use of } spot mode for the following reasons- } } 1. Just because the spot appears in a certain position on the screen } is this its correct position. I have found machines many microns out of } step in X and Y directions. } } 2. Specimens charge and switching out of spot mode does not give the } operator an easy opportunity to see if the spot had moved during the } analysis. } } 3. Spot mode gives a false sense of analytical volume, no matter how } small that spot may be we are almost certainly evaluating microns of } material. } } Do others worry about spot mode accuracy, do others test the spot mode } accuracy? Is it not better to simply increase the magnification watching } the area of interest all the way up before the analysis and all the way } down after the analysis? } } What do you think?
************************************** * Pete Ainsworth * * Dept. Geology & Applied Geology * * Lillybank Gardens * * University of Glasgow * * Glasgow G12 8QQ * * e-mail: ainswort-at-geology.gla.ac.uk * * Tel : 0141 330 5505 (direct) * **************************************
} ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } ----------------------------- } -----------------------------------------. } } I travel the world teaching practical electron microscopy and it } worries me } the emphasis that people place on "Spot Mode". I do not teach the use } of } spot mode for the following reasons- } } 1. Just because the spot appears in a certain position on the } screen } is this its correct position. I have found machines many microns out } of } step in X and Y directions. } } 2. Specimens charge and switching out of spot mode does not give } the } operator an easy opportunity to see if the spot had moved during the } analysis. } } 3. Spot mode gives a false sense of analytical volume, no matter } how } small that spot may be we are almost certainly evaluating microns of } material. } } Do others worry about spot mode accuracy, do others test the spot mode } } accuracy? Is it not better to simply increase the magnification } watching } the area of interest all the way up before the analysis and all the } way } down after the analysis? } } What do you think?
Stephen - you are absolutely right - infact the only way to be sure where the spot was is to look for the specimen damage after the fact! If you have to analyze a small area (first think about the excitation volume) you are better off doing a redcued area scan or go to high magnification so you can at least "see" where the beam is hitting the sample.
I am looking for an electron diffraction analysis software package. I have to identify some particles (wether they're cubic, tetragonal, etc) and there must be an easier way than to hand draw the expected diffraction patterns....Does anybody know of something useful on the Web?
Thanx Sara
========================================================== Sara Prins Surface and Structure Analytical Services Division for Material Science and Technology CSIR PO Box 395 Pretoria SOUTH AFRICA
Depending on the stability of your stage/specimen, you should verify the position of the spot at required intervals. My collegues and I and numerous grad. students and post-docs have used this approach for decades.
On Fri, 4 Jul 1997, Bill Miller wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Stephen Chapman wrote: } } } ------------------------------------------------------------------------ } } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } } America } } To Subscribe/Unsubscribe -- Send Email to } } ListServer-at-MSA.Microscopy.Com } } ----------------------------- } } -----------------------------------------. } } } } I travel the world teaching practical electron microscopy and it } } worries me } } the emphasis that people place on "Spot Mode". I do not teach the use } } of } } spot mode for the following reasons- } } } } 1. Just because the spot appears in a certain position on the } } screen } } is this its correct position. I have found machines many microns out } } of } } step in X and Y directions. } } } } 2. Specimens charge and switching out of spot mode does not give } } the } } operator an easy opportunity to see if the spot had moved during the } } analysis. } } } } 3. Spot mode gives a false sense of analytical volume, no matter } } how } } small that spot may be we are almost certainly evaluating microns of } } material. } } } } Do others worry about spot mode accuracy, do others test the spot mode } } } } accuracy? Is it not better to simply increase the magnification } } watching } } the area of interest all the way up before the analysis and all the } } way } } down after the analysis? } } } } What do you think? } } Stephen - you are absolutely right - infact the only way to be sure } where the spot was is to look for the specimen damage after the fact! If } you have to analyze a small area (first think about the excitation } volume) you are better off doing a redcued area scan or go to high } magnification so you can at least "see" where the beam is hitting the } sample. } } Bill Miller } }
} I have made an experiment with mites and than an embedding with Spurr=B4s } epoxy resin. After polymerisation the blocks are fragile and shatter. Is } there a possiblity to resolve Spurr=B4s and than to repeat the embedding of } the samples or other solutions?
Spurr's is notoriously susceptible to moisture which, when present, results in a brittle block that shatters during trimming. The moisture may be absorbed from the atmosphere if one allows the uncapped specimens to sit overnight (presumably to evaporate propylene oxide) or, of course, it may not have been removed properly from the specimen during the dehydration stage. I suspect that your resin has absorbed moisture from the atmoshere. Check this out by polymerizing a block of plain resin. If it shatters, toss out any components that may have absorbed moisture and start over. Spurr's is a good resin but it is toxic, potentially carcinogenic and sometimes a pain. Take precautions against contact (nitrile gloves), inhalation (fume or exhaust hood) and properly dispose of the components by polymerizing them.
########################### Dr. John Bozzola, Director Center for Electron Microscopy Southern Illinois University Carbondale, IL 62901 Phone: 618-453-3730 =46ax: 618-453-2665 ###########################
Dear friends, I have some problem in interpreting my transmittance electron microscopy. Please, let me know if you can help me. Sincerely Soroush Sardari PhD student, Faculty of Pharmacy, Univ. of Alberta, Edmonton, Canada, T6G 2N8 Fax: (403) 492-1217
I routinely demonstrate to users of our JEOL 840 that they need to be careful of hysteris during scanning. We have TV rate, a rapid scan, and two slow scan modes on our SEM. I switch on the cross hair display at TV rate and select a point of interest. I then switch to rapid scan and have to recenter the cross hairs. I then switch to slow scan mode and recenter again. Then I ask them which is correct? Of course the slower scan is most nearly correct. I can demonstrate by watching the brightness LEDs as I move the spot across a bright feature.
Fortunately, we are not using spot mode, per se, all that much. Both our scopes now have digital imaging with point-and-shoot x-ray analysis. It is much easier locating the point(s) of interest. But users still need to be aware that there may some lag in the scan while recording the digital image. Since we tend to use rather slow scans that is not much of a problem. But users still need to be aware.
Depending on the rush, I still often use your method of cranking up the manification so that the feature fills the field of view. I am fairly assured that what I see is what I get for x-rays.
At 03:23 AM 7/4/97 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
One way to record experimental data on objects, rooms, and even reciprocal-space, may be to record images along directions evenly-distributed over the surface of a sphere. As Bucky Fuller pointed out in our neck of the woods some years ago, the 20 face-normals of an icosahedron are especially convenient in this regard, in part because the pictures required can be shot with a single 24-exposure roll of film.
To illustrate, HTML templates for exploring and animating data-sets on a Mathematica-drawn Klein bottle, and our Philips EM430ST in its new low-vibration site, are now on the web*. A technical note on the angles for taking such pictures is linked to each template set. Let me know if you decide to make available similar views from your own labs, or have other application-related questions we might help with.
} A major problems with scanning at high mag are either the beam is } blanked while the beam waits for "start frame" or "start line" sync } signals, thus count times are inaccurate ... or, if the beam isn't } blanked then the volume analyzed is weighted to the upper left corner } and left edge ...
These aren't the only alternatives. I can't speak for other EDS manufacturers, but our system links the scan generator and the X-ray pulse processor so that raster retrace and settling times are treated as "dead time". So the counting (live) time is correct and the pixel weightings are equal.
} 1. Just because the spot appears in a certain position on the screen } is this its correct position. I have found machines many microns out of } step in X and Y directions.
In my case, I'm always worried about this. I have, however, found that for every electron-opaque object I have measured I have found the ap- propriate element(s) when the beam is on the object and not when it is ~1 micrometer away. In many cases, there is a dark spot at the measured site after the measurement, but not before. Since these spots are at the position where the beam was set, and since occasional specimen drifts are accompanied be shifts of the spot and lowered or absent peaks (compared to subsequent measurements), I think I can be confident about the positions of my analyses. } } 2. Specimens charge and switching out of spot mode does not give the } operator an easy opportunity to see if the spot had moved during the } analysis. } A distinct advantage of the use of high voltage (~1MV) and very low beam current is that charging is very seldom a problem. Since I don't have scanning capability, it is of little concern if I have to take a long time to accumulate counts. If I were to do mapping, however, low beam current would be very detrimental.
} 3. Spot mode gives a false sense of analytical volume, no matter how } small that spot may be we are almost certainly evaluating microns of } material. } Especially in my case where the typical specimen is a biological section ~1 micrometer thick. David Joy's Monte Carlo program shows pretty well what the measurement volumes are.
} Do others worry about spot mode accuracy, do others test the spot mode } accuracy? Is it not better to simply increase the magnification watching } the area of interest all the way up before the analysis and all the way } down after the analysis?
Having complete control of the spot size by being able to set the condenser aperture and two condenser lenses independently of the magnifica- tion, I just use standard settings for each analysis--10K mag, 30 micrometer C2 aperture, maximally excited C1 lens, C2 to crossover. Yours, Bill Tivol
Dear Colleagues: Recently we came across a sample for SEM which we have great difficulty to process. We would like to ask for your help: This is a kind of poly-electrolyte polymer, that has been treated to form microspheres ( 1 to 10 micron in diameter ) and cross-linked to maintein the folded shape. Since the beads are highly negatively charged they do not stick to poly-L-lysine coated cover slips. We tried to run the pellets in Eppendorf tubes through alcohol dehydration, critical point drying, and then sprinkled the dried ( as powder) onto double-sided scotch tape and sputter coated. The results are generally not satisfactory partly due to the fact that it is very difficult to dry these beads thoroughly in tubes. I wonder whether there have been better methods for this type of sample which we do not know of. Any suggestions will be very much appreciated. Best regards, Yuhui Xu
} I travel the world teaching practical electron microscopy and it worries me } the emphasis that people place on "Spot Mode". I do not teach the use of } spot mode for the following reasons- } } 1. Just because the spot appears in a certain position on the screen } is this its correct position. I have found machines many microns out of } step in X and Y directions. } } 2. Specimens charge and switching out of spot mode does not give the } operator an easy opportunity to see if the spot had moved during the } analysis. } } 3. Spot mode gives a false sense of analytical volume, no matter how } small that spot may be we are almost certainly evaluating microns of } material. } } Do others worry about spot mode accuracy, do others test the spot mode } accuracy? Is it not better to simply increase the magnification watching } the area of interest all the way up before the analysis and all the way } down after the analysis? } } What do you think?
While the spot mode leaves a lot to be desired, particularly if the user isn't aware of the problems, analysis in raster mode is not without its difficulties. Most SEMs use some sort of of sychronisation of the line and frame scan signals. If so, the analysis obtained in raster mode is biased towards one edge and one corner of an area which may not even be identical with the image area - the beam 'waits' at the beginning of each line and each frame to synch, and the display may actually be blanked for a small distance after the scan starts and before the scan ends (over scanning) to hide image distortions arising from hysteresis, which is usually most evident at the edges of the scans.
Get a nice dirty specimen and scan it with a coarse raster until the contamination builds up. Now image the area at a lower mag, and look at the contamination pattern - there will probably be a heavier contamination line down one edge (the line scan 'wait') and a spot at one corner (the frame 'wait'). In addition, you may also be unlucky enough to find that the lines making up the coarse raster are bent at each end - you don't see the distortion in your images however because this part of the frame is blanked from display, but it will contribute to an EDX analysis.
More modern SEMs have reduced these problems, but I believe that they are still present in many.
I think that this demonstrates that whatever 'employers' might want, and manufacturers try to provide, for anything beyond the most basic EM, you need a skilled and trained operator with a full understanding of the instrumentation, and the time to fully check and calibrate the machine (who needs to be properly paid for undertaking a highly sophisticated and skilled job). Otherwise you get results that are at best doubtful and at worst wrong.
Malcolm Thomas wrote: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Dear microscopists, } We will soon be purchasing an inverted metallograph for specimen prep. It } appears all the major brands are quite good, but perhaps there are subtle } advantages / disadvantages that are only evident after much use and experience. } If anyone has any strong recommendations, I would appreciate your input. } Please feel free to contact me off-line if you prefer. Thanks in advance } for your time and help. } } Sincerely, } Mick Thomas } } Materials Science Center } Cornell University } Ithaca, NY 14853 } mgt3-at-msc.cornell.edu Hi Mick,
Several of my customers have the new Leica. It is a low profile scope, provides easy access, very little stage drift and great amount of working distance. It is a new design and seems to be very popular. Call the 800 number for information for their number (800) 555-1212.
With regards to my recommendation for the LocTite 460, how did it perform?
Good Luck,
Sincerely,
Gary Liechty Allied High Tech Products 2376 E. Pacifica Place Rancho Dominguez, Ca 90220
800-675-1118 310-762-6808 Fax
Products for Materiallographic, SEM and TEM Sample Preparation
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I will soon need to get one or more new diamond knives for use with biological specimens. I haven't purchased a new knife for several years, and am curious about the quality of knives currently available. I'd appreciate hearing from anyone who has purchased and used a new knife in the last year or so.
Does anyone know of a method/stain that would localize glycogen in cryo sections of tissue (specifically goldfish neural retina) fixed with 4% paraformaldehyde? We are interested in the light level, not EM. Linda Barthel Research Associate II Department of Anatomy and Cell Biology University of Michigan lab (313) 764-7476 fax (313) 763-1166 barthel-at-umich.edu
You may simply put one drop of your sample onto a 0.45 u milipore filter or a normal stub and air dry it for 2-3 hours.
Good luck,
On Mon, 7 Jul 1997, yuhui xu wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Dear Colleagues: } Recently we came across a sample for SEM which we have great difficulty to } process. We would like to ask for your help: } This is a kind of poly-electrolyte polymer, that has been treated to form } microspheres ( 1 to 10 micron in diameter ) and cross-linked to maintein the } folded shape. Since the beads are highly negatively charged they do not stick } to poly-L-lysine coated cover slips. We tried to run the pellets in Eppendorf } tubes through alcohol dehydration, critical point drying, and then sprinkled } the dried ( as powder) onto double-sided scotch tape and sputter coated. The } results are generally not satisfactory partly due to the fact that it is very } difficult to dry these beads thoroughly in tubes. I wonder whether there have } been better methods for this type of sample which we do not know of. Any } suggestions will be very much appreciated. } Best regards, } Yuhui Xu }
*********************************************** * Ming H. Chen, PhD * * Medicine/Dentistry Electron Microscopy Unit * * University Of Alberta. * * Edmonton, Alberta, Canada * * * * Visit My Page At: * * http://www.ualberta.ca/~mingchen * ***********************************************
I have been trying to identify ESEM EDS resolution using a 50 micron diameter Nickel standard embedded in epoxy (C, O and Cl). My instrument parameters are 1: 15kv 2: 12.0 mm working distance 3: Chamber pressure=3.5T 4: Chamber Gas = N2 or H2O (did'nt make any difference which gas was used) 5: Condensor setting = 50% 6: Sample tilt = 30, 20, 10. Doing a spot analysis in the center of the Ni standard showed abundant C and O, and some Cl as well as the expected Ni counts. Is the skirting effect of the ESEM such that one can't perform EDS on smples of 50 microns or less? I would be interested in hearing from anyone who has any results for ESEM EDS resolution.
PAS will stain glycogen but also other things in frozen sections. Can the sections be treated to reomve lipids? Otherwise, how about Best's Carmine? Give a day or two and I'll look in Lillie and Fullmer.
Geoff -- *************************************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane Piscataway, NJ 08854 voice: (732)-235-4583; fax -4029 e-mail: mcauliff-at-umdnj.edu ***************************************************************
Responding to the message of {199707072115.AA24968-at-relay.ppco.com} from gllovel-at-ppco.com (Gary Lovell): } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } I have been trying to identify ESEM EDS resolution using a 50 micron } diameter Nickel standard embedded in epoxy (C, O and Cl). My instrument } parameters are 1: 15kv 2: 12.0 mm working distance 3: Chamber } pressure=3.5T 4: Chamber Gas = N2 or H2O (did'nt make any difference which } gas was used) 5: Condensor setting = 50% 6: Sample tilt = 30, 20, 10. } Doing a spot analysis in the center of the Ni standard showed abundant C and } O, and some Cl as well as the expected Ni counts. Is the skirting effect of } the ESEM such that one can't perform EDS on smples of 50 microns or less? I } would be interested in hearing from anyone who has any results for ESEM EDS } resolution. } } From the literature on this subject I think that the chamber pressure is too high; resulting in too much scattering of the incoming beam. There are several papers by Brendon Griffin in Australia, and by the Danish group in RISO - Horsewell, Appel and Bilde-Sorensen - where they have extensively documented the EDS resolution as a function of voltage, pressure and working distance. See MSA proceedings from 1995 and 1996 for papers from the ESEM symposia which cover this topic
Good luck,
__________________ Stuart McKernan stuartm-at-tc.umn.edu Microscopy Specialist CIE Characterization Facility, University of Minnesota Phone: (612) 626-7594 100 Union Street S. E., Minneapolis, MN 55455 Lab: (612) 624-6590
In my undergrad. zoology lab long ago we added 7-Up to slow down marine invertebrates. Essentially they are a bit oxygen starved from all the CO2. Nicotine would cause larger organisms, such as chitons, to curl and distort.
} } } } Further to my earlier note about tobacco smoke, I now have the } } reference: CFA OPantin. Notes on Microscopical Technique for } } Zoologists. Cambridge University Press. 1969., page 7. } } } } Funnily enough, guess what it is recommended for - Paramecium !! Also } } flagellates, the cilia of Mytilus and Hydra. Not very politically } correct (but } } in certain matters, I may not be!). Expedient, if not finally terminal } for } } both } } the Paramecium and the microscopist!! } } An aqueous extract of tobacco leaf usually has enough nicotine to make } them } comatose. And if you're a classroom volunteer, it's a dramatic & } definitely PC demonstration for the young folk... } Caroline } } Glen MacDonald Virginia Bloedel Hearing Research Center Box 35-7923 University of Washington Seattle, WA 98195-7923 (206) 616-4156 glenmac-at-u.washington.edu *---------------------------------------------------------------------* The box said "Requires Windows 95 or better.", so I bought a Macintosh. *---------------------------------------------------------------------*
Hi Guys, I'm currently acquiring mosaic images of muscle fibers with Mosaic Tiling under Oncor Image version 2.0.5d. However, the copy of Mosaic Tiling that I have doesn't allow me to get high resolution mosaic image. Does anyone know if there is a copy of Mosaic Tiling for Oncor or anyother alternative that will allow me to get high resolution mosaic images good for quantification. Thanks,
--Ciprian Have fun and keep the sun on your back and a smile on your face. __________________________________________________________ Ciprian A. Almonte University of Pittsburgh Center for Biologic Imaging Pittsburgh, PA 15261
Visit my web site at http://www.pitt.edu/~calmonte Laboratory's website: http://sbic6.sbic.pitt.edu __________________________________________________________
Dear Roadwalk, The main difference between a biological and metallurgical light microscope is that the metallurgical microscope views in the reflection mode, where the light comes through the objective lens and reflects off the opaque sample, since very few metal samples are transparent to light. Many metallurgical microscopes are inverted, that is, the sample is put upside down on a stage on the top of the microscope, with the objective nosepiece underneath, so there is very little restriction on sample size. Several of our microscopes are dual purpose and can work in either transmitted or reflected light mode. They look like an ordinary biological microscope.
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Regards, Mary Mary Mager Electron Microscopist Metals and Materials Eng., UBC 6350 Stores Rd. Vancouver, B.C. V6T 1Z4 CANADA tel:604-822-5648, fax:604-822-3619 e-mail: mager-at-unixg.ubc.ca
We would like to examine a pore size of cellulose acetate with SEM. Please advise us, how to prepare the sample becuase it is very sensitive to vacuum. Can we dry by using a CPD?
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Linda, Periodic Acid Schiff is first favourite. Any histochemistry text will give you the necessary details, failing that, e-mail me and I can send you my protocol. Ian.
well I now know why I am having difficulty finding the Amoeba since according to the book I have just bought, at least the classic Amoeba Proteus is rare in the wild!!! I remember being told by a retired microscopist that it was rare also. So at least I can take comfort in the fact that its more than likely there are none in my samples rather than my mis-identification! But I will not be giving up the search as everyone needs a mission in life *laugh*
Conrad
------------------------------------------------------------------------ ----------- "Any sufficiently advanced technology is indistinguishable from magic" ----------------------------------------------------------- Arthur C Clarke ----
I am trying to attatch various small samples (Diatoms, Microspores and Cecaria) onto Poly-L-lysine coated coverslips, but am finding that the samples do not attach.
Does anyone have any ideas?
My Method
Clean coverslips in 100% ethanol Air dry 0.1% Poly-L-lysine (MW60000) in distilled water for 1minute Rinse in distilled water Drop of fixative containg sample (30 - 60 minutes) rinse, dehydrate and CPD
I also remember reading somewhere about using Poly-D-lysine instead, has anyone tried this?
Thanks
Kevin Mackenzie Tillydrone E.M. Unit University of Aberdeen Tillydrone Avenue Aberdeen AB9 2NT
Tel 01224-272847 Fax 01224-272396 Web site- http://www.abdn.ac.uk/~nhi691/
} Responding to the message of {199707072115.AA24968-at-relay.ppco.com} } from gllovel-at-ppco.com (Gary Lovell):
} } I have been trying to identify ESEM EDS resolution using a 50 micron } } diameter Nickel standard embedded in epoxy (C, O and Cl). My instrument } } parameters are 1: 15kv 2: 12.0 mm working distance 3: Chamber } } pressure=3.5T 4: Chamber Gas = N2 or H2O (did'nt make any difference which } } gas was used) 5: Condensor setting = 50% 6: Sample tilt = 30, 20, 10. } } Doing a spot analysis in the center of the Ni standard showed abundant C and } } O, and some Cl as well as the expected Ni counts. Is the skirting effect of } } the ESEM such that one can't perform EDS on smples of 50 microns or less? I } } would be interested in hearing from anyone who has any results for ESEM EDS } } resolution. } } } } } } From the literature on this subject I think that the chamber pressure is too } high; resulting in too much scattering of the incoming beam. There are several } papers by Brendon Griffin in Australia, and by the Danish group in RISO - } Horsewell, Appel and Bilde-Sorensen - where they have extensively documented the } EDS resolution as a function of voltage, pressure and working distance. See MSA } proceedings from 1995 and 1996 for papers from the ESEM symposia which cover } this topic } } Good luck, } } Our group in Cambridge have also been investigating this, and will be presenting a paper at this years MSA conference in Cleveland :-
'Variations in the probe beam broadening with operating conditions in ESEM'
Tuesday 11:45 - Rm. 206
Ian Bache Research Student Polymers & Colloids Group, Cavendish Laboratory Cambridge University, Madingley Road, Cambridge CB3 0HE UK
SIGEE, D.C. and GILPIN, C.J. (1994) X-ray microanalysis with the environmental scanning electron microscope: Interpretation of data obtained under different atmospheric conditions. Scanning Microscopy Supplement 8: 219-229
1 GILPIN, C.J. and SIGEE, D.C. (1995) X-ray microanalysis of wet biological specimens in the environmental scanning electron microscope 1. Reduction of specimen distance under different atmospheric conditions. Journal of Microscopy 179: 22-28
Chris
Chris Gilpin Biological Sciences Electron Microscope Unit G452 Stopford Building Oxford Road Manchester M13 9PT phone +44 161 275 5170 fax +44 161 275 5171 http://www.biomed.man.ac.uk/biology/emunit/emhome.html
We do both theoretical and experimental work on high pressure SEM microscopy . (This is a noncommercial name for ESEM.) We have been looking at the effect of water vapour and water liquid layer on the emission characteristic x-rays. We have recently deduce from our Monte Carlo calculations that the true contribution from the 'image' area can be estimated by applying a correction factor. 'Resolved image area' in turn depends upon the thickness of the water layer or the pressure of the water vapour above the specimen. The exact proportion of the characteristic x-rays coming from the actual region being analysed increases with the 'increase' in the resolavable radius. Thus the value of the correction factor reduces as the resolution worsens. These results and others will be presented at the Meeting MSA.
Jitu Shah H.H. Wills Physics Laboratory, University of Bristol Royal Fort. Bristol BS 8 1TL UK email: jss-at-siva.bristol.ac.uk
} Stuart McKernan wrote: } } } Responding to the message of {199707072115.AA24968-at-relay.ppco.com} } } from gllovel-at-ppco.com (Gary Lovell): } } } } I have been trying to identify ESEM EDS resolution using a 50 micron } } } diameter Nickel standard embedded in epoxy (C, O and Cl). My instrument } } } parameters are 1: 15kv 2: 12.0 mm working distance 3: Chamber } } } pressure=3.5T 4: Chamber Gas = N2 or H2O (did'nt make any difference which } } } gas was used) 5: Condensor setting = 50% 6: Sample tilt = 30, 20, 10. } } } Doing a spot analysis in the center of the Ni standard showed abundant C and } } } O, and some Cl as well as the expected Ni counts. Is the skirting effect of } } } the ESEM such that one can't perform EDS on smples of 50 microns or less? I } } } would be interested in hearing from anyone who has any results for ESEM EDS } } } resolution. } } } } } } } } } From the literature on this subject I think that the chamber pressure is too } } high; resulting in too much scattering of the incoming beam. There are several } } papers by Brendon Griffin in Australia, and by the Danish group in RISO - } } Horsewell, Appel and Bilde-Sorensen - where they have extensively documented the } } EDS resolution as a function of voltage, pressure and working distance. See MSA } } proceedings from 1995 and 1996 for papers from the ESEM symposia which cover } } this topic } } } } Good luck, } } } } Ian Bache wrote: } Our group in Cambridge have also been investigating this, and will be } presenting a paper at this years MSA conference in Cleveland :- } } 'Variations in the probe beam broadening with operating conditions in } ESEM'
We do both theoretical and experimental work on high pressure SEM microscopy . (This is a noncommercial name for ESEM.) We have been looking at the effect of water vapour and water liquid layer on the emission characteristic x-rays. We have recently deduce from our Monte Carlo calculations that the true contribution from the 'image' area can be estimated by applying a correction factor. 'Resolved image area' in turn depends upon the thickness of the water layer or the pressure of the water vapour above the specimen. The exact proportion of the characteristic x-rays coming from the actual region being analysed increases with the 'increase' in the resolavable radius. Thus the value of the correction factor reduces as the resolution worsens. These results and others will be presented at the Meeting MSA.
Jitu Shah H.H. Wills Physics Laboratory, University of Bristol Royal Fort. Bristol BS 8 1TL UK email: jss-at-siva.bristol.ac.uk
} Stuart McKernan wrote: } } } Responding to the message of {199707072115.AA24968-at-relay.ppco.com} } } from gllovel-at-ppco.com (Gary Lovell): } } } } I have been trying to identify ESEM EDS resolution using a 50 micron } } } diameter Nickel standard embedded in epoxy (C, O and Cl). My instrument } } } parameters are 1: 15kv 2: 12.0 mm working distance 3: Chamber } } } pressure=3.5T 4: Chamber Gas = N2 or H2O (did'nt make any difference which } } } gas was used) 5: Condensor setting = 50% 6: Sample tilt = 30, 20, 10. } } } Doing a spot analysis in the center of the Ni standard showed abundant C and } } } O, and some Cl as well as the expected Ni counts. Is the skirting effect of } } } the ESEM such that one can't perform EDS on smples of 50 microns or less? I } } } would be interested in hearing from anyone who has any results for ESEM EDS } } } resolution. } } } } } } } } } From the literature on this subject I think that the chamber pressure is too } } high; resulting in too much scattering of the incoming beam. There are several } } papers by Brendon Griffin in Australia, and by the Danish group in RISO - } } Horsewell, Appel and Bilde-Sorensen - where they have extensively documented the } } EDS resolution as a function of voltage, pressure and working distance. See MSA } } proceedings from 1995 and 1996 for papers from the ESEM symposia which cover } } this topic } } } } Good luck, } } } } Ian Bache wrote: } Our group in Cambridge have also been investigating this, and will be } presenting a paper at this years MSA conference in Cleveland :- } } 'Variations in the probe beam broadening with operating conditions in } ESEM'
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Why would a silica shell of a diatom stick to a positively charge surface---- You can collect diatoms on a 0.22 micron or 0.45 micron silver filter sold by the electron microscopy houses. These filters are conductive if you are going to look at the diatoms in SEM.
George C. Ruben Dept. Biological Sciences Dartmouth College Hanover, NH USA
Mr-Received: by mta FWVX01; Relayed; Tue, 08 Jul 1997 08:37:27 -0400 Alternate-Recipient: prohibited Disclose-Recipients: prohibited
Gary,
I noticed a couple of things about your conditions which are probably not optimum for the best resolution. I assume you are using one of the Electroscan instruments. If so, you can reduce the beam skirt by using the long assembly secondary electron detector (17 mm) and a working distance of 19 mm. That leaves a 2 mm gas path length. Also, the voltage to the secondary detector has a profound effect on many of the measurement parameters such as the beam current (see Wight in 1996 MSA proceedings p838-839). Reducing that voltage probably has a beneficial effect As was mentioned previously, the size of the skirt is directly related to the gas pressure and one wants to operate as low as possible for the samples being examined. I use 2.0 torr for my polymer samples and lower for others. Also, higher accelerating voltages reduce the skirt size but at an obvious cost if doing EDS of light elements.
Notice how cleverly I gave an answer without answering your question? I should have been a lawyer or better a politician! Frankly, I don't think the final word has been written on EDS-ESEM resolution. The original work seemed to indicate that we should be talking millimeters not micrometers. More recent work, however, suggests that maybe micrometers are appropriate with sufficient care. There is a session on Tuesday morning on the ESEM at the MSA/MAS meeting and there will surely be more discussion of this issue there and in the literature.
By the way, I have been trying to locate the articles by Horsewell, Appel and Bilde-Sorenson mentioned by Stuart McKernan with no success. Does anyone have any suggesstions on how to get ahold of them?
Dear Yuhui; For particulate samples that cannot be air dried, I sandwich the sample between two Millipore filters held in a Millipore Swinney stainless steel filter holder. It is designed to fit on a syringe, but I cut the ends off (which allows for good fluid exchange), and run the holder through the dehydration series and CPD. Alternatively, some samples can be dried in HMDS or TMS, and you can skip the critical point dryer. Also, I recomend using conductive carbon tabs for your adhesive. They give a nice smooth background, and are more conductive than double-stick tape (mine came from Pella, but other companies may also offer them). Please feel free to contact me off-line for more details if necessary.
Leslie Eibest Zoology Dept., Box 90325 Duke University Durham, NC 27708 USA (919) 684-2547 leibest-at-duke.edu
} Assuming there is a device built on a 1cm by 1cm Si wafer and the } device has several 1-2nm thick layers and about 0.2um wide circuit } "wiring", how can you find failure points, preferably without breaking } the device? I know Sandi is doing X-TEM of devices and am really curious } about the way to find, say, a short circuit on the device. } Any comments are greatly appreciated. } Dear Chao-Ying, What are the compositions of the parts of the device you want to examine? That is, are the "wires" also mainly Si, are they a different material with similar Z--i.e. aluminum--or do they have much different Z? What I'm really asking is if there is a contrast-producing effect in the device. If all the device is Si with small amounts of various dopants for the different parts, you will have a difficult time, but if not, either imaging or element mapping could give you the info you want. Other questions are what resolution is necessary and what is the total thickness of the device--is there a thick backing? Yours, Bill Tivol
I have worked in biological EM for 13 years now, but am curious about metallurgical EM in Eng. and am wondering how difficult it would be for me to make a switch, because I've noticed that there seems to be more jobs for EM technologists in metallurgy than biology.
Are there any people out there that have switched from one side to the other? I think that looking after the microscope would be essentially the same, but I have no experience in electro-polishing and sample preparation for metallurgy. How difficult is this?
I am interested in hearing from anyone doing diagnostic EM with a company called MDS. We are about to be privatized here, but I have no idea what this company plans to do with electron microscopy. All the information that I have regarding MDS basically only applies to their high volume "core-lab" but not to their small specialized labs.
Responding to the message of {01IKZKXJZCKW90OHU4-at-mr.fwvx03.com} from "Robert A. CARLTON 610-454-3949" {CARLTRA-at-rpr.rpna.com} : } } By the way, I have been trying to locate the articles by Horsewell, } Appel and Bilde-Sorenson mentioned by Stuart McKernan with no success. } Does anyone have any suggesstions on how to get ahold of them? } The references I have are all to conference proceedings: MSA 1996 p847, Scandem 1996 Aarhus Denmark, Scandem 1997 Goteborg Sweden (and EUREM 1996 Dublin Ireland - proceedings on CR-ROM; incomplete and virtually useless!)
__________________ Stuart McKernan stuartm-at-tc.umn.edu Microscopy Specialist CIE Characterization Facility, University of Minnesota Phone: (612) 626-7594 100 Union Street S. E., Minneapolis, MN 55455 Lab: (612) 624-6590
} Date: Tue, 8 Jul 1997 08:51:14 +0100 (BST) } From: Kevin Mackenzie {nhi691-at-abdn.ac.uk} } To: Microscopy-at-sparc5.microscopy.com } Subject: Poly-L-lysine problem } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Hi } } I am trying to attatch various small samples (Diatoms, Microspores and } Cecaria) onto Poly-L-lysine coated coverslips, but am finding that the } samples do not attach. } } Does anyone have any ideas? } } My Method } } Clean coverslips in 100% ethanol } Air dry } 0.1% Poly-L-lysine (MW60000) in distilled water for 1minute } Rinse in distilled water } Drop of fixative containg sample (30 - 60 minutes) } rinse, dehydrate and CPD } } I also remember reading somewhere about using Poly-D-lysine instead, } has anyone tried this? } } Thanks } } Kevin Mackenzie } Tillydrone E.M. Unit } University of Aberdeen } Tillydrone Avenue } Aberdeen } AB9 2NT } } Tel 01224-272847 } Fax 01224-272396 } Web site- http://www.abdn.ac.uk/~nhi691/ } SAMPLES THAT ARE FIXED FREQUENTLY DON'T LIKE TO STICK TO ANYTHING ELSE. TRY COATING YOUR SURFACE (WE USED 1% PL LYS FOR 10 MIN, RINSE, AIR-DRY), ADDING YOUR SAMPLE (LET A ROUNDED DROP SIT AND DON'T LET IT OVERFLOW THE EDGE) ON THE COVERSLIP FOR 30 MIN COVERED IN A MOIST CHAMBER. THEN REMOVE THE EXCESS FLUID WITH A PASTEUR PIPET AND *!*GENTLY*!* ADD FIXATIVE TO ONE SIDE AND LET IT SIT IN A ROUNDED UP DROP FOR 10-20 MIN. WASH WITH BUFFER (GENTLY) AND PROCEED WITH WHATEVER ELSE YOU WANT TO DO.
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
} I have been trying to identify ESEM EDS resolution using a 50 micron } diameter Nickel standard embedded in epoxy (C, O and Cl). My } instrument parameters are 1: 15kv 2: 12.0 mm working distance 3: } Chamber pressure=3.5T 4: Chamber Gas = N2 or H2O (did'nt make any } difference which gas was used) 5: Condensor setting = 50% 6: } Sample tilt = 30, 20, 10. Doing a spot analysis in the center of the } Ni standard showed abundant C and O, and some Cl as well as the } expected Ni counts. Is the skirting effect of the ESEM such that } one can't perform EDS on smples of 50 microns or less? I } would be interested in hearing from anyone who has any results for } ESEM EDS resolution.
Dear Gary,
Approximately 75 % of the primary electrons will be scattered when you use a working distance of 12 mm and a pressure of 3.5 torr and a significant fraction of these scattered electrons will hit the sample further away from the beam target than 50 micrometer. (The scattered intensity is approximately given by Is/Io = 1 - exp(-psL/kT) where p is the pressure, s the total scattering cross section for electron scattering on the gas used, L the distance between the last pressure limiting aperture and the sample, k the Boltzmann constant and T the absolute temperature). Examples of skirt shapes are e. g. given in:
D. A. Moncrieff et al., J. Phys. D: Appl. Phys. vol. 12 (1979) 481-88. D. C. Joy, Microscopy and Microanalysis ' 96, Proc. Annual Meeting MSA, Minneapolis, Minnesota, 11 - 15 August 1996
It is to a large degree possible to correct for the beam skirt effects:
1) You can extrapolate from spectral measurements made at several different pressures to the result that would have been found without scattering provided that the measurements are made in the single scattering regime (i.e. pL { approx. 1.6 Pa.m for measurements in water vapour). In order to obtain single scattering conditions you can use a so-called X-ray bullet to reduce the working distance. 2) If there is plural scattering, you can take two spectra, one with a fine needle (of the kind used for field ion microscopy or scanning tunneling microscopy) inserted over the point of interest, and the other with the needle slightly retracted. Subtraction of the first from the second spectrum will approximately give the spectrum from the point of interest.
Neither method will give you as exact an analysis as you will get under high vacuum, but you can get rid of most of the skirt effects. The pressure variation method in particular yields pretty good results if carefully performed. The methods are described in:
J. B. Bilde-Soerensen and C. C. Appel, Proc. 48th Annual Meeting of the Scandinavian Society for Electron Microscopy, Aarhus, 2 - 5 June 1996, pp. 4 - 5. J. B. Bilde-Soerensen and C. C. Appel, Proc. 11th European Congress on Microscopy EUREM' 96, Dublin, 26-30 August 1996. Session T6. J. B. Bilde-Soerensen and C. C. Appel, Proc. 49th Meeting of the Scandinavian Society for Electron Microscopy, Gothenburg, 10 - 13 June 1997, pp. 12-15.
Best wishes, Jorgen.
J. B. Bilde-Soerensen Materials Research Department Risoe National Laboratory DK-4000 Roskilde Denmark
Job Title: TEM Analyst Manager: Dr. Carolyn Gondran Department: Materials Analysis Division: Internal Technical Support - SEMATECH 2706 Montopolis Dr. Austin, TX 78741-6499
(512) 356-3149 phone (512) 356-7008 FAX
Job Summary: Provide analytical support to SEMATECH and I300I projects and to the ATDF through TEM analysis of semiconductor materials and devices (in plan view and cross section) and direct interactions with internal customers. Operate the TEM (EDAX, STEM, PEELS, Electron diffraction etc.) interpret electron micrographs, electron diffraction patterns and EDAX data and provide written reports of all analyses. Work with and oversee the activities of TEM technician(s) including selection of sample preparation techniques, preparation of samples, photographic image processing, maintenance of lab equipment and supplies and the development/refinement of new sample preparation techniques as needed.
Qualifications: An advanced degree in Physics, Chemistry or Materials Science with proven TEM experience. 5 - 10 years of experience in TEM analysis and sample preparation and analysis of semiconductor materials and devices is desirable.
Hi All I have a need for a second hand Philips CM12 or CM20 preferably with compustage. If there is anyone with one to sell or is thinking of upgrading? I would take an instrument that is underused and give full access to the owner if that would help. The need is fairly urgent so a speedy reply would be appreciated even if only to express an interest Please reply directly to me and not the list. Many thanks
Chris
Chris Gilpin Biological Sciences Electron Microscope Unit G452 Stopford Building Oxford Road Manchester M13 9PT phone +44 161 275 5170 fax +44 161 275 5171 http://www.biomed.man.ac.uk/biology/emunit/emhome.html
} As was mentioned previously, the size of the skirt } is directly related to the gas pressure and one wants to operate as } low as possible for the samples being examined. I use 2.0 torr for my } polymer samples and lower for others. Also, higher accelerating } voltages reduce the skirt size but at an obvious cost if doing EDS of } light elements.
As people who know me will already know I am a biologist using ESEM. I almost always view hydrated samples. Try keeping a sample wet at 2.0 Torr! There are many people who use an ESEM to look at dry uncoated samples and in this case there are a number of parameters available for change. In any general discussion on the list remember that not all samples and imaging requirements are the same. Sounds like lots of discussion for the ESEM session at MSA and also the users group meeting.
Chris
Chris Gilpin Biological Sciences Electron Microscope Unit G452 Stopford Building Oxford Road Manchester M13 9PT phone +44 161 275 5170 fax +44 161 275 5171 http://www.biomed.man.ac.uk/biology/emunit/emhome.html
Hi, We have a large staff and many students. We have about 22 thousand dollars worth of various kinds of diamond knives. We always buy DIATOME. The quality is excellent and the service and support superior. I would not consider buying any other knife from another company. In 15 years of buying and using these knives, I have never been sent a poor one or one with any detectable flaw. I do not even "check" them out anymore when we get one resharpened. I know it is good. (I have no stock in EMS who sells these knives). Sincerely, Hildy
Kevin Mackenzie wrote: =============================================== I am trying to attatch various small samples (Diatoms, Microspores and Cecaria) onto Poly-L-lysine coated coverslips, but am finding that the samples do not attach.
Does anyone have any ideas? ================================================ So long as these are relatively "free flowing" powders, larger than about 4 um, then one of our Tacky Dot (TM) Slide products should work fine in this application. In addition, there is the added bonus that the particles are mounted in an orthogonal array, making it possible to do analytical work on the powder far more quickly if not also more accurately.
Information about Tacky Dot Slides can be found on our website.
Disclaimer: SPI is the sole worldwide manufacturer, under license from DuPont, of Tacky Dot Slides so we have a vested interest in seeing that more are used! We know of no other product like this one so there are no other references to be given.
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 ############################ ==================================================
Paiboon NUANNIN wrote: ================================================ We would like to examine a pore size of cellulose acetate with SEM. Please advise us, how to prepare the sample becuase it is very sensitive to vacuum. Can we dry by using a CPD? ================================================= Could you give more information about your system, for example, a) what is the cellulose acetate "wet" with?, b) what would be the expected pore size, and c) what is the physical form of the sample, is it a thin film coating on a substrate or is it more of a bulk sample? With regard to cellulose acetate, I don't think we have ever found porosity in that polymer system. But then again, maybe we did not look hard enough either.
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 ############################ ==================================================
I have a customer who will be using SEM/EDS to characterize a mixed oxide layer (Fe-, Ni-, Cr-, Mn-, Si-oxides, approximately 1 micrometer thick) furnace-grown from the superalloy substrate. While measuring the desired features will be successful at a comfortable rate in the lab, the ultimate goal is to develop a low-prep, rapid, reliable process that can be used on the plant floor by semi-skilled workers.
I checked around and it looks like x-ray thickness measurement tools may do the trick. Many fluorescence units are portable, easily used, etc. and are appropriate to our sample size (approximately 1 CM2)
My questions: Do any of you use industrial-duty x-ray fluorescence thickness gages for measuring oxide layers over metal substrates? If so, how accurate, repeatable, etc. Any caveats?
If you would prefer to respond directly, I can be reached at:
Harold J. Crossman OSRAM SYLVANIA INC. Lighting Research Center 71 Cherry Hill Dr. Beverly, MA 01915 Phone: (508) 750-1717 E-mail: crossman-at-osi.sylvania.com
Our web sites: www.sylvania.com www.siemens.com --
well I now know why I am having difficulty finding the Amoeba since according to the book I have just bought, at least the classic Amoeba Proteus is rare in the wild!!! I remember being told by a retired microscopist that it was rare also. So at least I can take comfort in the fact that its more than likely there are none in my samples rather than my mis-identification! But I will not be giving up the search as everyone needs a mission in life *laugh*
Conrad
------------------------------------------------------------------------ ----------- "Any sufficiently advanced technology is indistinguishable from magic" ----------------------------------------------------------- Arthur C Clarke ----
I finally got a good book on Protozoa identification. In the end I found that the place I got my Microscope had a good book. Only 4.40 uk pounds! Thats the kind of price I like :)
Conrad
------------------------------------------------------------------------ ----------- "Any sufficiently advanced technology is indistinguishable from magic" ----------------------------------------------------------- Arthur C Clarke ----
Hello, I agree completely with the praise to DIATOME. Well deserved indeed. Sally
On Tue, 8 Jul 1997, HILDEGARD CROWLEY wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } Hi, } We have a large staff and many students. We have about 22 thousand } dollars worth of various kinds of diamond knives. We always buy } DIATOME. The quality is excellent and the service and support superior. } I would not consider buying any other knife from another company. In 15 } years of buying and using these knives, I have never been sent a poor one } or one with any detectable flaw. I do not even "check" them out anymore } when we get one resharpened. I know it is good. (I have no stock in EMS } who sells these knives). } Sincerely, } Hildy }
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Hi YuHui, You can try using Cryo-SEM. Put a drop of beads in suspension onto a substrate, blot excess solution using filter paper, and fast-freeze it. In order to expose the surface for SEM viewing, you need to etch the ice on surface, but the bottom part of beads are still embedded in ice. Then apply cryo-coating and cryo-observation in a cryo-SEM. If you have any questions, please contact me offline.
Ya Chen
Ya Chen
========================================================================= \ / Integrated Microscopy Resource (IMR)-- \ / __ an NIH Biomedical Research Resource TEL : 608-263-8481 \/ / / University of Wisconsin-Madison FAX : 608-265-4076 / / / 1675 Observatory Drive #159 Email1:ychen14-at-facstaff.wisc.edu / /__/_ Madison, WI 53706 Email2:chen-at-calshp.cals.wisc.edu ========================================================================= IMR WWW Home Page: http://www.bocklabs.wisc.edu/imr.html
The Integrated Microscopy Resource and Carnegie Mellon University will be sponsoring a symposium and short course on multi-photon excitation imaging, August 9-10, 1997, in Cleveland Ohio.
We have never gotten a bad diamond knife from MicroStar in the 10 plus years we have been dealing with them. We have at least a dozen knives and get 2 or more resharpened every year.
} } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } At 12:16 PM 7/8/97 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America Scientific Director, ICBR Electron Microscopy Core Lab PO Box 118525 Fax: 352-846-0251 University of Florida E-mail: gwe-at-biotech.ufl.edu Gainesville, FL 32611 http://www.biotech.ufl.edu/~emcl/ Home of the #1 Gators ***** "Many shall run to and fro, and knowledge shall be increased" Daniel 12:4
We are trying to fix diatomaceous algae for SEM. Does anyone have any suggestions or warnings they could pass along.
Specifically, we are interested in the best solution for long-term storage (greater than 3 months).
The four suggested so far are: 1)lugol's solution, 2)formalin, 3)1% glutaraldehyde, 3)2% paraforaldehyde EM grade, and 4)a combination of 1% glutaraldehyde and .1% paraformaldehyde.
If you would like to post directly to me, I would be happy to submit a summary to the group.
Dear Microscopists, For the Microscopy and Microanalysis '98 meeting in Atlanta, the local arrangements committee is seeking a volunteer to organize a fun 5K run/walk for Sunday morning, July 12, 1998. For further information, please contact me off-line, at my e-mail address: yolande.berta-at-mse.gatech.edu Yolande Berta Georgia Tech School of Materials Science and Engineering 778 Atlantic Dr. Atlanta, GA 30332-0245 (404)894-2545
My two cents worth on EDS at elevated pressures, I published results of quantitative analysis of 70 microns Cr-spinel in Mg-olivine matrix at pressure in the specimen chamber from 1 Tr to 16.6 Tr with still good GSE resolution. Even at 10,000x magnification and analytical window width 4.5 microns, there is small but significant contribution from Mg-olivine towards Cr-spinel composition. And yes, changing working distance (shorter) and accelerating voltage (higher) can minimise skirt. My advice is if you have to do it in ESEM, go to the lowest possible pressure in the specimen chamber to avoid charging and try WD and kV for the best results. For biological specimen use a cold stage and keep a specimen at minimum temp. close to 0 deg. In this case you will still deal with a fully hydrated specimen at relatively low pressure of water vapor in the specimen chamber ( 4.647 Tr at 0.2 deg. C) Wis Jablonski OiC EM/X-ray Microanalysis Unit, CSL, Uni of Tasmania
Ref: W Jablonski, ESEM-2020-a key research tool in the university environment. The Third Biennial Symposium on SEM Imaging and Analysis: Applications and Techniques, Proceedings, Australian Microbeam Society, Feb 15-17, Melbourne 1995, pages16-17.
PS More recent and quantitative work by Bilde-Soerensen could be a good help.
Dear Garry, If you have done biological EM then metallurgical EM should be a breeze. Although my degree is in Microbiology, all my EM has been in Metallurgy and Materials Engineering. There is much more SEM than TEM and a lot of EDX, including the problems of quantitative EDX. Specimen preparation is straight forward and can consist of just putting the metal piece on a stub and into the SEM. You need to learn all about EDX and some basic principals of metallurgy. Electropolishing is just recipe following and there are some neat instruments, like ion beam thinners, to help you. It depends on what the particular lab you work at specializes in. I don't think there is anything as difficult as biological TEM specimen preparation and ultramicrotoming in the metallurgical field. The trick is to get the training you need from someone who knows it well.
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Mary Mager Electron Microscopist Metals and Materials Eng., UBC 6350 Stores Rd. Vancouver, B.C. V6T 1Z4 CANADA tel:604-822-5648, fax:604-822-3619 e-mail: mager-at-unixg.ubc.ca
Further to my mail on EDS at elevated pressures: I would like to clarify the sentence No2: Even at 10,000x magnification and analytical window width 4.5 microns, there is a small but significant contribution from Mg-olivine towards Cr-spinel composition at the lowest ( in this case) 1 Tr specimen chamber pressure. Hope this will help. WJ
} Help! I need a source for indium foil. I need to use this to pick up } particles and residue from the equipment in our wafer fab.
Try ESPI, phone toll free (800) 638-2581, fax (818) 889-7098, at 5310 Derry Avenue, Agoura, CA 91301. Their catalog lists 9 different thicknesses each in 3 purity grades!!
I have no connection with them, I merely drool over their catalog occasionally.
Ritchie
Ritchie Sims phone: 64 9 3737599 ext 7713 Department of Geology fax: 64 9 3737435 University of Auckland Private Bag 92019 Auckland New Zealand
I have to admit a very need solution. Being restricted on budget AND the exchange rate! (totally unfair) We are using a cheap solution. We produce envelopes from a ~ 5cm x 3cm piece of lint free paper (lens paper), staple it closed and CPD the normal way.
} For particulate samples that cannot be air dried, I sandwich the } sample between two Millipore filters held in a Millipore Swinney stainless } steel filter holder. It is designed to fit on a syringe, but I cut the } ends off (which allows for good fluid exchange), and run the holder through } the dehydration series and CPD. Alternatively, some samples can be dried } in HMDS or TMS, and you can skip the critical point dryer. } Also, I recomend using conductive carbon tabs for your adhesive. } They give a nice smooth background, and are more conductive than } double-stick tape (mine came from Pella, but other companies may also } offer them). } Please feel free to contact me off-line for more details if necessary. ## [########] ## ## ## ## ## Stephan H Coeztee Electron Microscope Unit Private Bag 3 Wits 2050 South Africa
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G'day,
Do any of you have hints and advises to give me about studies of gels with TEM ? 1) What kind of preparation would you use ? 2) What would you characterise first ? 3) Would try to go for cryo TEM ? 4) Would you do replica or try to use a cold stage ?... Your experience is more than welcommed ! Thanks.
I have made a note in my palmtop to remember to bring the details of the protozoan book that I bought into work so I can send them to this list.
I can't say how good it is when compared to other possible books due to not having seen other books to compare it with! but I thought my little Observers Book was good value for 2 UK pounds and that book was concerned mainly with pond life in general. This book costs just over twice as much and is full of detailed pictures of various protozoan including of course several types of Amoeba which I am currently searching for...!
It is quite interesting though how different pictures in books can be with respect to certain creatures since I think the little observers book has a better drawing of COLEPS than this protozoan book in my opinion as it resembles a 'knurled barrel' whereas in this protozoan book it doesn't give so much an impression of a 'knurled' surface which is how it appeared to me under the 'scope!
I liked the warning about culturing Amoeba in that if you do it at too high a temperature you can favour the culture of a LETHAL PATHONOGENIC species of amoeba!!! although it says if careful it is unlikely as the temperatures it quotes are above 35C which is hot for a country like the UK!!! unless of course the central heating is turned up.....
For any amateurs on this list in the UK I obtained this book from Brunel Microscopes for the cost of 4.40 UK Pounds.
Conrad
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Dear all, we are in a desperate search for an old vacuum electronic module E-U12A Philips EM400 (Philips Cat. No. 532269514353) as our old one is down and we cannot afford to buy a new one from Philips - the old parts are outrageously expensive. Isn't there someone owning an old Philips 400 that is being replaced and could give us the module?
Thank you for any help or suggestion.
Pavel Hozak
__________________________
Pavel HOZAK, PhD Inst. of Experimental Medicine Dept. of Cell Ultrastructure & Molecular Biology Videnska 1083 142 20 Prague 4 - Krc Czech Republic
Laura L. Estok Asst. to the President M.E. Taylor Engineering, Inc. 21604 Gentry Lane Brookeville, MD 20833 Phone: 301-774-6246 * FAX: 301-774-6711 * e-mail: Metengr-at-aol.com
I'm a Metallugical engineer taking Ph.D. course on Biomaterials. My thesys is on titanium implants for odontology. My M.Sc. thesis was on a beta titanium alloy for aircraft industry and I did TEM characterization of this alloy. Now, I'm on the opposite way: I need to do histological cuts and analisys of the specimens (titanium implants inserted into the tibiae of rabbits).
I'd like you to give me some information about the interpretation of histological specimens on laser confocal microscopy or optical microscopy. First, about specimens preparation: We have a diamond wheel machine (ISOMET). I intend to embed the specimen with resin and gently cut (low speed, low weight). To obtain a histological slice, must I grind it? Till how many microns? Second: How can I quantitatively characterize de degree of osseointegration?
I hope you can help me.
Yours sincerely,
Marcelo Henrique Prado PEMM - COPPE/UFRJ Po.Box.:68505 Cidade Universitaria - Ilha do Fundao Rio de Janeiro-R.J. CEP.: 21941-900
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REGARDING Indium foil
Beckey,
The best source is the Indium Corporation of America, 1-800-4-indium. I've used the foil to capture particles for years with excellent results. I work as a semiconductor failure analyst, 19 years here at Motorola.
Hope this is the information you need. Have a nice day!
Hi, I hope that the I didn't imply that the little book that I have bought had photographs in it! It has only drawings but still at 4.40 its good value I think. I intend to post the book details when I remember to bring them in!
Conrad
------------------------------------------------------------------------ ----------- "Any sufficiently advanced technology is indistinguishable from magic" ----------------------------------------------------------- Arthur C Clarke ----
Mary and Garry wrote about switch from Biological to Metallurgical EM:
Although I agree that sample prep can be far more demanding with biological specimens, as far as TEM is concerned, one thing biologists will most likely lack is training in crystallography. Most biologists I've met are not fond of reciprocal space. Garry, if you want to switch to crystalline solids, take a few courses in crystallography. Materials Science is far more than just generating a conventional TEM image.
Ciao for now, Ken
Kenneth JT Livi Department of Earth and Planetary Sciences 34th and Charles Streets Johns Hopkins University Baltimore, Maryland 21218 USA Phone: (410) 516-8342 Fax: (410) 516-7933 e-mail: klivi-at-jhu.edu
Can any of our German colleagues help with translation please? Came across the term PHOSPHORWOLFRAM acid in a technical paper. Suspect it is phosphoric acid but want to be sure before proceeding. Thanks in advance Ronnie Houston Texas Scottish Rite Hospital for Children Dallas
In message {33C3BC88.5577-at-airmail.net} writes: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Can any of our German colleagues help with translation please? Came } across the term PHOSPHORWOLFRAM acid in a technical paper. Suspect it is } phosphoric acid but want to be sure before proceeding. } Thanks in advance } Ronnie Houston } Texas Scottish Rite Hospital for Children } Dallas
Ronnie, wolfram in the old Germanic (I think) name for tungsten, so you have phosportungstic acid there, which when dissolved in water to a few percent is good ol' PTA, commonly used as a negative stain for viruses, bacteria, particulates in transmission electron microscopy.
--
Gib Ahlstrand, MMS Newsletter Editor Electron Optical Facility, University of Minnesota, Dept. Plant Pathology 495 Borlaug Hall, St. Paul, MN 55108 (612)625-8249 612-625-9728 FAX, giba-at-puccini.crl.umn.edu
Plato: "When the mode of the music changes, the walls of the city will shake."
Chuck Berry: "There's a whole lotta shakin' goin' on!"
I have used spot modes to do EDX "maps" on the fly.
If I suspect a high concentration of a particular element in a specific local area of an image, I start acquiring and then move the spot onto and away from the feature of interest while simultaneously watching the growth of a peak for the element of interest. It is a crude but effective way of matching the location of particular to features observed.
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} I travel the world teaching practical electron microscopy and it worries me } the emphasis that people place on "Spot Mode". I do not teach the use of } spot mode for the following reasons- } } 1. Just because the spot appears in a certain position on the screen } is this its correct position. I have found machines many microns out of } step in X and Y directions. } } 2. Specimens charge and switching out of spot mode does not give the } operator an easy opportunity to see if the spot had moved during the } analysis. } } 3. Spot mode gives a false sense of analytical volume, no matter how } small that spot may be we are almost certainly evaluating microns of } material. } } Do others worry about spot mode accuracy, do others test the spot mode } accuracy? Is it not better to simply increase the magnification watching } the area of interest all the way up before the analysis and all the way } down after the analysis? } } What do you think?
While the spot mode leaves a lot to be desired, particularly if the user isn't aware of the problems, analysis in raster mode is not without its difficulties. Most SEMs use some sort of of sychronisation of the line and frame scan signals. If so, the analysis obtained in raster mode is biased towards one edge and one corner of an area which may not even be identical with the image area - the beam 'waits' at the beginning of each line and each frame to synch, and the display may actually be blanked for a small distance after the scan starts and before the scan ends (over scanning) to hide image distortions arising from hysteresis, which is usually most evident at the edges of the scans.
Get a nice dirty specimen and scan it with a coarse raster until the contamination builds up. Now image the area at a lower mag, and look at the contamination pattern - there will probably be a heavier contamination line down one edge (the line scan 'wait') and a spot at one corner (the frame 'wait'). In addition, you may also be unlucky enough to find that the lines making up the coarse raster are bent at each end - you don't see the distortion in your images however because this part of the frame is blanked from display, but it will contribute to an EDX analysis.
More modern SEMs have reduced these problems, but I believe that they are still present in many.
I think that this demonstrates that whatever 'employers' might want, and manufacturers try to provide, for anything beyond the most basic EM, you need a skilled and trained operator with a full understanding of the instrumentation, and the time to fully check and calibrate the machine (who needs to be properly paid for undertaking a highly sophisticated and skilled job). Otherwise you get results that are at best doubtful and at worst wrong.
Ronnie Houston wrote: } } ------------------------------------------------------------------------} The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------.} } Can any of our German colleagues help with translation please? Came } across the term PHOSPHORWOLFRAM acid in a technical paper. Suspect it is } phosphoric acid but want to be sure before proceeding. } Thanks in advance } Ronnie Houston } Texas Scottish Rite Hospital for Children } DallasDear Ronnie,
This sounds like a "common" rather than proper chemical name. If it helps at all, Wolfram is the source for the chemical symbol "W", for tungsten. I would try looking up tungsten salts of phosphoric acid.
Good luck.
Barbara Foster Consortium President Microscopy/Marketing & Education 53 Eton Street Springfield, MA 01108 USA (413)736-6931 fax: (413)746-9311 email: mme-at-map.com (
Linda Barthel asked for a method to localize glycogen in cryosections. I sent her some background information needed in the microscopic study of glycogen, and I repeat them for all researchers interested in the subject.
The problem of glycogen is more complex than commonly appreciated, and the understanding of this complexity is a sine qua non condition in the microscopic study of glycogen.
Glycogen in the cell appears in the organelles, GLYCOSOMES, composed of glycogen and enzymes involved in glycogen synthesis and degradation. The structures stained by uranium and lead salts, and interpreted in EM as "particles of glycogen" represent in fact the protein component of glycosomes. Glycogen does not react with the ionic stains, but it can be demonstrated histochemically, by the PAS technique (periodic acid - Schiff reagent) in LM, and by the modification of this procedure (Thiery technique) in EM.
The problem is that glycosomes are easily destroyed during tissue processing. The most common destructive factor is the change in pH which breaks the bond between glycogen and protein. The effect is that the soluble protein component (enzymes) is washed out, and glycogen which is not fixed, floats in the cell and aggregates into clumps. The acidic treatment is inherent in the PAS procedure where periodic acid is used, therefore, in the vertically processed slides, the unfixed glycogen often accumulates as crescents at the bottom of the cells (the effect well known in the classical histochemistry).
In EM the common destructive factor is uranyl acetate (strongly acidic) used before tissue dehydration. In tissue processed without uranyl acetate glycosomes appear intact, even after the priodic acid treatment in Thiery technique, because the histochemical reaction is performed on sections which are already embedded in the resin. This embedding prevents the floating of the unfixed glycogen.
Freezing seems to be another destructive factor for glycosomes. Raether et al, 1977 (Z.Parasitenkunde, 54, 149) used deep-freezing of Entamoeba cultures, and found that only a few amoebae retained normal structure. Their micrographs indicate that in the destroyed organisms the glycosomes were destroyed.
Additional complication is that the described factors affect only free glycosomes, whereas others, which are bound to different cell structures remain resistant.
The review of glycosomes was published by K.K.Rybicka, 1996, Tissue & Cell 28 (3) 253-267.
} Can any of our German colleagues help with translation please? Came } across the term PHOSPHORWOLFRAM acid in a technical paper. Suspect it is } phosphoric acid but want to be sure before proceeding. } Thanks in advance } Ronnie Houston } Texas Scottish Rite Hospital for Children } Dallas
Ronnie,
"Wolfram" is the former chemical name for the element "Tungsten," hence the symbol of "W" on the periodic table. I think it's safe to assume that PHOSPHORWOLFRAM ACID is the same as PHOSPHOTUNGSTIC ACID. Phosphotungstic acid is listed in references as a stain for EM work, but I have no experience with it myself.
Jim Passmore Analytical Chemist Cryovac North America Duncan, SC
This may have nothing to do with your problem and others on this listserver may understand my story better than I do, but you may also want to note the batch of your collodion as well as the cytochrome c. We do routine TEM autoradiography which involves coating slides with a 1% collodion/amyl acetate solution, then placing the sections on the slides, carbon coating, processing the sections through autoradiography and placing grids on the sections, then removing the collodion before scanning on the TEM. At one point a couple years ago we ran into real problems removing the collodion after the procedure - the sections were removed before the collodion!! After much investigation and many trials a friend of mine finally spoke with the chemist working at the company we bought our stock collodion solution from. Apparently they had switched to a newer? better? safer? method of preparing the stock collodion. We now specially order collodion prepared somehow with ethanol and ethyl ether - and have never had a problem since!!!
Pat Hales McGill University Dept. of Anatomy & Cell Biology hales-at-hippo.medcor.mcgill.ca
Anyone knows anything about pyroxylin? I would like to know what it is exactly, and if it might be more stable than some other films for coating grids for the TEM. (Note that I'm looking at thick stuff - no thin sections here!)
I will be purchasing a TOPCON SEM soon and I am interested in studying clays. I am most interested in the preparation techniques that investigators have used to get good pictures of kaolinite, smectite, illite, etc.
I have been using an X-ray diffractometer w/clays for years but wonder about how to mount oriented/unoriented specimens on those stubs.
I'd agree with the assessment of Diatome knives as being excellent. We have 6 of them here, and they all seemed to be of excellent quality. Of course, I also like Dupont and DDK.
My thoughts,
Garry
} ---------- } From: Greg Erdos[SMTP:gwe-at-biotech.ufl.edu] } Sent: 8 July, 1997 12:55 } To: Microscopy-at-Sparc5.Microscopy.Com } Subject: Re: Good diamond knives } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Mounting the specimen may be as simple as placing it onto a stub using a conducting adhesive, waiting 20 minutes for it to dry, and then running your new microscope at a low ( {5kV) accelerating voltage. Keep the specimen as small as possible to limit contamination. Do not fall into t= he trap of believing that every specimen should be evaluated at 25kV. Vary the kV to see how the information you visualise changes with beam penetration, there is always one particular kV that will best present the=
specimen features. Obtain a simple Monte Carlo program so that you can calculate the depth from which the backscatter and x-ray information is coming. Changing Z and tilt will vary the backscatter contribution to yo= ur image and change the way the information is presented.
However if you wish to carry out EDX analysis in the microscope you will=
need up to 25kV to enable a full range spectrum evaluation. In this case=
if the material is non conducting you will need to coat it, preferably wi= th carbon. The simplest carbon coating systems are desk top and may either = be self contained or as an accessory for a sputter coater. In either case a=
carbon string system should be all that is required. The coating system i= s not the most important feature but the type of string you use will decide=
how easy it is to coat the specimen. We find the thick string like a boo= t lace is the best. If you new SEM is of the variable pressure variety th= is will help you reduce any problems of specimen charge
For both imaging atomic number contrast and to better visualise your specimen for EDX evaluation you need to use a backscattered electron detector. Varying the kV, viewing by backscatter differing depths of information, you will effectively be able to section the specimen, from 5= kV through to the microscopes maximum.. The Monte Carlo program may be used=
Diffraction pattern indexing can be difficult even if you know the general structure, i.e. orthorhombic, because of the different length sides of the unit cell (let alone angles in other structures). Then, just because you can find a set of reciprocal space planes at the right distances and angles from each other doesn't mean that the structure factor allows for that spot to be strong in reciprocal space. So any indexing software usually requires that you know positions of atoms in the unit cells of all the structures that would potentially fit the pattern that you are trying to index. EMS can be run on unix or vax machines. You must not only know the possible structures but also a range of camera length.
But,...I have found an alternate although cumbersome and limited method: I have generated and used successfully in Microsoft Excel a 3 sheet spreadsheet that notifies you of all planes that match the input specifications of the pattern to be indexed. You must also input the cell parameters and cell angles. This does not take into account any structure factors. And, it is limited by computer RAM. If you are interested, I can tell you more how to generate it yourself -- better that my old one.
When staining sections of Douglas-fir twigs, I use safranine/fast green. I would like to see a stain difference between early and late wood, but with this stain I can only tell the difference by the size of the vessels and the thickness of their walls. I would like a cleaner method. Any ideas?
Hello everyone! I'm sending this message to you in behalf of Ms. Louise Tayl or because I believe that her question might get more replies from this netw ork. Please send your replies to Ms Taylor directly at this address:
179LOU-at-chiron.wits.ac.za Thanks!
At 1:51 PM 97.7.9, Ms Louise Taylor wrote: } Hi there histonetters } } I have a query from an electron microscopist in our department. They } are having some problems doing IHC on their samples, particularly } with HMB45 (DAKO). } Are there any suggestions out there regarding optimum dilution of the } antibody or conversely another source that works well at EM level. I } realise that this is not necessarily a histonet query, but I would } appreciate whatever contacts, info etc I can get. } } Many Thanks } Louise Taylor
Laurent NORMAND wrote: ================================================ Do any of you have hints and advises to give me about studies of gels with TEM? 1) What kind of preparation would you use ? 2) What would you characterise first? 3) Would try to go for cryo TEM ? 4) Would you do replica or try to use a cold stage ?... ================================================= There might be as many philosophies of approach as there are people doing this kind of work. My first thought is always to determine what kind of features are you really trying to resolve in the gel structure. For example , certain greases (they act as gels because they have "thickeners") contain various metal stearates that form fibrillar type networks, and the differences between samples show up by just taking the gel, often times not even diluting it but just smearing it out on a glass slide to the thinnest possible film, and after RT drying, carrying out Pt/C shadowing and replication and looking at it that way. You see the metal stearate structures quite nicely and they usually, or at least some of them, get picked up with the replica so you can even do ED and DF studies. And the experimental procedure is very definitely not a complicated one, using just conventional equipment that just about everyone has at their fingertips in their laboratories.
On the other hand, if you have something like an aviation jet fuel kind of gel, and one is interested in seeing the network features of the polymeric additives (e.g. the ones that are responsible for the gelling of the system) , freeze fracture TEM is more appropriately indicated. If you are seeking to resolve "micropores", you might see them this way as well. If the gel, such as a so-called "hydro gel" type system is being looked at, being aqeuous based, some freeze etching should also be done.
If there is some kind of a non-organic type additive, that would have enough electron density in its own right, and the goal is to see its degree of dispersion in the gel, then cryo-TEM would be our preference. If the gel is aqueous based, and the presence of pores is what is to be resolved, then we have looked for ways to precipitate silver chloride into the pores to serve as a "decorator".
The conspicuous absence of mention of SEM was not accidental. We have almost never found SEM to be good enough to resolve the kinds of features people want to see when doing this kind of work.
It did not sound like you are asking about things like "gel" spots in polymer films which of course would require entirely different approaches, none requiring cryo, except maybe for diamond knife thin sectioning.
Disclaimer: We have no connection with any of these techniques except that we do offer them as a service for clients wanting to do this type of work.
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 ############################ ==================================================
Dear Hank, When I prepare clays to see the crystal structure, I usually suspend the clay in alcohol to the consistancy of milk and then put a drop on a SEM stub with a small glass cover slip on top, to get a very smooth surface. Dry, then gold coat. This will allow you to see the "books" that are characteristic of kaolinite. I use this method to look at the different crystal shapes of kaolinite, halloysite and illite for a lab. You wrote: } } I will be purchasing a TOPCON SEM soon and I am interested in studying } clays. } I am most interested in the preparation techniques that investigators have } used } to get good pictures of kaolinite, smectite, illite, etc. } } I have been using an X-ray diffractometer w/clays for years but wonder } about how } to mount oriented/unoriented specimens on those stubs. } } I'd appreciate any help or references. } } Hank Bart Regards, Mary
Mary Mager Electron Microscopist Metals and Materials Eng., UBC 6350 Stores Rd. Vancouver, B.C. V6T 1Z4 CANADA tel:604-822-5648, fax:604-822-3619 e-mail: mager-at-unixg.ubc.ca
as it might be of interest for some of you, I wish to point you to a new listserver at the medical faculty of the Westfalian Wilhelms-University. It discusses anatomical and related topics. However, it is announced for the english and for the german language. On the other hand it has just started and traffic is still low; therefore it might become common to discuss in English as Germans appear sometimes to be a bit offish :-)
To subscribe send an e-Mail to Majordomo-at-medweb.uni-muenster.de with the following body text: subscribe ANATOMIE-D {your-eMail-address}
Sorry if I have annoyed people but with all due respect Ray, I was asked to share the information about this book by fellow amateurs. I admit that I may have rambled on a bit. Although I have been on the net for a long time, I guess with respect to this list I am one of those 'dreaded newbies'. and thought it was like a newsgroup and forgot it was composed mainly of proffesionals.
I only posted because it would have taken too much time to reply to everyone and I had better end this here before this becomes another rambling post!
Regards,
Conrad
} -----Original Message----- } From: Ray Hicks [SMTP:rh208-at-cus.cam.ac.uk] } Sent: Wednesday, July 09, 1997 6:54 PM } To: Conrad Perfett } Subject: Re: protozoan book } } Conrad, } } It's good that you've found some information useful to you, but why not } wait until someone asks before you share it? You'll notice that there } isn't very much spontaneous posting to the list by anyone else, about } microscopy but especially about their research interests. Remember that } the common interest of the list is microscopy, not microbiology, histology } or metallurgy. If, for instance, someone has a microscopy related } metallurgical question they post it and get an answer, generally from } another metallurgist, you don't tend to get "I looked at a piece of } pearlite yesterday - interesting eh?" type comments. If you did you might } get twenty anecdotes a day from each of the members of the list, and it } would stop being so useful. } } How about lurking around until someone asks a question that's in your area } of expertise, and then helping them out? } } By the way, my school teachers used to provide amoebae for practical }
Dear all, we are in a desperate search for an old vacuum electronic module E-U12A Philips EM400 (Philips Cat. No. 532269514353) as our old one is down and we cannot afford to buy a new one from Philips - the old parts are outrageously expensive. Isn't there someone owning an old Philips 400 that is being replaced and could kindly give us the module or to suggest a solution?
Thank you for any help or suggestion.
Pavel Hozak
__________________________
Pavel HOZAK, PhD Inst. of Experimental Medicine Dept. of Cell Ultrastructure & Molecular Biology Videnska 1083 142 20 Prague 4 - Krc Czech Republic
I am working on single aerosol particle analysis using windowless SEM/EDX system. I hope to quantitatively analyze the contents of C, N, and O in microparticles. CITZAF program has been known to work for particles, so I am looking for where I could get it. I know it is a shareware and was distributed by Caltech. However, Dr. Armstrong seems not working over there any more. Would someone kindly help me to have a copy of CITZAF program? Many thanks in advance.
Sincerely yours, Chul-Un Ro
Chul-Un Ro, Ph.D Department of Chemistry University of Antwerp B-2610 Wilrijk Belgium
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Howdy, We have a JEOL 2010 STEM that must be isolated from a huge vibrating air compressor in a different part of the building (no they won't move it or isolate the compressor). Our plan is to cut the floor (8" of high density reinforced concrete sitting on dirt) around the microscope. I am interested in tips and lessons learned,e.g., width of cut, filler material in the cut, is it ok to tile over the gap, how far from the scope to place the cut, etc.
Brad Storey Materials Scientist Argonne National Lab - West P.O. Box 2528 Idaho Falls, ID 83403 Ph. 208-533-7685 Fax 208-533-7683 e-mail brad.storey-at-anl.gov
Greetings, Any of you erudite types out there know the origins of the word "Wolfram"? or why or when the word "tungsten" (which I believe comes from sweedish words for "heavy + steel") came to be the prefered word? Please forgive my curiousity if this is too far off the microscopical axis for your taste.
Short article on pyroxylin in Merck Index. Cellulose nitrate in old film base and plastics is not stable over decades, but this probably does not matter for your needs. Note hazards.
I am currently staining milk proteins with phosphotungtic acid (2%/water ph 7.3) and uranyl acetate (1%/water) and often I do get what I called a background consisting of very small (about 20-40nm) bubbles-like structures that interfere with my samples...
I also recall seeing the same thing with viruses but this time it is rather annoying because it also looks a lot like my protein...
Does somebody out there has a explanation to this phenomenon....I though maybe it was perhaps a hydrophobic reaction between the formvar and the stain or irregularities/defects in the formvar film....
thank you,
Diane Montpetit Food research center agriculture canada st-hyacinthe, quebec, Canada fax 514 773 8461 tel; 514 773 1105 e mail; montpetitd-at-em.agr.ca
The tungsten mineral wolframite was known in the tin mines of the Saxony-Bohemia region long before the element itself was discovered. In 1781, the Swedish chemist Scheele, who had been working with the stony mineral, elucidated the composition of this mineral to be a compound of calcium with an unknown acid. The acid-forming element thus discovered was named tungsten by A. F. Cronstedt in 1755. He derived this name, from the Swedish words "tung", meaning heavy, and "sten", meaning stone.
Wolfram
In 1783, the brothers J.J. and F. de Elhujar found that wolframite also contained tungsten. After success in obtaining metallic tungsten from wolframite, and gave it the name "wolfram" The origin of the word is not clear, but it is assumed to be derived from German words "Wolf" and "Rahm" or Swedish word "wolfrig".
Wolfram is the official international alternate name for tungsten. Tungsten is preferred in the United States.
Hello fellow microscopists, I'm in the process of using the DGD resinless embedding technique developed by Capco, Krochmalnic and Penman and I'm looking for: 1)Short cuts-the ethanol, nbutanol to DGD transition is very long (6 hours) and I am working with monolayers, so is there a poblem with this version: Graded ethanol dehydration to 100% 30 min 100% ethanol 4 changes 1 hr 1:1 nbutanol:100% ethanol 30 min 100% nbutanol 1 hr 1:1 nbutanol:DGD 45 min 100% DGD (with DMSO) 1 hr RT harden O/N
2) Also any suggesstions on how to harden this wax on tissue culture dishes, coverslips (glass) and platic slides will be greatly appreciated. Thank you.
Dear all, we are in a desperate search for an old vacuum electronic module E-U12A Philips EM400 (Philips Cat. No. 532269514353) as our old one is down and we cannot afford to buy a new one from Philips - the old parts are outrageously expensive. Isn't there someone owning an old Philips 400 that is being replaced and could give us the module?
Thank you for any help or suggestion.
Pavel Hozak
__________________________
Pavel HOZAK, PhD Inst. of Experimental Medicine Dept. of Cell Ultrastructure & Molecular Biology Videnska 1083 142 20 Prague 4 - Krc Czech Republic
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Hi Brad, I have done a bit of work staining core samples and twig sections of Doug fir in an attempt to age the wood, and safranin staining was the best way to evaluate this. Unfortunately, the only difference between early and late wood is vessel diameter (ie. secondary cell wall thickness). There may be differences in minor cell wall components from early to late wood but none of them (I predict) would show as good a demarcation as safranin does in staining lignin. Don't overstain with fast green as you can lose contrast, in fact, it can be left out altogether because you are not interested in staining cytoplasmic components. Remember that in Doug fir there will be a gradual transition from early to late wood. You have to estimate where the change is based on an arbitrary cell wall thickness that you get to choose (I get a real rush making executive decisions like that). Cheers, John
================= C. John Runions, Ph.D Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
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Hi Brad, I have done a bit of work staining core samples and twig sections of Doug fir in an attempt to age the wood, and safranin staining was the best way to evaluate this. Unfortunately, the only difference between early and late wood is vessel diameter (ie. secondary cell wall thickness). There may be differences in minor cell wall components from early to late wood but none of them (I predict) would show as good a demarcation as safranin does in staining lignin. Don't overstain with fast green as you can lose contrast, in fact, it can be left out altogether because you are not interested in staining cytoplasmic components. Remember that in Doug fir there will be a gradual transition from early to late wood. You have to estimate where the change is based on an arbitrary cell wall thickness that you get to choose (I get a real rush making executive decisions like that). Cheers, John
================= C. John Runions, Ph.D Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
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Hi Brad, I have done a bit of work staining core samples and twig sections of Doug fir in an attempt to age the wood, and safranin staining was the best way to evaluate this. Unfortunately, the only difference between early and late wood is vessel diameter (ie. secondary cell wall thickness). There may be differences in minor cell wall components from early to late wood but none of them (I predict) would show as good a demarcation as safranin does in staining lignin. Don't overstain with fast green as you can lose contrast, in fact, it can be left out altogether because you are not interested in staining cytoplasmic components. Remember that in Doug fir there will be a gradual transition from early to late wood. You have to estimate where the change is based on an arbitrary cell wall thickness that you get to choose (I get a real rush making executive decisions like that). Cheers, John
================= C. John Runions Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
In message {s3c4b19e.001-at-EM.AGR.CA} Diane Montpetit writes: } hello everyone, } } I am currently staining milk proteins with phosphotungtic acid (2%/water } ph 7.3) and uranyl acetate (1%/water) and often I do get what I called a } background consisting of very small (about 20-40nm) bubbles-like } structures that interfere with my samples... } } I also recall seeing the same thing with viruses but this time it is rather } annoying because it also looks a lot like my protein... } } Does somebody out there has a explanation to this phenomenon....I though } maybe it was perhaps a hydrophobic reaction between the formvar and } the stain or irregularities/defects in the formvar film.... } } thank you, } } Diane Montpetit } Food research center } agriculture canada } st-hyacinthe, quebec, Canada } fax 514 773 8461 } tel; 514 773 1105 } e mail; montpetitd-at-em.agr.ca
Perhaps you could modify your formvar/grid preparation techniques to see if it makes any improvement. You could try: 1. Coat the Formvar coated grids with a thin layer of evaporated carbon. 2. Treat the Formvar coated grids in a glow discharge device if one is available. Either treatment may improve spreading and negative staining of your samples.
I looked at milk proteins, too, a few years ago using PTA staining and I don't recall any problems. The "bubbles" you see in the background may be due to contaminant that got on the Formvar film during preparation or handling, so look at your Formvar coating technique to see if you can clean it up; use pure double distilled water to float film off a previously cleaned glass slide, water in clean trough, etc, etc.
Good Luck!
--
Gib Ahlstrand, MMS Newsletter Editor Electron Optical Facility, University of Minnesota, Dept. Plant Pathology 495 Borlaug Hall, St. Paul, MN 55108 (612)625-8249 612-625-9728 FAX, giba-at-puccini.crl.umn.edu
Plato: "When the mode of the music changes, the walls of the city will shake."
Chuck Berry: "There's a whole lotta shakin' goin' on!"
} Date: Wed, 9 Jul 1997 16:22:22 -0400 (EDT) } From: Leclerc Jean {leclercj-at-magellan.umontreal.ca} } To: Microscopy Association of America {Microscopy-at-Sparc5.Microscopy.Com} } Subject: Pyroxylin coating
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Hi there!^ } } Anyone knows anything about pyroxylin? I would like to know what it is } exactly, and if it might be more stable than some other films for coating } grids for the TEM. (Note that I'm looking at thick stuff - no thin } sections here!) } } Thanks! } Our lab uses Pyroxylin, also known as parlodian or collodion, frequently, as it is easy to use. We buy as a 2% solution in amyl acetate from an EM supplier. The method: one drop on DDW in a 10mm diameter dish, let the film dry and then remove with the tip of a glass pipet to remove any dust particles, followed by another drop and let dry (interference colors become visible when viewed at a low angle). Then very gently place clean 10 to 25 grids dull side down on the film's surface.Carefully lay a piece of parafilm on the surface, pull the parafilm back up, and the grids plus film come along. Place in petri dish to thoroughly dry, followed by carbon evaporation. It is a very quick method and suitable for a number of applications.
you didn't mention what sort of coating you are using on your grid. Could these bubble structures be defects in the plastic coating caused by moisture absorbed into the liquid plastic stock solution? I must admit they sound very small but you can get small semi-perforate holes when you try to make holey plastic films.
If they are present before sample and stain they would be more difficult to see but should show up with a small objective aperture and lower KV.
Sorry if you've already thought of this.
Malcolm Haswell e.m. unit University of Sunderland UK ----------
hello everyone,
I am currently staining milk proteins with phosphotungtic acid (2%/water ph 7.3) and uranyl acetate (1%/water) and often I do get what I called a background consisting of very small (about 20-40nm) bubbles-like structures that interfere with my samples...
I also recall seeing the same thing with viruses but this time it is rather annoying because it also looks a lot like my protein...
Does somebody out there has a explanation to this phenomenon....I though maybe it was perhaps a hydrophobic reaction between the formvar and the stain or irregularities/defects in the formvar film....
thank you,
Diane Montpetit Food research center agriculture canada st-hyacinthe, quebec, Canada fax 514 773 8461 tel; 514 773 1105 e mail; montpetitd-at-em.agr.ca
We have an older, DOS-based version of Image Pro Plus image analysis software that has us stymied. We are capturing video images of chloroplast movements and dumping them into a DOS-based computer. We want to be able to sum all of the white areas in a field and then express that total white area as a percentage of the total image area. It seems simple enough, but it's not. We can identify all of the "white" objects, we can define what we are calling 'white', we can get the area of each individual white object, but we cannot sum the areas of all of the white objects. Image Pro Plus corporation is no help. They sold the rights to the DOS version to another company and no one a IPP knows anything about it. They are quite willing to sell us the Windows version for $3500 but cannot help us with their older version. Does anyone out there in microscope land have experience with this program?
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Greetings, Any of you erudite types out there know the origins of the word "Wolfram"? or why or when the word "tungsten" (which I believe comes from sweedish words for "heavy + steel") came to be the prefered word? Please forgive my curiousity if this is too far off the microscopical axis for your taste.
Back in my early graduate school days when I used to be a botanist, we used a stain called phloroglucinol which gave excellent demarcation of wood based on lignin content. I remember that you have to be very careful not to overstain, as everything will become crimson. The rest of the tissue, parenchyma etc. was contrasted with fast green. The stain can be modified to distinguish between deciduous and conifer wood (excluding Ginko), although I never tried this. The technique is described in Johansen's "Plant Microtechnique". I'm not sure if this stain would apply to your case, but it might be worth a try.
-=W.L. Steffens=- Department of Veterinary Pathology College of Veterinary Medicine University of Georgia Athens, GA 30602 USA http://www.vet.uga.edu/vpp/wls/steffens.html
The "bubbles" on the substrate that you describe are wetting artifacts...at least that is what I was told by an oldtimer. I get them consistently on negative stained viral samples, unless I incorporated a small amount (a very small amount) of bacitracin into the negative stain. I suppose some other surfactants would work, but bacitracin does the trick for me every time.
-=W.L. Steffens=- Department of Veterinary Pathology College of Veterinary Medicine University of Georgia Athens, GA 30602 USA http://www.vet.uga.edu/vpp/wls/steffens.html
My experience is better with isopropyl alcohol, more dilute suspension plus in some stubborn cases a litle ultrasonic bathing... Everything will be visible! Kris
} When I prepare clays to see the crystal structure, I usually suspend the } clay in alcohol to the consistancy of milk and then put a drop on a SEM stub } with a small glass cover slip on top, to get a very smooth surface. Dry, } then gold coat. This will allow you to see the "books" that are } characteristic of kaolinite. I use this method to look at the different } crystal shapes of kaolinite, halloysite and illite for a lab.
Disclaimer first: I'm not a chemistry/physics person, so please excuse me if this question is incredibly stupid :) Is it possible for a pigment to scatter/reflect light strongly enough to look like fluorescence? If so, how could you distinguish such behaviour from "real" fluorescence? I'm trying to work with some pigmented plant samples that "glow" under my rhodamine/Tx Red excitation/emission filters (standard fluorescence scope); the pigmented areas look normal under FITC and UV conditions. The pigment itself is red under regular light. We'd like to know if the pigment autofluoresces or if this is just some light/pigment interaction. I'm lost and our library isn't heavy on this type of references. Any of you "hard science" types willing to try to educate me?!
On Thu, 10 Jul 1997 wise-at-vaxa.cis.uwosh.edu wrote: } We have an older, DOS-based version of Image Pro Plus image } analysis software that has us stymied. We are capturing video images of } chloroplast movements and dumping them into a DOS-based computer. We want } to be able to sum all of the white areas in a field and then express that } total white area as a percentage of the total image area. It seems simple } enough, but it's not. We can identify all of the "white" objects, we can } define what we are calling 'white', we can get the area of each individual } white object, but we cannot sum the areas of all of the white objects. } Image Pro Plus corporation is no help. They sold the rights to the DOS } version to another company and no one a IPP knows anything about it. They } are quite willing to sell us the Windows version for $3500 but cannot help } us with their older version. } Does anyone out there in microscope land have experience with this } program?
Yes, a bit. Is the remaining non-white area contiguous? If so, measure that as the reciprocal of the white areas.
I'm trying to find a vendor who might sell a dissecting microscope that has 2 heads. We need something like this for demonstration purposes. It's a lot easier to use one of those than to keep switching chairs and asking if they see what we're talking about. So if anybody knows please send me the information so I can check into it further.
Thanks!
Paula = )
Paula Sicurello UC Berkeley Electron Microscope Lab psic-at-uclink4.berkeley.edu
} I am currently staining milk proteins with phosphotungtic acid (2%/water } ph 7.3) and uranyl acetate (1%/water) and often I do get what I called a } background consisting of very small (about 20-40nm) bubbles-like } structures that interfere with my samples... } } I also recall seeing the same thing with viruses but this time it is rather } annoying because it also looks a lot like my protein...
This annoying phenomenon, sometimes called "champagning", may be caused by several factors. In my own experience, I would see it whenever using PTA in preparations with high protein concentrations. If you are quick, you can actually see the phenomenon taking place as the beam vaporizes the PTA. I believe that the proteins serve to sequester water which is then enrobed in PTA. When the beam strikes the hydrated proteins, the water is vaporized and bursts through the PTA crust (like erupting gases in magma) leaving behind uniformly sized, spherical, electron light areas that may be mistaken for viruses or protein subunits. One workaround may be to dilute the protein, use phosphomolybdic or silicotungstic acids as the negative stain and to dry the grids in a 60 degree oven over the weekend.
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing Southern Illinois University Carbondale, IL 62901-4402 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
I am looking for a software package to construct and visualize an amorphous 3-D SiO2 network with options that allow for insertion of other elements such as nitrogen, carbon, and hydrogen and also the possibility of forming various molecular groups such as silanol, siloxane, etc..
Does anyone know if such a software package exists?
} I am looking for an electron diffraction analysis software package. I have } to identify some particles (wether they're cubic, tetragonal, etc) and } there must be an easier way than to hand draw the expected diffraction } patterns....Does anybody know of something useful on the Web?
Try the EMS On-line soft running at WWW URL http://cimewww.epfl.ch/. This is a short version of P. Stadelmann EMS programme.
Sara, You can build a library of possible structures (lattice parameters, space group and if known the atom position in the cell to account for kinematical extinctions). Then you enter two or three diffracted vectors (lengths on the pattern, angles, approximate camera length) and the programme tells you on which zone axis of which structure you took the diffraction pattern plus the exact camera length needed to do the fit. If there are some ambiguities, you can also view/print the computed pattern=8A and more!
Philippe-A. Buffat
__________________________________________________________________ Philippe Buffat Ecole Polytechnique Federale de Lausanne (EPFL) Centre Interdepartemental de Microscopie Electronique Address: EPFL-CIME, Batiment MX-C, CH-1015 Lausanne, Switzerland Phone: +41(21)693 29 83 Fax: +41(21)693 44 01 (Central European Time) E-mail: philippe.buffat-at-cime.uhd.epfl.ch, WWW URL http://cimewww.epfl.ch/ ______________________________ Eudora F2.1 ___________________________
At 08:49 10/07/1997 -0600, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Wolfram is a German word and it's a reference to the gray color of "Tungsten" similar to the color of a wolf. Tungsten is a swedish word and means heavy stone. We use the word "Tunstene" in France, Wolfram is an alternate word but not very common. Salutations. ========================================================== Jacky Larnould mailto:larnould-at-mnet.fr voice:33 (0)4 67 72 28 26 fax :33 (0)4 67 79 54 90
I came across some sort of porous superlattice silicon(I think) in my rearch, I would be happy if someone provides me some information on diffraction patterns of porous superlattice silicon.
Thanks in advance
S Suder
Suli Suder Joule Physics Lab University of Salford Salford M5 4WT United Kingdom Tel: 0161 745 5000 ext. 53264 Fax: 0161 745 5119 E-mail: s.suder-at-eee.salford.ac.uk
Mark et.al. - We have the confocal listserver linked in our comprehensive listing of microscopy sites. When you go to it you will get this notice:
"Season's Greetings Our server is down for repairing. We will gradually bring the web up in the coming week. Please visit us after the New Year. Have a happy holiday!"
I think they are a bit premature! Seasons greetings Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
_________________________ } Dear all, } Could someone please give me the address of the listserver for } confocal microscopy. } } Thanks a lot, } } Mark Munro } SAC Aberdeen } m.munro-at-ab.sac.ac.uk
Mark et.al. - We have the confocal listserver linked in our comprehensive listing of microscopy sites. When you go to it you will get this notice:
"Season's Greetings Our server is down for repairing. We will gradually bring the web up in the coming week. Please visit us after the New Year. Have a happy holiday!"
I think they are a bit premature! Seasons greetings Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
_________________________ } Dear all, } Could someone please give me the address of the listserver for } confocal microscopy. } } Thanks a lot, } } Mark Munro } SAC Aberdeen } m.munro-at-ab.sac.ac.uk
Jean and all: Here is a copy from our on-line catalogue:
"PARLODION Pyroxylin, in strips. A highly purified form of cellulose nitrate; prepared for embedding tissu= es for sectioning and for making support films for EM grids from 1% parlodio= n in Amyl Acetate."
I prefer parlodion over formvar because it seems to release better when cast on a microscope slide, also is seems more suitable for thicker film= s. It comes in hard strips. Cut and weigh a suitable piece and then calcula= te the solvent required to make the percentage solution. Because it is very flammable parlodion it is frequently stored under water. Blot, then dry = in an incubator before weighing the small piece. Parlodion, like formvar requires=20 a carbon coating for stability in TEM. Butvar does not.=20 ProSciTech and I trust all other EM suppliers handle Parlodion. =20 Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS =20 ************************ http://www.proscitech.com.au =20 ---------- } From: Leclerc Jean {leclercj-at-magellan.umontreal.ca} } Hi there!=88 } =20 } Anyone knows anything about pyroxylin? I would like to know what it is } exactly, and if it might be more stable than some other films for coati= ng } grids for the TEM. (Note that I'm looking at thick stuff - no thin } sections here!) } =20 } Thanks! } ---------- } =20
At 10:49 AM 7/10/97 -0400, Michael Delannoy wrote: I'm in the process of using the DGD resinless embedding technique developed by } Capco, Krochmalnic and Penman and I'm looking for: 1)Short cuts- } the ethanol, nbutanol to DGD transition is very long (6 hours) and I am working } with monolayers
{snip}
It is often possible to dramatically speed up the dehydration steps of TEM specimen preparation by carrying them out in a laboratory microwave. The technique is described on pp352-353 of _The Microwave Cookbook for Microscopists._
disclaimer: Energy Beam Sciences manufactures laboratory microwaves, and has a vested interest in seeing more people utilizing this technique.
Best regards, Steven E. Slap, Vice-President ******************************** Energy Beam Sciences, Inc. Adding Brilliance To Your Vision ebs-at-ebsciences.com http://www.ebsciences.com/ ********************************
O.K. you professionals here's a real puzzler for you:
I have a user here who has about 24 SPI Slide-A Grid Boxes. They were a little dusty (they may have been used before - heavens not to say they didn't come absolutely pristine from SPI), and so he throughly washed them and rinsed them with Ethanol, only now he can not get them back open any longer! They are really stuck.
I don't know if he removed some type of lubication on the box, maybe the "natural" lube from the manufacturing process, or if he caused the plastic to swell with the EtOH, but I could certianly use any suggestion. Maybe we should throw this up to the group - surely someone has cleaned grib boxes before.
We can get the box lids to move upto 5mm or so but then they stop and with work we can get them to move back, but we still have freed any lids. We've tried prying up the lids a little from the back edges (where they slide along rasied ridges) - didn't help much. We've even tried soaking them again in water or EtOH hoping to provide some "lubrication" (there are no grids in the boxes, but didn't really want to soak them in some thing oily) - still with no success.
I oringinally asked this of Charles Garber (of SPI) and neither of us have come up with any ideas, but he did throw in the following:
Of the box parts:
} The plastic is an antistatic formulation however, alcohol being so } polar is not going to swell the base piece at least. If the alcohol } was hot or if it was in contact with the plastic for some extended } period of time, then who knows, but I just can not believe that the } alcohol would have done anything to it in the way you are } suggesting. } } The sliding top might be another story. I forgot the plastic that is } used for it, but it might be a clear PVC. But even then, room } temperature alcohol should not have had time to do anything. }
Any suggestions?
The things fools will do ....
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 352 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513-529-5712 Fax: 513-529-4243 E-mail: edelmare-at-muohio.edu
"640K ought to be enough for anybody." -- Bill Gates, 1981
ebs-at-ebsciences.com wrote: } It is often possible to dramatically speed up the dehydration steps of TEM } specimen preparation by carrying them out in a laboratory microwave. The } technique is described on pp352-353 of _The Microwave Cookbook for } Microscopists._
Another way to speed up dehydration is to do it chemically - use acidified 2,2 dimethoxypropane (DMP). For a monlayer complete dehydration will occur in a few min. I usually dehydrate small blocks (up to 3x3x3 mm) of plant tissue for 15 min, and then go directly to embedding medium:dimethoxypropane 1:1. Any residual water from the dehydration step will be removed during the first embedding step. However, I do not know if DGD can be mixed or react with DMP, so you have to test it first.
Any reason why you wouldn't use an anti-vibration platform? We have a Micro-g isolation table made by Technical Manufacturing Corporation for our JEM 1010. We were in a room next to the furnace and air conditioners for the whole building and NEVER had any problems. If you are interested, I have the information, so should JEOL, since they both have offices in Peabody, Massachusetts.
Cheri Owen Detroit Neurotrauma Institute Wayne State University Detroit
Microscopy & Microanalysis '97 in Cleveland is less than a month away! The deadline for early registration is this coming Tuesday, July 15. If you have not yet done so, register now to qualify for the lower rates. If you need registration forms, please call the business office at (800) 538-3672 . If you have internet access to the www, forms are available for downloading at http://www.bright.net/~strecker/msno/mm97.html.
If you are planning on attending and have not yet reserved hotel space, you must contact the hotels directly. Hotel information is also available from our web site or from the business office. I can also e-mail you this information if required. At our last planning meeting it was announced that there are no rooms available downtown Cleveland for Saturday, August 9, so if you need a room, please act now.
A special note to those who have been following the Mars Pathfinder program. A debate on the pros and cons of life on Mars will take place on Monday, August 11. Following this will be a presentation by Al Worden, Apollo 15 astronaut. Mr. Worden's talk will feature many of his breath-taking photographs from space.
---------------------------------------------------------------- Dave Strecker mailto:dave.strecker-at-ab.com Rockwell Automation/Allen-Bradley Phone: (216)646-3250 Component Engineering ND246 Fax: (216)646-3416 1 Allen-Bradley Dr. Mayfield Hts., Ohio 44124 USA ----------------------------------------------------------------
Sorry Geoff! I guess I was forgetting that other people may want to know the details of this book, so I am posting the details here:
A beginners guide to the collection, isolation, cultivation and identification of Freshwater Protozoa
Pblished in 1988 by Culture Collection of Algae Associaction (CCAP)
Freshwater Biological Association The Ferry House Ambleside Cumbria United Kingdom
ISBN 1 871105 03 X
but remember it has drawings not photos! still for 4.40 uk pounds you can't complain :)
} -----Original Message----- } From: Geoff McAuliffe [SMTP:mcauliff-at-UMDNJ.EDU] } Sent: Friday, July 11, 1997 4:34 PM } To: Conrad Perfett } Subject: protozoa book?? } } Dear Conrad: } } You have been on the MSA list praising a book on microscopy of protozoa } quite often. So what is the title of the book??????? I keep reading your } posts hoping to find some mention of the title, author, publisher, } something! Either post the title or email it to me, please. } } Geoff } -- } *************************************************************** } Geoff McAuliffe, Ph.D. } Neuroscience and Cell Biology } Robert Wood Johnson Medical School } 675 Hoes Lane Piscataway, NJ 08854 } voice: (732)-235-4583; fax -4029 e-mail: mcauliff-at-umdnj.edu } ***************************************************************
Greetings, I received many replies to my original query, and in case others are interested, here is the gist of them. Thanks to all of you who answered.
Hi Tamara,
If you have a proper filter setup, the emission filter should be excluding all the exitation wavelength. Therefore I think you are looking at autofluorescence. Any light that has not changed wavelength should be blocked.
Bob
On Thu, 10 Jul 1997, Tamara Howard wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Disclaimer first: I'm not a chemistry/physics person, so please } excuse me if this question is incredibly stupid :) } Is it possible for a pigment to scatter/reflect light strongly } enough to look like fluorescence? If so, how could you distinguish such } behaviour from "real" fluorescence? I'm trying to work with some pigmented } plant samples that "glow" under my rhodamine/Tx Red excitation/emission } filters (standard fluorescence scope); the pigmented areas look normal } under FITC and UV conditions. The pigment itself is red under regular } light. We'd like to know if the pigment autofluoresces or if this is just } some light/pigment interaction. I'm lost and our library isn't heavy on } this type of references. Any of you "hard science" types willing to try to } educate me?! } } TIA! } } Tamara Howard } CSHL } howard-at-cshl.org } } } }
Does anyone know where I can get a used LogEtronics 55-EM enlarger (or something similar with auto-dodge & burn-in capabilities)in good working condition?
Thanks,
Michael Coviello Lab Manager The University of Texas -at- Arlington
Hello again, DGD stands for diethylene glycol disterarate, which is a waxy "solid" at room temp (although sweats/melts at r.t. like the London resins-white,gold) and liquid at 70 deg.C. The references are: Pennman (1995) PNAS. 92:5251-5257 review Capco etal (1984) JCB. 98:1878-1885 These will get you started. Right now I've run some samples on tissue culture plates, coverslips and a glass slide (which got trashed). Separating the DGD from the plates was easy (light hammer tapping) but some cells did remain on the dish. The coverslips separated easily, so I've mounted a few and will be cutting soon (I can't see the cells so I'm sectioning blindly). One problem is the block sweating at the cell surface, I've placed everything in the fridge hopefully to stop this-cutting should be interesting. The second paper used glass petris which I don't have, but may have to get. The n-butanol did not dissolve the culture plate. I'll keep you posted, more suggesstions are welcome.
Sweating in Baltimore, Mike D.
P.S. I got my DGD from Polysciences who also sent a protocol. This stuff may work better with tissue blocks and pellets, monolayers are tricky.
An excellent source for used photographic equipment of all kinds, including digital imaging, is a monthly magazine called "Shutterbug". I think it's put out by the same people who do the "Computer Shopper" magazine. They began as an advertising magazine dedicated to used photo stuff, but have expanded into a large format glossy, general photography magazine. They still have loads of ads and classifieds for all sorts of used you-name-it relating to photography. It can be found in many camera shops and magazine newstands.
Randy Tindall Center for Electron Microscopy Southern Illinois University at Carbondale
Hi, In 25 years of being in a microscopy laboratory I have witnessed impossibly sticky grid boxes 3 times. Twice they were manufactured by LKB, once they came from another source. Each time the grid boxes were washed in alcohol. No time were we able to rescue the boxes. We threw them away. We now strictly wash all grid boxes with soap and water only. So sorry, HHC
In Image-Pro for DOS, after you count and measure all the white objects, the total area of these objects can be found by going to View-Statistics. The Sum measurement is the total area of all counted objects.
Additionally, to clarify a few points, Image-Pro for DOS was not sold to anyone - Media Cybernetics - the original developer of the Image-Pro family of products - still owns all rights to these products. However, the DOS versions of Image-Pro were discontinued quite some time ago (nearly 4 years). Although we make our best effort to support all of our products and customers, there have been at least 6 new Windows versions published since the last DOS version, and the vast majority of our customers have taken advantage of the upgrades or extended maintenance plans in order to stay 'current'. If you have registered your product with us, then you have probably seen the announcements and upgrade offers over the years.
If you, or any other DOS users out there, would like to upgrade to the latest version - Image-Pro Plus 3.0, please feel free to contact me directly.
-------------------------------------- Scott D. Ireland Regional Sales Manager Eastern North America & Latin America Media Cybernetics, LP "The Imaging Experts" Tel: (716) 473-0222 Fax: (716) 473-8048 scott-at-mediacy.com http://www.mediacy.com ---------------------------------------
---------- } From: wise-at-vaxa.cis.uwosh.edu } To: Microscopy-at-sparc5.microscopy.com } Subject: Image Pro Plus } Date: Thursday, July 10, 1997 9:21 AM } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } To all, } } We have an older, DOS-based version of Image Pro Plus image } analysis software that has us stymied. We are capturing video images of } chloroplast movements and dumping them into a DOS-based computer. We want } to be able to sum all of the white areas in a field and then express that } total white area as a percentage of the total image area. It seems simple } enough, but it's not. We can identify all of the "white" objects, we can } define what we are calling 'white', we can get the area of each individual } white object, but we cannot sum the areas of all of the white objects. } Image Pro Plus corporation is no help. They sold the rights to the DOS } version to another company and no one a IPP knows anything about it. They } are quite willing to sell us the Windows version for $3500 but cannot help } us with their older version. } Does anyone out there in microscope land have experience with this } program? } } Yours in computer frustration, } } Bob } }
} } Does anyone know a TEM protocol for "Tungsten Shadowing" say, for DNA } strand enhancement? } } Thanks! } } Cheri Owen } Detroit Neurotrauma Institute } Wayne State University } Detroit
Cheri, There are two sections on tungsten shadowing in Chap 7 " High Resolution Shadowing" by Henry S. Slayter in the book: Electron Microscopy in Biology, J.R. Harris, ed. 1991, IRL Press at Oxford Univ. Press. ISBN 0-19-963215-4 (pbk)
cheers ed
Edward J. Basgall, PhD The Pennsylvania State University Surface Chemistry Group ejb11-at-psu.edu Materials Research Institute Building Ph: 814-865-0493 University Park, PA 16802-7003 FAX: 814-863-0618
Often in our lab we are in a big rush to get prints right after the user has finished using the EM. I've sometimes just washed the negatives (3.25X4" Kodak EM film) just 2 minutes instead of 20, and then after I've printed them all, I go back and wash them for 30 minutes to remove any trace fixative that might still be remaining in the emulsion. This way I can shave 18 minutes off of the delivery time. I also print them with just a bit of water at their edges so that they are not completely dry, and shave another 5 minutes off of the drying time, so that they are 23 minutes out the door faster.
Does anyone else have any long term experience with this wash-after-wash technique. I'd like to know more about the long term effect, i.e.: whether I can indeed wash them later and be sure to remove all the traces of fix from the film emulsion.
} However, the DOS } versions of Image-Pro were discontinued quite some time ago (nearly 4 } years). Although we make our best effort to support all of our products } and customers, there have been at least 6 new Windows versions published } since the last DOS version, and the vast majority of our customers have } taken advantage of the upgrades or extended maintenance plans in order to } stay 'current'.
There is a difference in perspective about these issues for many of us.
'Nearly 4 years' is not a long time for the life of a useful product after discontinuation and, in fact, I am often loath to upgrade software if the current version is working despite the 'enhancements' of newer versions. For example, previous correspondance has revealed that IP has changed some of the ways in which it measures certain parameters and I am concerned about data compatibility. In addition, interface changes are tedious even if the newer interface is better; one has to unlearn too many things.
So, your statement that IPWin has had at least 6 versions since the last DOS version (which I have) is an indication of progress but it is also a matter of great concern about support for these lapsed versions.
} Often in our lab we are in a big rush to get prints right after the user } has finished using the EM. I've sometimes just washed the negatives } (3.25X4" Kodak EM film) just 2 minutes instead of 20, and then after } I've printed them all, I go back and wash them for 30 minutes to remove } any trace fixative that might still be remaining in the emulsion. This } way I can shave 18 minutes off of the delivery time. I also print them } with just a bit of water at their edges so that they are not completely } dry, and shave another 5 minutes off of the drying time, so that they } are 23 minutes out the door faster. } } Does anyone else have any long term experience with this wash-after-wash } technique. I'd like to know more about the long term effect, i.e.: } whether I can indeed wash them later and be sure to remove all the } traces of fix from the film emulsion. } Dear Garry, We don't use the wash-after-wash technique; rather, we wash for ~1 min in H2O, 1 min in Permawash and ~1 min in H20. This removes the fixer and negs which have been processed this way keep for several years at least. I guess if a user were *really* in a hurry, we could save ~1 min, but our way the user can take the negs home. ;-) Yours, Bill Tivol
} } } Often in our lab we are in a big rush to get prints right after the user } } has finished using the EM. I've sometimes just washed the negatives } } (3.25X4" Kodak EM film) just 2 minutes instead of 20, and then after } } I've printed them all, I go back and wash them for 30 minutes to remove } } any trace fixative that might still be remaining in the emulsion. This } } way I can shave 18 minutes off of the delivery time. I also print them } } with just a bit of water at their edges so that they are not completely } } dry, and shave another 5 minutes off of the drying time, so that they } } are 23 minutes out the door faster. } } } } Does anyone else have any long term experience with this wash-after-wash } } technique. I'd like to know more about the long term effect, i.e.: } } whether I can indeed wash them later and be sure to remove all the } } traces of fix from the film emulsion. } } } Dear Garry, } We don't use the wash-after-wash technique; rather, we wash for } ~1 min in H2O, 1 min in Permawash and ~1 min in H20. This removes the } fixer and negs which have been processed this way keep for several } years at least. I guess if a user were *really* in a hurry, we could } save ~1 min, but our way the user can take the negs home. ;-) } Yours, } Bill Tivol
Dr. Lawrence F. Allard Senior Research Staff Member High Temperature Materials Laboratory Oak Ridge National Laboratory 1 Bethel Valley Road Bldg. 4515, MS 6064 PO Box 2008 Oak Ridge, TN 37831-6064
Please recommend, if it's possible, suitable, detailed references or your own procedure, how to isolate leukocyte-rich plasma and process it without sedimentation media, for electron microscopy.
Thanks for your understanding, Margarita Chrysanthou E-mail : cozzika-at-athena.copmulink.gr
The Department of Biological Sciences of the Exacts and Natural Sciences Faculty of the Buenos Aires University are looking for donation of a TEM and-or SEM equipment, not older than 15 years and still woorking. We will paid any handling and shipping cost. It would be very helpfull for us any other kind of assistance. Thank you vey much in advance.
Please respond to me at the addresses and numbers listed below.
Gabriel Adriano Rosa Area Microscopia Electronica, Depto. Cs. Biologicas Fac. de Ciencias Exactas y Naturales, Universidad de Buenos Aires Ciudad Universitaria, 4 piso, Pab. II, CP 1428, Buenos Aires, ARGENTINA
Cheri and all - Specific to DNA strands, I know this as the Kleinschmitt technique, now over 25 years old. The osmotically opened DNA strands are rotary shadowed. For that the specimen grids are rotated at about 60 rpm, 60-100mm from the source and at an 6-10 degree angle of incidence. The rotary shadowing makes it easier to check the continuity and to follow the under/over of the fibers. I used to evaporate Pt/Pd or for finer grain, Pt and C simultaneously. Ed Basgall's posting should help with the tungsten evaporation. Tungsten is finer still but really requires special equipment. It is possible to evaporate W - "Wolfram" in a normal evaporator, but it is slow and a lot of heat is generated. A heat shield for the specimen with a suitable aperture is advised. But why that trouble? The coarser grain from other evaporation material does not impair tracing and measuring of the DNA fibres and fibre detail is better using negative staining, or positive staining using methanolic UA. Regards Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
---------- } From: Cheri Owen {cowen-at-cmb.biosci.wayne.edu} } To: Microscopy-at-sparc5.microscopy.com } Subject: Tungsten shadowing } Date: Friday, 11 July 1997 23:33 } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } Does anyone know a TEM protocol for "Tungsten Shadowing" say, for DNA } strand enhancement? } } Thanks! } } Cheri Owen } Detroit Neurotrauma Institute } Wayne State University } Detroit }
A Research Associate position is open in the dDpartment of Radiology, University of Texas Health Science Center at San Antonio, as the primary technician for the operation of a new atomic force microscope (Nanoscope, Digital Instruments). The person in this position would participate in studies of novel vascular biomaterial surfaces as well as in studies of vascular cell responses to mechanical forces. Applicants should have a Master's degree or a bachelor's degree with several years experience . Preferred areas of training and/or experience include scanning electron microscopy, image analysis, and cell biology. The position is currently open and we seek to hire someone as soon as possible. All applicants must apply through Human Resources, UTHSCSA. Inquiries should be directed to Dr. Gene Sprague, 210 567-5564, e-mail sprague-at-uthscsa.edu
We have used the Struers Tenupols here for many years and are generally happy with them. They wear out after a while because of all the nasty electropolishing solutions that people want to use in them but they seem to be good for at least 3-5 years (probably a lot longer if they did't get the battering that our students and RFs give them). Even then, the first things to go are the jets and these can be replaced (at a price).
You do (at least with tenupols) have to think a bit about how you will cool your electrolytes. We have a system where alcohol cooled with either dry ice or liquid nitrogen is pumped through the cooling coils on the tenupol. The temperature of the electrolyte is controlled (to give between 0 and -40 degrees C) by a homebuilt control box using the reading from a electrical resistance thermometer to control a relay that turns the pumping of the cooling alcohol on or off.
Hope this helps
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ Ian MacLaren, Tel: (44) (0) 121 414 3447 IRC in Materials for FAX: (44) (0) 121 414 3441 High Performance Applications, email: I.MacLaren-at-bham.ac.uk The University of Birmingham, http://web.bham.ac.uk/I.MacLaren/ Birmingham B15 2TT, England. ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
When I had worked as a newspaper photographer back in my college days, time was always something that we were trying to cheat on. We used the EtOH wash technique to get things to dry faster.
One thing that has not been addressed is what are the photos going to be used for. Previously a typical morning at the EM would produce 80 to 100 films. We would print 8X10 images of these photos. These would later be used to keep track of the films and to keep track of the portions that would were later digitized. The printing to RC paper and processing with an Ilford developer went fairly quickly. I would hate to think of how much time it would take to send 100 several meg images to the Fugi Pixtograph printer. First there would be the storage problem, the output problem, lower resolution, and then the cost.
Dana noji-at-snowman.med.umn.edu __~0_\___ Minneapolis, Minnesota (_________) (612) 624-4687 O O http://www.tc.umn.edu/nlhome/g118/post-doc/dtn.htm
Margarita: One can simply make a buffy coat by spinning the sample in a wintrobe tube ( a skinny glass tube obtained from hematology) immediately and fixing the buffy coat by aspirating off most of the serum and replacing with glutaraldehyde/paraformaldehyde EM fixative. then cut the top and bottom off the wintrobe tube (after diamond scoring) and float this half inch long segment in the fixative until the buffy starts to get firm. Simply take a wooden applicator stick and poke the pellet out of the tube segment, process for EM and embed on edge to see all the leukocyte types in layers. bob M
We have a Fishione unit which we have used for the past fifteen years. We found the unit to be very reliable. Our unit came with a glass dish used as the reservoir. This tended to affect the response of the light detector used to stop the electropolishing when a hole appeared. In most instances however this was not a severe limitation for us. The jets and the specimen holder do need to be aligned occasionally. Also, we did have to replace the jets and the holder once since the electrodes do have a finite life. As long as the unit is rinsed thoroughly after use it should last a long time. We found the unit to be very compact and it was a simple matter to cool the electrolyte using a double beaker with the larger beaker containing the coolant. Please not that I am not familiar with the newer units and that configuration might have changed by now.
I need your suggestions on the twin-jet electropolishers for preparation of TEM samples. I need a reliable brand with reasonable price. Right now, I have information from Struers,SouthBay Technology and Fischione. Fischione has a reasonable price. Does anyone have any experience with Fischione?
Thank you for your all suggestions. Please reply my own adress.
Ibrahim Karaman Mechanical Engineering Department University of Illinois 1206 W. Green Street Urbana, IL 61801
I've never heard about Permawash. I've heard of "hypoclear" from Kodak, but the instructions state that this should be discarded after 24 hours, so it would be expensive and time consuming to make this up all the time. Is this Permawash stuff stable enough that I could keep it in a stainless steel covered tank for a week or so, to use as required?
Garry
} } Dear Garry, } } We don't use the wash-after-wash technique; rather, we wash for } } ~1 min in H2O, 1 min in Permawash and ~1 min in H20. This removes the } } fixer and negs which have been processed this way keep for several } } years at least. I guess if a user were *really* in a hurry, we could } } save ~1 min, but our way the user can take the negs home. ;-) } } Yours, } } Bill Tivol } }
It would have been my thoughts that the film need not be washed for such a long period of time, but I'm just trying to follow the instructions as supplied by Kodak as a film insert. Somehow they felt that 20 minutes was the minimum time required unless one used hypoclear or Permawash.
Garry
} Garry, } } It's probably not necessary to use the wash-after-wash technique. William } Tivol's reply probably provides the best compromise. Film, as opposed to a } print on paper, is not really very absorbent and can be washed pretty clean } pretty fast. Fix tends to rinse out readily. (I repeat, this is NOT } necessarily true for resin-coated prints, although they also wash pretty } fast, and is CERTAINLY not true for fiber-based prints.) } } As an example, in their IlfoPro newsletter, Ilford recently recommended a } wash technique for their 35mm and roll films that involved filling the } developing tank with water and inverting it 5 times, refilling it and } inverting it 10 times, and filling it a third time and inverting it 20 } times. They claim this is an archival wash technique. } } After only a couple minutes wash, I'd bet that your negatives will show no } signs of change for many years. (I can already hear the screams of } outrage!) Hypo clearing agents, PermaWash, etc., are excellent for } assuring that the negatives will outlive the people taking them, and I } certainly recommend their use for safety's sake. I still have negatives, } however, that were taken when I was 12 years old and processed in a } makeshift darkroom (it required nighttime in the country to make it } light-tight, IF the moon wasn't too bright). They were washed pretty } sloppily and still show no signs of deterioration 30 years later. } } Regarding digital imaging---in my opinion, silver-based photography is } still the most cost-effective, highest information content, easily stored } method for image CAPTURE. After capture, however, digital is more and more } the way to go for manipulation and publication. Nothing yet beats a } negative as an imaging starting point, especially considering the sizeable } investment in decent digital imaging equipment. Most labs already have } darkrooms. (I bet this will generate some comments!) } } Hope this helps. } } Randy Tindall } Center for Electron Microscopy } Southern Illinois University at Carbondale } } }
We are doing diagnostic electron microscopy as part of the Department of Pathology, in a major health care centre. The users that demand fast turn around time are the Pathologists especially when we are processing a fine needle aspirate biopsy sample. Our technique spares the patient from more invasive procedures. Because of our speed, we have been able to deliver the micrographs within 8 working hours from fresh tissue, to dispell the myth that EM as applied to biological tissue is necessarily very very slow. Often we get micrographs after 6 working hours or less. So, speed is of the essence for us. (at least that is what they tell us)
Garry
} ---------- } From: Brian G. Demczyk[SMTP:demczyk-at-erxindy.rl.plh.af.mil] } Sent: 13 July, 1997 06:35 } To: Garry Burgess } Subject: Re: RUSH PRINTS! Rush Lab. } } Just out of curiosity, what industry are you in that users } demand such a rapid turn around?! }
Sometimes when I want to make a transparency for seminars, or workshops or whatever, I put the EM negative into the enlarger and project it to a small 2X2" size on to yet another piece of EM film that I've cut in half for this purpose. (you need a special enlarger lens to get this small!). Then I cut it to size and mount it in a glass slide mount, and voila, I have supersize black and white slides, with good resolution and contrast. (at least better than 35mm projection slides) But sometimes in the past, if I tried to rush things, I notice that the image turned brown. To fix this problem though, I simply re-fix and re-wash this image, and the brown discoloration (which is probably some residual silver halide and fix) is removed, and all is well again.
Garry
} ---------- } From: rtind-at-siu.edu[SMTP:rtind-at-siu.edu] } Sent: 14 July, 1997 14:51 } To: Garry Burgess } Subject: RE: Rush Lab } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
from what i understand, Permawash is just soapy water. i simply use a mild detergent solution instead and it works fine
} I've never heard about Permawash. I've heard of "hypoclear" from Kodak, } but the instructions state that this should be discarded after 24 hours, } so it would be expensive and time consuming to make this up all the } time. Is this Permawash stuff stable enough that I could keep it in a } stainless steel covered tank for a week or so, to use as required?
} } } We don't use the wash-after-wash technique; rather, we wash for } } } ~1 min in H2O, 1 min in Permawash and ~1 min in H20. This removes the } } } fixer and negs which have been processed this way keep for several } } } years at least. I guess if a user were *really* in a hurry, we could } } } save ~1 min, but our way the user can take the negs home. ;-)
Evans blue is commonly used as a counterstain for flourescence work. It stains the specimen for observation by white light, and since it has no autoflourescence, it is invisible under the UV wavelengths.
For this application, a bottle should last an academic lifetime...I can't imagine what a case would be used for (except maybe spending surplus year-end funds)
-=W.L. Steffens=- Department of Veterinary Pathology College of Veterinary Medicine University of Georgia Athens, GA 30602 USA http://www.vet.uga.edu/vpp/wls/steffens.html
The photos are going to be used by Pathologists in a hospital for diagnostic purposes.
Garry } } One thing that has not been addressed is what are the photos going to be } used for. Previously a typical morning at the EM would produce 80 to 100 } films. We would print 8X10 images of these photos. These would later be } used to keep track of the films and to keep track of the portions that } would were later digitized. The printing to RC paper and processing with } an Ilford developer went fairly quickly. I would hate to think of how } much time it would take to send 100 several meg images to the Fugi } Pixtograph printer. First there would be the storage problem, the output } problem, lower resolution, and then the cost. } } Dana } noji-at-snowman.med.umn.edu __~0_\___ } Minneapolis, Minnesota (_________) } (612) 624-4687 O O } http://www.tc.umn.edu/nlhome/g118/post-doc/dtn.htm } } }
} I've never heard about Permawash. I've heard of "hypoclear" from Kodak, } but the instructions state that this should be discarded after 24 hours, } so it would be expensive and time consuming to make this up all the } time. Is this Permawash stuff stable enough that I could keep it in a } stainless steel covered tank for a week or so, to use as required? } Dear Garry, The bottle of Permawash says that working solution will oxidise in an open tank/tray in 6-8 hrs. A tank with a floating lid will keep for 30 days and a stoppered bottle for 90 days. The undiluted stock solution will keep a lot longer. We have our working solution in a SS covered tank like yours; I'm confident that it keeps easily for the week or so you need. Yours, Bill Tivol
Sorry to do this to you, but there was a discussion a while back about inexpensive light microscopes for children. In particular there was a recommendation from someone in the Seattle area about a $99 model (Tasco brand?). Of course, I deleted the messages but was recently asked by a friend for a recommendation. I just about went blind looking (unsuccesfully) through the June archive so it must have been further back then that! To save my eyes from further abuse, could the person posting about that microscope please just e-mail me the company name and address? I would greatly appreciate it.
Oh, also any recommendations for things to look at for kids ages 6-10 would be appreciated.
Thanks in advance.
Cheers,
John Vetrano Pacific Northwest National Laboratory P.O. Box 999 Richland, WA 99352
I need your suggestions on the twin-jet electropolishers for preparation of TEM samples. I need a reliable brand with reasonable price. Right now, I have information from Struers,SouthBay Technology and Fischione. Fischione has a reasonable price. Does anyone have any experience with Fischione?
Thank you for your all suggestions. Please reply my own adress.
Ibrahim Karaman Mechanical Engineering Department University of Illinois 1206 W. Green Street Urbana, IL 61801
We have just received an enquiry about SEM analysis for: a) pigments particles dispersed in mineral oils b) pigments particles in oil-water emulsions
Does any of you have any experience in that subject? Unfortunately, we do not count with equipment for sample preparation other than the sputter-coater and carbon evaporator.
Any help will be very welcome.
Thanks in advance,
Nora Pratta Centro Regional de Investigacion y Desarrollo Santa Fe - Argentina
} } A *very* good argument for Digital Imaging! } } Larry
Yes! but at what cost! with dwindling funds and tighter budgets all arguments are in vain......... Neelima Shah Regards... :-) :-) :-) :-) :-) :-) :-) :-) :-) :-) ;-) ;-) Visit us at http://www.med.upenn.edu/~path/core/EMCMAIN1.htm
----------------snip---------------} } 3] Is there any fixative other than Osmium which would best retain phenolics } in small root explants? } } I am not using a cryoprotectant like DMSO because I want to avoid soaking the
} roots in it, as this could allow phenolics to diffuse from the plant cells. } However, small root explants are soaked in Os-KI for several hours under } vacuum, to overnight at 4 C prior to microtomy. This allows for Os } penetration. I am relying on Os to "fix" the phenolics. Perhaps there is } something else which would penetrate more rapidly and fix or condense the } phenolics. } } Mahalo = Hawaiian for Thanks !!!!!!!! }
Dave,
have you tried RuO4? I have found it works well with a number of unsaturated ring structure compounds as a fixative. I make it up from the chloride just before use and it seems to act quite rapidly.
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Dear Conrad } } } well I now know why I am having difficulty finding the Amoeba since } } according to the book I have just bought, at least the classic Amoeba } } Proteus is rare in the wild!!! I remember being told by a retired } } microscopist that it was rare also. } I did send a previous reply to the list. Since I got no copy I have to assume it is lost? Depending on what you see as "wild", "nature" and "natural", Amoeba is easy to find in large concentrations. At water refinery plants they are in high concentrations in the activated sludge. There is normally a buildup of material on the edge above water level, different species and even "waterbears" can be found. (Remember to use gloves collecting and handling samples.)
## [########] ## ## ## ## ## Stephan H Coeztee Electron Microscope Unit Private Bag 3 Wits 2050 South Africa
Stephan-at-Gecko.biol.WITS.ac.za
Tell: +27 11 716 2419 Fax : +27 11 339 3407
} ## [########] ## ## ## ## ## Stephan H Coeztee Electron Microscope Unit Private Bag 3 Wits 2050 South Africa
If time is of the essence have you considered doing a final wash in ethanol (70% or thereabouts)? It must be clean of course but you should knock a few more drying minutes off. Try it on something unimportant first though.
Malcolm Haswell e.m. unit University of Sunderland UK ----------
Often in our lab we are in a big rush to get prints right after the user has finished using the EM. I've sometimes just washed the negatives (3.25X4" Kodak EM film) just 2 minutes instead of 20, and then after I've printed them all, I go back and wash them for 30 minutes to remove any trace fixative that might still be remaining in the emulsion. This way I can shave 18 minutes off of the delivery time. I also print them with just a bit of water at their edges so that they are not completely dry, and shave another 5 minutes off of the drying time, so that they are 23 minutes out the door faster.
Does anyone else have any long term experience with this wash-after-wash technique. I'd like to know more about the long term effect, i.e.: whether I can indeed wash them later and be sure to remove all the traces of fix from the film emulsion.
We have been using two methods (Fe Acetate & Osmium KI) to stain for phenolics in thick (30-60u) CRYOsections of plant roots. We checked these methods on coffee leaves which have a lot of phenolics, and while both worked, the Os-KI method was superior and more specific in terms of the literature. In our research specimens (roots of alfalfa) neither of these works very well by itself. However, we get a strong staining reaction from Os-KI treated specimens if we add Toluidine Blue just prior to observation. This reveals cells with large dark deposits in their vacuoles. These could be phenolics but they could be something else. We are planning to screen several more protocols, as it would be encouraging to have several methods which yield approximately the same results, because the specificity of histochemical methods is often not understood. I have the following questions.
1] Has anyone used Toluidine Blue in consort with other treatments (especially Os-KI) to localize phenolics? In other words is this staining reaction indicative of the presence of phenolics?
2] Are there any other histochemical procedures which are known to be specific for phenolics that may be appropriate for frozen plant sections?
3] Is there any fixative other than Osmium which would best retain phenolics in small root explants?
I am not using a cryoprotectant like DMSO because I want to avoid soaking the roots in it, as this could allow phenolics to diffuse from the plant cells. However, small root explants are soaked in Os-KI for several hours under vacuum, to overnight at 4 C prior to microtomy. This allows for Os penetration. I am relying on Os to "fix" the phenolics. Perhaps there is something else which would penetrate more rapidly and fix or condense the phenolics.
We have been using two methods (Fe Acetate & Osmium KI) to stain for phenolics in thick (30-60u) CRYOsections of plant roots. We checked these methods on coffee leaves which have a lot of phenolics, and while both worked, the Os-KI method was superior and more specific in terms of the literature. In our research specimens (roots of alfalfa) neither of these works very well by itself. However, we get a strong staining reaction from Os-KI treated specimens if we add Toluidine Blue just prior to observation. This reveals cells with large dark deposits in their vacuoles. These could be phenolics but they could be something else. We are planning to screen several more protocols, as it would be encouraging to have several methods which yield approximately the same results, because the specificity of histochemical methods is often not understood. I have the following questions.
1] Has anyone used Toluidine Blue in consort with other treatments (especially Os-KI) to localize phenolics? In other words is this staining reaction indicative of the presence of phenolics?
2] Are there any other histochemical procedures which are known to be specific for phenolics that may be appropriate for frozen plant sections?
3] Is there any fixative other than Osmium which would best retain phenolics in small root explants?
I am not using a cryoprotectant like DMSO because I want to avoid soaking the roots in it, as this could allow phenolics to diffuse from the plant cells. However, small root explants are soaked in Os-KI for several hours under vacuum, to overnight at 4 C prior to microtomy. This allows for Os penetration. I am relying on Os to "fix" the phenolics. Perhaps there is something else which would penetrate more rapidly and fix or condense the phenolics.
It's probably not necessary to use the wash-after-wash technique. William Tivol's reply probably provides the best compromise. Film, as opposed to a print on paper, is not really very absorbent and can be washed pretty clean pretty fast. Fix tends to rinse out readily. (I repeat, this is NOT necessarily true for resin-coated prints, although they also wash pretty fast, and is CERTAINLY not true for fiber-based prints.)
As an example, in their IlfoPro newsletter, Ilford recently recommended a wash technique for their 35mm and roll films that involved filling the developing tank with water and inverting it 5 times, refilling it and inverting it 10 times, and filling it a third time and inverting it 20 times. They claim this is an archival wash technique.
After only a couple minutes wash, I'd bet that your negatives will show no signs of change for many years. (I can already hear the screams of outrage!) Hypo clearing agents, PermaWash, etc., are excellent for assuring that the negatives will outlive the people taking them, and I certainly recommend their use for safety's sake. I still have negatives, however, that were taken when I was 12 years old and processed in a makeshift darkroom (it required nighttime in the country to make it light-tight, IF the moon wasn't too bright). They were washed pretty sloppily and still show no signs of deterioration 30 years later.
Regarding digital imaging---in my opinion, silver-based photography is still the most cost-effective, highest information content, easily stored method for image CAPTURE. After capture, however, digital is more and more the way to go for manipulation and publication. Nothing yet beats a negative as an imaging starting point, especially considering the sizeable investment in decent digital imaging equipment. Most labs already have darkrooms. (I bet this will generate some comments!)
Hope this helps.
Randy Tindall Center for Electron Microscopy Southern Illinois University at Carbondale
Herbert McLean Evans was chair of Anatomy at Univ of Calif Berkeley, from 1915. His early research included use of vital dyes for studies on blood volume, macrophages, ovarian estrous cycle, etc. In a 1920 paper (Dawson, Evans, Whipple. "Blood volume studies. III. Behavior of large series of dyes introduced into the circulating blood." Amer J Physiol 51:232-256) it was shown that a blue azo dye (T-1824) was "slightly superior" for blood volume measurement than the vital red series previously used. The dye came to be know as "Evans blue." Criteria that were important were: (1) non-toxic, (2) not stored in tissue, (3) color allows accurate determination, (4) removed quite slowly from the blood stream.
An example of more recent use: Bergh, Damber, 1988, Int J Androl 11:449.
Kent (A. Kent Christensen, Dept of Anatomy and Cell Biology, University of Michigan Medical School), {akc-at-umich.edu} )
------------------------------
On Sun, 13 Jul 1997, Gilbert T. Groehn wrote:
} Our lab acquired a case of EVANS BLUE stain } from one of our clients and we can not locate } this in any of our chemistry manuals. } } What are the normal uses for this stain (eg. } histology, hematology, etc.) ? We would be } using it only for light microscopy. } } What is the standard mixing ratios and } formulas for various applications? } } Any help on this item would be most appreciated. } } TIA } Gil Groehn } ULTRAMED, INC. } } ============================================================= } ULTRAMED, INC. Research Div. ad408-at-detroit.freenet.org } Grosse Pointe Farms, MICH 48236 USA Biomedical Consultants } =============================================================== }
We use the spot mode pretty routinely to identify inorganic particles in our polyester films. The size of these particles is anywhere from about 0.5 =B5m to 10 =B5m. So far it has worked quite well for us. When we = switch back to full scan after using the spot mode, we invariably see a "burn" mark which is exactly where we think it ought to be. We often look at a 2nd spot away from the structure that interests us to be sure that we're not just picking up a background.
} I travel the world teaching practical electron microscopy and it = worries me } the emphasis that people place on "Spot Mode". I do not teach the use = of } spot mode for the following reasons-
} 1. Just because the spot appears in a certain position on the = screen } is this its correct position. I have found machines many microns out = of } step in X and Y directions.
} 2. Specimens charge and switching out of spot mode does not give = the } operator an easy opportunity to see if the spot had moved during the } analysis.
} 3. Spot mode gives a false sense of analytical volume, no matter = how } small that spot may be we are almost certainly evaluating microns of } material.
} Do others worry about spot mode accuracy, do others test the spot mode } accuracy? Is it not better to simply increase the magnification = watching } the area of interest all the way up before the analysis and all the way } down after the analysis?
Youre vibration problem resembles a problem I had about 7 years ago when I was working with scanning tunneling microscopy at the university of Nijmegen (The Netherlands). Our STM in a UHV system was on the 4th floor, so we had some trouble with vibrations, mainly stemming from air ventilation compressors in the base of the building. In the same basement there were three vibration free sites consisting of concrete bloks (2x2x2 m) placed in sand an having no connection to the foundation of the building. We measured the vibrations on these bloks and found that they were only a little bit better than on the 4th floor!! We found out that the vibrations from the compressors propagates not only through the building construction but also through the ground or sand. So the effect of de-coupling the concrete blocks from the building foundation did not eliminate the vibration-coupling, but it did reduce the (effective) mass of the blocks. Hence the vibration amplitude growths! The foundation of the building, although directly coupled to the floors of the compressors, showed a quit clean vibration spectrum, because the mass is high (effectively the whole building).
So, back to your question: cutting the floor around your microscope will only help if this floor is the only vibration coupling to the microscope. If also the ground (dirt) or air (acoustic) couples vibrations to your microscope, the end result will be worse, since the mass of your system is reduced.
Success, Bart
Bart Nelissen
Akzo Nobel Central Research
Dept of Applied Physics, Microscopy
Tel: +31 26 366 1371
Fax: +31 26 366 5272
eMail: Bart.B.J.Nelissen-at-Akzo.nl
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to respond to this message (the sender-address is distorted by our 'gate-keeper').
I remember many years ago trying to clean some grid boxes (the white LKB type with a clear lid) with ethanol and it all went disastrously wrong. I can't remember whether both types of plastic deteriorated but the boxes were unusable. It may be that SPI boxes are made of similar material - try asking them. I now just give boxes a wash with soapy water (maybe a quick ultrasonic clean) and rinse in distilled water then dry for a couple of days I don't know if this helps.
Malcolm Haswell e.m. unit University of Sunderland UK ----------
O.K. you professionals here's a real puzzler for you:
I have a user here who has about 24 SPI Slide-A Grid Boxes. They were a little dusty (they may have been used before - heavens not to say they didn't come absolutely pristine from SPI), and so he throughly washed them and rinsed them with Ethanol, only now he can not get them back open any longer! They are really stuck.
I don't know if he removed some type of lubication on the box, maybe the "natural" lube from the manufacturing process, or if he caused the plastic to swell with the EtOH, but I could certianly use any suggestion. Maybe we should throw this up to the group - surely someone has cleaned grib boxes before.
We can get the box lids to move upto 5mm or so but then they stop and with work we can get them to move back, but we still have freed any lids. We've tried prying up the lids a little from the back edges (where they slide along rasied ridges) - didn't help much. We've even tried soaking them again in water or EtOH hoping to provide some "lubrication" (there are no grids in the boxes, but didn't really want to soak them in some thing oily) - still with no success.
I oringinally asked this of Charles Garber (of SPI) and neither of us have come up with any ideas, but he did throw in the following:
Of the box parts:
} The plastic is an antistatic formulation however, alcohol being so } polar is not going to swell the base piece at least. If the alcohol } was hot or if it was in contact with the plastic for some extended } period of time, then who knows, but I just can not believe that the } alcohol would have done anything to it in the way you are } suggesting. } } The sliding top might be another story. I forgot the plastic that is } used for it, but it might be a clear PVC. But even then, room } temperature alcohol should not have had time to do anything. }
Any suggestions?
The things fools will do ....
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 352 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513-529-5712 Fax: 513-529-4243 E-mail: edelmare-at-muohio.edu
"640K ought to be enough for anybody." -- Bill Gates, 1981
} } Can any of our German colleagues help with translation please? Came } across the term PHOSPHORWOLFRAM acid in a technical paper. Suspect it is } phosphoric acid but want to be sure before proceeding. } Thanks in advance } Ronnie Houston I think you will find that it is phosphotungstic acid (hence the atomic symbol W for tungsten)
Chris
Chris Gilpin Biological Sciences Electron Microscope Unit G452 Stopford Building Oxford Road Manchester M13 9PT phone +44 161 275 5170 fax +44 161 275 5171 http://www.biomed.man.ac.uk/biology/emunit/emhome.html
I have a client who will be purifying a retrovirus using a sucrose gradient.
The question is what medium should the sample be in to do the negative stain. Sucrose? Buffer? If anyone has worked with retrovirus I would like to hear your suggestion.
Thank you in advance
Karen Vaughn,EM Technician phone: (352)392-1184 University of Florida fax: (352)846-0251 Electron Microscopy Core Lab. e-mail: klv-at-biotech.ufl.edu 214 Bartram Hall web: http://www.biotech.ufl.edu/~emcl/ Gainesville, Fl. 32611
I'm not sure what is causing the brown staining, especially since you indicate that it's temporary. My GUESS is that it's inadequate fixing time or exhausted fixer. What are your fixing times?
Randy,
Usually I fix for 2-3 minutes, but in this case, it might be more, since the slides were actually just piled up in a staining beaker filled with hypo. Since the slides were in contact with each other, I'm quite sure that this explains the inadequate fixation. Actually usually I look at them under the safelight to ensure that they are adequately fixed, but sometimes, when I had a huge volume, then I must have missed a couple, because I had so many. (about 200 or so)
I use some sort of Kodak fixer, but I can't remember the name right at this second. I usually do not use hypochek, because as I've said, usually it's obvious when the fixer is losing it's strength, because it starts to take significantly longer than 2 minutes to fix our EM negatives. Because we work under safelight, this is something that can be checked each and every time that we develop film.
What fix do you use? Do you check your fix for exhaustion with HypoChek or a similar product that gives you a white precipitate when the fixer is saturated with silver compounds? Interesting.
I agree with you about the confusion about Permawash. It would have to be something more chemically like Hypo Clear than soap, because I can't imagine a soapy solution removing fix from the emulsion, though it's an interesting thought. However, I also don't think that I'd like a soapy solution as a substitute for Photoflo, because I think that a soapy solution might leave a residue, which is probably the last think that one would want in the last step of their film developing.
Incidentally, someone on the server said that they thought PermaWash was a soap solution. I think they're confusing it with PhotoFlo, which, as you know, is just a wetting agent to prevent water drops from marking the film when it dries. PermaWash is more like Hypo Clear in that it chemically removes fixer or changes it into more soluble, harmless compounds (I forget which!). Another very good product is Orbit Bath, which is very cheap and effective, but unfortunately seems to be now sold under a different name. Ask in a good photo shop, if you're interested. There is also Hypo Eliminator, another Kodak product, which the Kodak folks once told me is different than Hypo Clear and, I believe, is meant strictly for films, rather than both film and paper.
The history of these compounds seems to be kind of interesting, because I've read somewhere that ordinary sea water works as a hypo clearing agent and that it forms the basis for the modern products. Take it for what it's worth...
Neat method of making slides, by the way. I'm going to remember it for possible future use.
Yes, it works fine. But the biggest obstacle for someone doing this for the first time is to make sure that they have a lens that is capable of making such a small focused image. You also have to make sure that you use glass slide holders, because negative film cannot stand up to the heat of a slide projector without glass protection, because it's not as tough as slide film.
Garry
} ---------- } From: rtind-at-siu.edu[SMTP:rtind-at-siu.edu] } Sent: 15 July, 1997 09:08 } To: Garry Burgess } Subject: RE: Rush Lab + Projection Slides } } Garry, } } I'm not sure what is causing the brown staining, especially since you } indicate that it's temporary. My GUESS is that it's inadequate fixing time } or exhausted fixer. What are your fixing times? } } Randy, } } Usually I fix for 2-3 minutes, but in this case, it might be more, since the } slides were actually just piled up in a staining beaker filled with hypo. } Since the slides were in contact with each other, I'm quite sure that this } explains the inadequate fixation. Actually usually I look at them under the } safelight to ensure that they are adequately fixed, but sometimes, when I had } a huge volume, then I must have missed a couple, because I had so many. } (about 200 or so) } } I use some sort of Kodak fixer, but I can't remember the name right at this } second. I usually do not use hypochek, because as I've said, usually it's } obvious when the fixer is losing it's strength, because it starts to take } significantly longer than 2 minutes to fix our EM negatives. Because we work } under safelight, this is something that can be checked each and every time } that we develop film. } } } What fix do you use? Do } you check your fix for exhaustion with HypoChek or a similar product that } gives you a white precipitate when the fixer is saturated with silver } compounds? Interesting. } } I agree with you about the confusion about Permawash. It would have to be } something more chemically like Hypo Clear than soap, because I can't imagine } a soapy solution removing fix from the emulsion, though it's an interesting } thought. However, I also don't think that I'd like a soapy solution as a } substitute for Photoflo, because I think that a soapy solution might leave a } residue, which is probably the last think that one would want in the last } step of their film developing. } } Incidentally, someone on the server said that they thought PermaWash was a } soap solution. I think they're confusing it with PhotoFlo, which, as you } know, is just a wetting agent to prevent water drops from marking the film } when it dries. PermaWash is more like Hypo Clear in that it chemically } removes fixer or changes it into more soluble, harmless compounds (I forget } which!). Another very good product is Orbit Bath, which is very cheap and } effective, but unfortunately seems to be now sold under a different name. } Ask in a good photo shop, if you're interested. There is also Hypo } Eliminator, another Kodak product, which the Kodak folks once told me is } different than Hypo Clear and, I believe, is meant strictly for films, } rather than both film and paper. } } The history of these compounds seems to be kind of interesting, because } I've read somewhere that ordinary sea water works as a hypo clearing agent } and that it forms the basis for the modern products. Take it for what it's } worth... } } Neat method of making slides, by the way. I'm going to remember it for } possible future use. } } Yes, it works fine. But the biggest obstacle for someone doing this for the } first time is to make sure that they have a lens that is capable of making } such a small focused image. You also have to make sure that you use glass } slide holders, because negative film cannot stand up to the heat of a slide } projector without glass protection, because it's not as tough as slide film. } } Garry } }
} Sorry to do this to you, but there was a discussion a while back about } inexpensive light microscopes for children. In particular there was a } recommendation from someone in the Seattle area about a $99 model (Tasco } brand?). Of course, I deleted the messages but was recently asked by a friend } for a recommendation.
Younger children do MUCH better with an erect image, so the first decision is "dissecting" vs. compound. And another choice is monocular vs. binocular. Young children have problems with both the eye spacing and the convergance required by binocs, so the extra cost of two oculars may not be justified. } } Oh, also any recommendations for things to look at for kids ages 6-10 would be } appreciated.
Children want to DO things, rather than just look. My grandson was fascinated with watching epsom salt crystals grow when he was 6. You'll find a wealth of suggestions in the Project MICRO bibliography (see below); look in section IIA, "The microscopic world".
Caroline Schooley Educational Outreach Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/
} Date: Mon, 14 Jul 1997 15:54:34 -0500 } From: David Knecht {knecht-at-uconnvm.uconn.edu} } To: microscopy-at-sparc5.microscopy.com } Subject: ConA-labeled } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Does anyone know any good sources besides Sigma for conA labeled with } Rhodamine, Texas Red or another non-FITC dye. Thanks- Dave } } Dr. David Knecht } Department of Molecular and Cell Biology } University of Connecticut } U-125 } Storrs, CT 06269 } Knecht-at-uconnvm.uconn.edu } Linscott's Directory of Immunological and Biological Reagents (ISBN: 0-9604920-4-6, Phone: 415-544 9555, about $70) is a fantastic reference that lists almost every antibody made by every company. It is a valuable book I recommend for anyone doing immunostaining.}
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
Hello Gilbert and cyber-folk Evans blue can also be used to determine cellular integrity in plant cell suspensions, squashes, etc. The dye will not penetrate intact plasmalemma, while staining damaged cells.
Yet another use for your case of stain! Have you considered a "garage sale"?
James Wesley-Smith EM Unit University of Natal Durban, South Africa
------------------------------
On Sun, 13 Jul 1997, Gilbert T. Groehn wrote:
} Our lab acquired a case of EVANS BLUE stain } from one of our clients and we can not locate } this in any of our chemistry manuals. } } What are the normal uses for this stain (eg. } histology, hematology, etc.) ? We would be } using it only for light microscopy. } } What is the standard mixing ratios and } formulas for various applications? } } Any help on this item would be most appreciated. } } TIA } Gil Groehn } ULTRAMED, INC. } } ============================================================= } ULTRAMED, INC. Research Div. ad408-at-detroit.freenet.org } Grosse Pointe Farms, MICH 48236 USA Biomedical Consultants } =============================================================== }
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Mike,
It would be interesting to know a situation where this enlarger would be the preferred method of producing prints as opposed to taking the negatives that need dodge/burn, scan them and then digitally "correct" them? Good luck on finding this enlarger used etc,!
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Hello Everyone:
Does anyone know where I can get a used LogEtronics 55-EM enlarger (or something similar with auto-dodge & burn-in capabilities)in good working condition?
Thanks,
Michael Coviello Lab Manager The University of Texas -at- Arlington --IMA.Boundary.790299868 Content-Type: text/plain; charset=US-ASCII; name="RFC822 message headers" Content-Transfer-Encoding: 7bit Content-Description: cc:Mail note part Content-Disposition: inline; filename="RFC822 message headers"
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} from what i understand, Permawash is just soapy water. i simply use a } mild detergent solution instead and it works fine } You may be thinking of Photoflo - which is essentially a wetting agent, a detergent - used to break the hydrophobicity of film and permit sheeting of the water. This is similar to the wetting agents used in dishwashing machines (e.g., for spotless glassess). Permawash should contain some chemical scavengers used to remove resisual fixers in the film/papers. Any photo-chemical types listening to this?
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing Southern Illinois University Carbondale, IL 62901-4402 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
I am interested in the size of particle that you talk about, what densit= y do you see in your material having carried out the analysis?
For materials similar to carbon the volume of material involved with an electron beam of an energy suitable for most EDX analysis (say 15kV) wil= l be in excess of 2.7 microns? To carry out an analysis of 0.5 micron structures with material the density of carbon would require a beam energ= y of less than 6kV or for iron less than 10kV.
The mark you call a "burn" is this not really contamination? There is usually no damage to a specimen if you see a dark mark, contamination. T= he problem is that with your materials you may well drive off the volatile components and then you will burn out an area of specimen, the depth dependant upon the time of your observation or analysis.
However you seem to have conclusive proof that you are analysing the area=
that you have made your target. My point was not that a spot analysis is=
taboo but that we should not blindly use a spot analysis without knowing more about its accuracy. Some replies have taken the attitude that becau= se they have used the technique for many many years there cannot be a proble= m! Unless you know exactly what is happening in a microscope I do not agree=
that this attitude is sufficient, I can show you instruments where this attitude would be a disaster.
In the consultancy business the biggest problem that one confronts is the=
"scientist" who has been doing things his way for 20 years and will not change. I understand that we may well compare driving a microscope to driving a car. Who would by a new car and then go on a course to learn t= o drive it? The same approach in SEM however would be constructive; new instruments open up new techniques and new areas of investigation. From what I see the topic of scanning electron microscopy changes considerably=
within a two year cycle and operators who persist in using their 20 year old techniques, even on an old instrument, are somewhat lost! The "I onl= y use 25kV" and " an SEM will not work above 5,000X" bunch really worry me and they should also worry their supervisors! =
Is a true scientist someone who experiments, most SEM operators do not! =
This is not a one country thing it is the same world wide, just because w= e are microscopists we seem to forget we are also scientists. What do the masses think?
} Now, with negative, flatbed scanners one could scan the negative (10 } minutes) and print an image on transparency paper with an inkjet } (Epson 1440 dpi or a dyesub printer). Haven't tried this with TEM } negatives but I know people who do. } I have no doubt that this would work, but it requires yet more capital equipment. And here in Manitoba Canada, the money is NOT AVAILABLE. Hence, I have to dismiss all the talk about digital images and scanners, and that sort of thing. It's something like arguing that a Mercedes is a nice car. I have no doubts that it is, but I nevertheless won't be driving one in the near future, unless I win a rather large lottery. } Seems like about one hour would do it. } } What about a positive TEM film that you could project directly. } I have not heard about the existence of this sort of TEM film, but even if we did have the film, we would still suffer from the fact that we would have to crop a considerable amount out of the frame in order to get it to fit our slide projector, even with the supersize size. (ie: 2X2")
Secondly, we usually don't know ahead of time that we will need the transparencies. The urgent images are the 8X10" prints, of which I'm rushed to produce as fast as possible. There is no time for a Pathologist to scope the case twice, once with positive film. And in any event, we would need a record on negative film in any event.
I'm sure that I could develop this normal TEM film in a reversal sort of way, in the same manner that one might work with T-MAX film developed with reversal chemistry, but this produces positives directly, and there is no negative produced, so that I would have the above problems, plus the cropped images.
It seems to be quite satisfactory to produce my positive slides the way I do, and usually speed is not a big factor with these positive images.
Garry } } } } } ______________________________ Reply Separator } _________________________________ } Subject: RE: Rush Lab + Projection Slides } Author: GBurgess (GBurgess-at-exchange.hsc.mb.ca) at unix,mime } Date: 7/14/97 2:02 PM } } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} the biggest problem that one confronts is the "scientist" who has been = doing } things his way for 20 years and will not change.
That's certainly not the case for me! I'm a chemist who was been partially using myself and partially supervising a new SEM for less than a year. I've subscribed to the list in the hopes that I'd learn something.
} I am interested in the size of particle that you talk about, what = density } do you see in your material having carried out the analysis?
} For materials similar to carbon the volume of material involved with = an } electron beam of an energy suitable for most EDX analysis (say 15kV) = will } be in excess of 2.7 microns? To carry out an analysis of 0.5 micron } structures with material the density of carbon would require a beam = energy } of less than 6kV or for iron less than 10kV.
I guess I didn't express myself very clearly. We're looking at low-level amounts of inorganic particulate contaminants or additives in plastic (C,H,O) and what we want is QUALITATIVE information, such as "Does this particle contain calcium or does it contain silicon?" We wouldn't dare to try say anything about amounts (other than perhaps "lots" or "hardly any") because we're never sure how much of the polymer matrix we're hitting along with the particle. (The particles are at varying depths.)
In my--admittedly still amaturish--opinion, spot analysis can be OK if what you want is qualitative information. You have to make sure that the customers don't think the results are in any way quantitative. (They always want to and you have to keep on banging on 'em.)
} The mark you call a "burn" is this not really contamination? There is } usually no damage to a specimen if you see a dark mark, contamination.
I take the placement of the "burn" mark as proof that the positioning of the beam in spot mode is OK. As to what the dark "burn" mark actually is, I'm open. (I put those quotes on the word for a reason!) However I think it's thermal degradation of the polymer, which melts around 260=B0 = C and softens and shrinks before that. I think this because we don't get these marks when we look at metals, etc.
Cindy Bennett Hoechst Diafoil
Disclaimer: these are my opinions only and not those of my company.
we are drifting well away from the main theme, but you can still make slides with your enlarger even if you haven't got close focussing on your lens. It's more fiddly but if you have a light box you simply put the e.m. negative on the light box under the enlarger and the film to be exposed in the negative carrier. The tricky bit is getting the position and focus right but you do most of that by enlarging something of the right format first onto the light box. Then of course when you're exposing the film in the enlarger you must remember to turn on the light box and not the enlarger.
I have used this a couple of times and it works fine in an emergency although of course if you were doing a lot it would be easier to make the slides with a close-up 35mm camera and a light box..
Malcolm Haswell e.m. unit University of Sunderland UK ----------
{SNIP} Neat method of making slides, by the way. I'm going to remember it for possible future use.
Yes, it works fine. But the biggest obstacle for someone doing this for the first time is to make sure that they have a lens that is capable of making such a small focused image. You also have to make sure that you use glass slide holders, because negative film cannot stand up to the heat of a slide projector without glass protection, because it's not as tough as slide film.
I will tried the glow discharge and the carbon film to change the hydrophobicity...and also putting my grids (specimen and stain) in the oven 60C for 24 hours before looking at them to get rid of possible water trapping. I really appreciate your help, especially during summer time when a lot of people are absent...
thanks again,
Diane Montpetit microscopist Food research center agro alimentaire canada st-hyacinthe, quebec, canada tel 514 773 1105 fax 514 773 8461 e mail montpetitd-at-em.agr.ca
First, someone working 20 years in the industry is, by definition, successful. This means that he/she is providing information that is useful to others in solving their problems.
Second, our employers could care less about kv's, resolution, magnification, etc. Their interest is in solving problems. Doing the same thing for 20 years (even with a 20 year old microscope) may be what it takes. I have seen old-timers with a 20 year old scope work circles around newbies with newbie microscopes.
Finally, I agree whole-heartedly with Steve--we must be open to new ideas to solve difficult problems. I worked for a major computer manufacturer to solve a circuit board plating problem (wavy circuit traces). They ALWAYS USED 25 kv to examine the photo resist on the boards that produced the circuit traces. At 25 kv they looked straight and clean. I showed them what they looked like at 5kv and at 2kv. The edges were full of unremoved resist film whose pattern matched the wavy lines identically.
Providing INFORMATION to solve problems is our goal.
Jim Harper Amoco Fabrics and Fibers ______________________________ Reply Separator _________________________________
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I am interested in the size of particle that you talk about, what density do you see in your material having carried out the analysis?
For materials similar to carbon the volume of material involved with an electron beam of an energy suitable for most EDX analysis (say 15kV) will be in excess of 2.7 microns? To carry out an analysis of 0.5 micron structures with material the density of carbon would require a beam energy of less than 6kV or for iron less than 10kV.
The mark you call a "burn" is this not really contamination? There is usually no damage to a specimen if you see a dark mark, contamination. The problem is that with your materials you may well drive off the volatile components and then you will burn out an area of specimen, the depth dependant upon the time of your observation or analysis.
However you seem to have conclusive proof that you are analysing the area that you have made your target. My point was not that a spot analysis is taboo but that we should not blindly use a spot analysis without knowing more about its accuracy. Some replies have taken the attitude that because they have used the technique for many many years there cannot be a problem! Unless you know exactly what is happening in a microscope I do not agree that this attitude is sufficient, I can show you instruments where this attitude would be a disaster.
In the consultancy business the biggest problem that one confronts is the "scientist" who has been doing things his way for 20 years and will not change. I understand that we may well compare driving a microscope to driving a car. Who would by a new car and then go on a course to learn to drive it? The same approach in SEM however would be constructive; new instruments open up new techniques and new areas of investigation. From what I see the topic of scanning electron microscopy changes considerably within a two year cycle and operators who persist in using their 20 year old techniques, even on an old instrument, are somewhat lost! The "I only use 25kV" and " an SEM will not work above 5,000X" bunch really worry me and they should also worry their supervisors!
Is a true scientist someone who experiments, most SEM operators do not! This is not a one country thing it is the same world wide, just because we are microscopists we seem to forget we are also scientists. What do the masses think?
Hello all. last week I stupidly stuck a Cu grid on top of the area of interest of a TEM cross section. Having tried to remove it for a few days, I thought I'd turn to the microscopy community for some help! The sample is Si, polished to less than ten microns thick (orangey colour), with Al and SiO2 on the top surface. It's the only one I have. I stuck the 2x1mm slot grid onto the sample with 5-minute epoxy (devcon); I didn't realise the grid was in the wrong place until a couple of hours later. Since then I've soaked it in dmf (dimethylformamide) [3 days] acetone [a few hours] and ashed it in nitrogen and oxygen [a couple of hours]. The sample is still in one piece and stuck to the grid. I've tried pushing it about with a fine hair but it's still well fixed. So, while it is soaking for a little longer in dmf and being ashed periodically, has anyone got any bright ideas how to rescue my sample?
Many thanks in advance,
Richard Beanland, Gmmt Ltd., Caswell, Towcester, Northants NN12 8EQ
Tel +44 1327 356363 Fax +44 1327 356775 e-mail richard.beanland-at-gecm.com
For all who are interested in burns in polymers under SEM, one does not only get them in Spot Mode, but when looking at high magnification in raster mode. Strange things can happen when looking at banded spherulites of polyethylene, for example: because of the periodic variations in crystal orientation, one gets mass transport (not mass transit, which is the underground railway in Hong Kong!) and the banded spherulitic structure appears to "develop" as if by etching, but what appears is in fact an artifact which mimics the real thing.
Similar developments occur when trying to look at these beasties directly in sections under the TEM, which is one reason why staining (chlorosulphonic acid, RuO4) and etching (permanganic) techniques were developed.
There is a rather poor quality picture of a banded spherulite on my home page: URL as in the signature, but to go straight there type:
I have looked through all of my EM catalogues, checking the section on Photography for anything like Permawash, but the only product that I can find that comes close to that idea of hypo remover is the Kodak product Hypo Eliminator. My problem with the Hypo Eliminator is that the instructions suggested that the working solution only lasts 24 hours, but I wanted to keep it in a stainless steel tank for about a week, and just use it conveniently, without having to make up a working solution every day.
That said, providing there were no water marks on the film, I'm still wondering if I can rewash it later for 20 minutes, after the negatives have already dried, and effectively remove any fixer that might have been left in the emulsion the first time for a very abbreviated wash.
Other people have suggested that Kodak was being very conservative with their wash times, but I'm not sure if this is true. Kodak had no difficulty shortening the wash times for RC paper when they felt that a 2 minute wash time was sufficient for this photographic paper. So, if they really felt that 2 or 3 minutes wash time was sufficient for their EM film, then I would suspect that that would be their recommendation.
Garry } } } from what i understand, Permawash is just soapy water. i simply use a } } mild detergent solution instead and it works fine } } } You may be thinking of Photoflo - which is essentially a wetting agent, a } detergent - used to break the hydrophobicity of film and permit sheeting } of the water. This is similar to the wetting agents used in dishwashing } machines (e.g., for spotless glassess). Permawash should contain some } chemical scavengers used to remove resisual fixers in the film/papers. Any } photo-chemical types listening to this? } } } #################################################################### } John J. Bozzola, Ph.D., Director } Center for Electron Microscopy } Neckers Building, Room 146 - B Wing } Southern Illinois University } Carbondale, IL 62901-4402 } U.S.A. } Phone: 618-453-3730 } Fax: 618-453-2665 } Email: bozzola-at-siu.edu } Web: http://www.siu.edu/departments/shops/cem.html } #################################################################### } } }
The thread of this conversation appears to be drifting toward making projection slides from EM negatives.
The technique that I have used for years is to place the EM negative (SEM or TEM) on a light table with a cardboard mask around the negative to block peripheral light. Use Kodak Technical Pan film in a standard 35 mm camera with a macro lens. Bracket exposures 1/2 f-stop and develop for maximum contrast. (From start to mounted slides, less than 1 hour).
One gets crisp slides with perfect contrast (assuming that the original negatives were good). The advantage is that one can crop the EM negative by adjusting distance of the camera from the negative. Disadvantage is that one cannot label the slide unless one is willing to put rub-on letters on the EM negatives (I don't).
______________________________________________________________________ Donald L. Lovett e-mail: lovett-at-tcnj.edu Assoc. Professor, Dept. of Biology voice: (609) 771-2876 The College of New Jersey fax: (609) 771-2674 Trenton, NJ 08650-4700
Taking epoxy off without damaging the silicon or aluminum is tough. Acetic acid will work but may damage the silicon film. It does not attack bulk aluminum (much) but at the thin film level it might. Methyl cloride is also useful for dissolving epoxy--some paint removers will also dissolve epoxy.
best regards mark
Mark W. Lund, PhD Director } } Soft X-ray Web page http://www.moxtek.com { { MOXTEK, Inc. 452 West 1260 North Orem UT 84057 801-225-0930 FAX 801-221-1121 lundm-at-xray.byu.edu
"The state is good at simple tasks, like killing people and seizing their wealth. It has far more trouble reaching inside individuals and making them good." Doug Bandow
Malcolm, This sounds bizarre. Are you talking about a contact print here on to film? Or do you mean leave the negative carrier in the carrier holder of the enlarger, and use it upside down? (sort of like using the enlarger as an upside down camera.)
Garry
} Garry } } we are drifting well away from the main theme, but you can still make slides } with your enlarger even if you haven't got close focussing on your lens. } It's more fiddly but if you have a light box you simply put the e.m. } negative on the light box under the enlarger and the film to be exposed in } the negative carrier. } The tricky bit is getting the position and focus right but you do most of } that by enlarging something of the right format first onto the light box. } Then of course when you're exposing the film in the enlarger you must } remember to turn on the light box and not the enlarger. } } I have used this a couple of times and it works fine in an emergency } although of course if you were doing a lot it would be easier to make the } slides with a close-up 35mm camera and a light box.. } } Malcolm Haswell } e.m. unit } University of Sunderland } UK } ---------- } From: Garry Burgess } To: 'Microscopy Society of America } Subject: RE: Rush Lab + Projection Slides } Date: 15 July 1997 18:12 } } {SNIP} } Neat method of making slides, by the way. I'm going to remember it for } possible future use. } } Yes, it works fine. But the biggest obstacle for someone doing this for the } first time is to make sure that they have a lens that is capable of making } such a small focused image. You also have to make sure that you use glass } slide holders, because negative film cannot stand up to the heat of a slide } projector without glass protection, because it's not as tough as slide film. } } Garry } }
Hi Richard, Acetone disolves araldite so it seems a good start. Try soaking some of your cured epoxy (not your specimen) in acetone for a while and see if it softens, if it does then soak your specimen in it for longer. It is probably very difficult for the acetone to get right under the copper grid so it may need quite a long time. Plasma ashing seems to spread epoxy around the specimen but does not appear to remove it.
Good luck.
Ron =========================================================================== Mr. Ron Doole e-mail ron.doole-at-materials.ox.ac.uk Department of Materials, phone +44 (0) 1865 273701 University of Oxford, fax +44 (0) 1865 283333 Parks Road. Oxford. OX1 3PH. UK. ============================================================================
To get slides with labelling, I overlay a piece of acetate with the letters, arrows etc. rubbed onto it, placed carefully on top of the negative. Provided the acetate is clean, it works very well. Of course, my negatives are 3in x 4in; it might be more difficult with 35mm film.
Lesley Weston.
On Wed, 16 Jul 1997, Donald Lovett wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } The thread of this conversation appears to be drifting toward making } projection slides from EM negatives. } } The technique that I have used for years is to place the EM negative (SEM } or TEM) on a light table with a cardboard mask around the negative to } block peripheral light. Use Kodak Technical Pan film in a standard } 35 mm camera with a macro lens. Bracket exposures 1/2 f-stop and develop } for maximum contrast. (From start to mounted slides, less than 1 hour). } } One gets crisp slides with perfect contrast (assuming that the original } negatives were good). The advantage is that one can crop the EM negative } by adjusting distance of the camera from the negative. Disadvantage is } that one cannot label the slide unless one is willing to put rub-on } letters on the EM negatives (I don't). } } ______________________________________________________________________ } Donald L. Lovett e-mail: lovett-at-tcnj.edu } Assoc. Professor, Dept. of Biology voice: (609) 771-2876 } The College of New Jersey fax: (609) 771-2674 } Trenton, NJ 08650-4700 } } } }
1. How long can I store rat tissue in glutaraldehyde fixative before conpleting the tissue preparation process and not experience tissue degradation? Can I go as long as three weeks? At this point, I don't know if I will be processing for SEM (CPD) or TEM - depends on the LM results.
2. Should I store in buffer rather than the fixative? Some other solution?
3. Would I use a different formulation of fixative for this kind of delayed processing of tissue?
So much for quick questions and thanks so much for your help! Now, I have to go answer someone else's question.
1. How long can I store rat tissue in glutaraldehyde fixative before conpleting the tissue preparation process and not experience tissue degradation? Can I go as long as three weeks? At this point, I don't know if I will be processing for SEM (CPD) or TEM - depends on the LM results.
2. Should I store in buffer rather than the fixative? Some other solution?
3. Would I use a different formulation of fixative for this kind of delayed processing of tissue?
So much for quick questions and thanks so much for your help! Now, I have to go answer someone else's question.
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Dear Richard:
I was intrigued by your question and decided to contact Devcon directly. I was told by their applications engineer that the 5 minute epoxy will withstand DMF, acetone and most anything else. He said the only thing it doesn't stand up too well against is water!
He suggested soaking the sample in warm water for a few hours and then prying it apart with a razor blade. When I explained how fragile the sample was, he just said to soak it longer and it will come apart. If you still have trouble getting it apart, try contacting Devcon directly. They have a help section on their web site - unfortunately, I forgot to copy down their web site address, but I found it just by searching on "Devcon".
David Henriks TEL: 800-728-2233 (toll-free in USA) South Bay Technology, Inc. 714-492-2600 1120 Via Callejon FAX: 714-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of Precision Sample Preparation Equipment and Supplies for Metallography, Crystallography and Electron Microscopy.
Message text written by Richard Beanland +44 1327 356363 } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hello all. last week I stupidly stuck a Cu grid on top of the area of interest of a TEM cross section. Having tried to remove it for a few days, I thought I'd turn to the microscopy community for some help! The sample is Si, polished to less than ten microns thick (orangey colour), with Al and SiO2 on the top surface. It's the only one I have. I stuck the 2x1mm slot grid onto the sample with 5-minute epoxy (devcon); I didn't realise the grid was in the wrong place until a couple of hours later. Since then I've soaked it in dmf (dimethylformamide) [3 days] acetone [a few hours] and ashed it in nitrogen and oxygen [a couple of hours]. The sample is still in one piece and stuck to the grid. I've tried pushing it about with a fine hair but it's still well fixed. So, while it is soaking for a little longer in dmf and being ashed periodically, has anyone got any bright ideas how to rescue my sample?
Many thanks in advance,
Richard Beanland, Gmmt Ltd., Caswell, Towcester, Northants NN12 8EQ
I am planning to purchase a video or digital camera to document, analyze, and print images of paint and fiber samples using polarized light microscopy and fluorescence microscopy. Previous threads re video or digital cameras have focused largely on black and white images obtained using SEM or TEM and printed using high-DPI inkjets or dye-sub printers.
May I ask for recommendations or comments on purchasing a system (input through output) to deal with colored images of paint cross-sections, fibers, petrographic samples, etc.? This system would allow still images to be captured, analyzed/manipulated using image analysis software (including Photoshop), embedded in reports, and printed with near photographic quality in color.
I'll be pleased to provide more details, as requested.
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Continuing the discussion of this topic. I used twin jet electropolishing a bit about 1970, but was frustrated by not being able to determine when a shiny surface was produced. (I had been able to see the sample in an old glass system operated manually). About that time, one of the early South Bay 550 polishers was ordered by me here at Argonne because it permitted magnified, IN SITU, viewing of the sample during polishing. This was needed to thin 1 or 2 new materials every week! The ease of use permitted accumulating reproducible data published in a 65 page report, (ANL-80-120), used world wide, a journal cover photo, and 12 other published articles. This work brought me the MSA "Technologist of the Year" award in 1994. About six 550 polishers are used exclusively at Argonne-some as long as 25 years! The unit can do jetting from one side, (for back-thinning to a special surface-lacquer protected), or both sides by inverting the sample after jet polishing about half way thru it. Microshield lacquer, (from South Bay), works great to protect surfaces from etching, dissolves in acetone, and may be thinned to reduce shrinkage when thinning soft, annealled copper for example. Also, the entire 3 m.m. disc surface can be polished via a 3 m.m. jet; using an external "timer/switch",and a D.C. power supply, planar "sectioning" of as little as 100 nm. can be removed from a surface. The jet polishing electrolyte and conditions will work. The large jet can be used to etch a surface for optical photos by simply reducing the "polishing" voltage about 20% for a couple of seconds! The 300 volt, 150 mA. capacity power supply exeeds other manufacturer's units and makes use of non-acid "BK-2" type electrolytes possible-a must for many materials. The line-of-sight optical shut off system can be fitted with a variety of color spectrum light sources for special uses. The standard infra red LED and detector bias may be independently adjusted to give the desired sensitivity setting. It will make electron transparant regions in pure annealled metal such as aluminum-with no hole! Of course the setting is normally set for a 20 micron hole with a very thin edge (quite reproducible, of course). Alignment of the parts is easy and stays set a long time. Even saphire light pipes are available for hydrofluoric acid or bromine/alcohol solutions. PVC plastic parts are available and may be substituted for metal ones for such strong chemical baths. Low temperatures of -50 degrees C. are no problem. The sample is accesible for rapid rinsig after swinging the detent-equipped jet support to one side. In 25 years of use, these instruments have saved one man per year in labor cost, (roughly $100,000/yr.), or $2,500,000--due to the ease and speed with which excellent samples can be made. About 90 to 95% of the samples attempted are good once- conditions are established. In my opinion, all the jet polishers have improved with time, but the South Bay 550 C and 550 D units are unmatched when it comes to working with the newer, difficult materials which should be viewd DURING thinning. They permit me to thin about 300 TEM foils/year in my spare time.
Might as well throw in my two cents worth on this topic. We generally make our projections slides from prints, rather than negatives. This, of course, requires making prints first, but since this is often done anyway, it's usually not a problem.
We use a seldom-mentioned film known as Kodak 5468, a direct positive film used, I think, as a motion picture stock. It's bright red and translucent---strange looking stuff. Our exposures on a 4-light copy stand range from 6-10 seconds at a lens opening of f/3.5 on a Canon 55mm macro lens, so it obviously requires a lot of light. Development is in Dektol diluted 1:1 with water. Yes, Dektol, the paper developer. Stop with water, fix with whatever you usually use and wash normally.
This film is cheap (still less than $25/100 ft., last I checked), development is easy, and the slides are quite good. The problem is that this film tends to undergo reduction-oxidation (redox) after a few years, giving a solarized appearance. To prevent this, use Kodak Brown Toner (TOXIC---use plenty of ventilation and gloves) diluted about 1:50 for a 30-second dip. This will often give your slides a slight brown tone. Some folks like this, some don't.
Not a perfect solution, but very useful for many purposes.
Randy Tindall Center for Electron Microscopy Southern Illinois University at Carbondale
Gerry et al., Perma Wash is advertised as an archival high speed wash for both film and paper that reduces wash time by 90%. It's made by Heico (a Cambrex Co.) along with other kinds of photographic chemicals. I've used it for years both as an EM specialist and as a photographer with no problems at all. You should check with local photographic darkroom supply house instead of any EM suppliers. Two suppliers that I know who do offer it are listed if you still can't find it in your area. For film Heico states archival permanence with one minute first water wash, one minute Perma Wash, one minute final water wash; for r.c. paper, it's two minutes, two minutes, two minutes; finally, for double weight papers, five minutes, five minutes, five minutes does the job. I hope this washes well for you. :-) :-) :-}
Camera World Crimson Tech PO Box 9426 325 Vassar Street 1809 Commonwealth Ave Cambridge, MA 02139 Charlotte, NC 28205 Pho: 800-868-3686 800-868-5150 704-375-8453 617-868-5150 Fax: 704-376-1826 617-499=4777
HEICO Chemicals Inc. Route 611 Delaware Water Gap, PA 18327
717-420-3900
Contains ammonium sulfite, sodium sulfite and water.
The container say it specifically removes silver sulfate.
Tom
Thomas Moninger moninger-at-emiris.iaf.uiowa.edu University of Iowa Central Microscopy Research Facility http://www.uiowa.edu/~cemrf Views expressed are mine.
} last week I stupidly stuck a Cu grid on top of the area of interest } of a TEM cross section. Having tried to remove it for a few days, I thought } I'd turn to the microscopy community for some help! } The sample is Si, polished to less than ten microns thick (orangey colour), } with Al and SiO2 on the top surface. It's the only one I have. I stuck the } 2x1mm slot grid onto the sample with 5-minute epoxy (devcon); I didn't real- } ise the grid was in the wrong place until a couple of hours later. Since } then I've soaked it in dmf (dimethylformamide) [3 days] acetone [a few } hours] and ashed it in nitrogen and oxygen [a couple of hours]. The sam- } ple is still in one piece and stuck to the grid. I've tried pushing it } about with a fine hair but it's still well fixed. } So, while it is soaking for a little longer in dmf and being ashed } periodically, has anyone got any bright ideas how to rescue my sample? } } Many thanks in advance, } } Richard Beanland,
} I was intrigued by your question and decided to contact Devcon directly. } I was told by their applications engineer that the 5 minute epoxy will with- } stand DMF, acetone and most anything else. He said the only thing it does- } n't stand up too well against is water! } He suggested soaking the sample in warm water for a few hours and then pry- } ing it apart with a razor blade. When I explained how fragile the sample } was, he just said to soak it longer and it will come apart.
} David } Dear Richard, Here's a thought: If just soaking won't do, you might try micro- waving the water-soaked specimen. That will heat up the water, but maybe not the Si, Al or Cu. I know metal is a no-no in a microwave oven, but there is so little that there shouldn't be a disaster here. I'd put ~50 ml of H2O in a beaker in the oven at the same time. Of course, I'd try it first with something other than the specimen. Good luck. Yours, Bill Tivol
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Continuing the discussion of this topic. I used twin jet electropolishing a bit about 1970, but was frustrated by not being able to determine when a shiny surface was produced. (I had been able to see the sample in an old glass system operated manually). About that time, one of the early South Bay 550 polishers was ordered by me here at Argonne because it permitted magnified, IN SITU, viewing of the sample during polishing. This was needed to thin 1 or 2 new materials every week! The ease of use permitted accumulating reproducible data published in a 65 page report, (ANL-80-120), used world wide, a journal cover photo, and 12 other published articles. This work brought me the MSA "Technologist of the Year" award in 1994. About six 550 polishers are used exclusively at Argonne-some as long as 25 years! The unit can do jetting from one side, (for back-thinning to a special surface-lacquer protected), or both sides by inverting the sample after jet polishing about half way thru it. Microshield lacquer, (from South Bay), works great to protect surfaces from etching, dissolves in acetone, and may be thinned to reduce shrinkage when thinning soft, annealled copper for example. Also, the entire 3 m.m. disc surface can be polished via a 3 m.m. jet; using an external "timer/switch",and a D.C. power supply, planar "sectioning" of as little as 100 nm. can be removed from a surface. The jet polishing electrolyte and conditions will work. The large jet can be used to etch a surface for optical photos by simply reducing the "polishing" voltage about 20% for a couple of seconds! The 300 volt, 150 mA. capacity power supply exeeds other manufacturer's units and makes use of non-acid "BK-2" type electrolytes possible-a must for many materials. The line-of-sight optical shut off system can be fitted with a variety of color spectrum light sources for special uses. The standard infra red LED and detector bias may be independently adjusted to give the desired sensitivity setting. It will make electron transparant regions in pure annealled metal such as aluminum-with no hole! Of course the setting is normally set for a 20 micron hole with a very thin edge (quite reproducible, of course). Alignment of the parts is easy and stays set a long time. Even saphire light pipes are available for hydrofluoric acid or bromine/alcohol solutions. PVC plastic parts are available and may be substituted for metal ones for such strong chemical baths. Low temperatures of -50 degrees C. are no problem. The sample is accesible for rapid rinsig after swinging the detent-equipped jet support to one side. In 25 years of use, these instruments have saved one man per year in labor cost, (roughly $100,000/yr.), or $2,500,000--due to the ease and speed with which excellent samples can be made. About 90 to 95% of the samples attempted are good once- conditions are established. In my opinion, all the jet polishers have improved with time, but the South Bay 550 C and 550 D units are unmatched when it comes to working with the newer, difficult materials which should be viewd DURING thinning. They permit me to thin about 300 TEM foils/year in my spare time.
We are having a mini-debate here over whether one can confidently say what side of the membrane an epitope on a membrane faces (e.g., towards the cytoplasm or lumen of the RER) based on the distribution of gold labeling. If most the labeling is on one side, would you be confident the epitope is solely on that side? One of my collaborators had a paper criticized by a reviewer who said this couldn't be done reliably.
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility 3 Tucker Hall University of Missouri Columbia, MO 65211 (573)-882-4712 (voice) (573)-882-0123 (fax)
OOPs--I intended to say that acetic acid may damage the aluminum film. It doesn't attack silicon from my experience.
best regards mark
Mark W. Lund, PhD Director } } Soft X-ray Web page http://www.moxtek.com { { MOXTEK, Inc. 452 West 1260 North Orem UT 84057 801-225-0930 FAX 801-221-1121 lundm-at-xray.byu.edu
"The state is good at simple tasks, like killing people and seizing their wealth. It has far more trouble reaching inside individuals and making them good." Doug Bandow
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Richard Beanland described a problem involving a TEM grid stick to a sample in the wrong place, glued with a 5-Minute (Devcon) epoxy.
One is his comments was: ================================================ I've soaked it in dmf (dimethylformamide) [3 days] acetone [a few hours] and ashed it in nitrogen and oxygen [a couple of hours]. The sample is still in one piece and stuck to the grid. I've tried pushing it about with a fine hair but it's still well fixed. ================================================ Oxygen plasma etching, in an isotropic (I stress isotropic as opposed to anisotropic) plasma etcher should remove the epoxy. If you were correct in that you used "nitrogen and oxygen", then this is why it did not work. You have to use pure oxygen, no nitrogen. Even a small leak in the system, allowing just a small partial pressure of nitrogen will literally kill the etching rate. In air, literally nothing will happen.
Now, if in fact you were using pure oxygen, there could still be a leak problem, another reason why you did not get any etching. But the technique should work. Make sure the power is not more than 100 watts or else the sample might heat up to temperature that would not be acceptable.
Disclaimer: SPI manufactures an isotropic plasma etcher for doing this kind of etching of organic materials. You can see further information on our website as well as an explanation of the differences between isotropic vs. anisotripic etching.
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 ############################ ==================================================
Does anyone have a tried and true method for the fixation and embedment of Tetrahymena? I would like to do some preembedding staining for nuclear proteins, and have been having trouble once the dehydration is started. The cells are blasted apart by the time I examine them under the electron microscope. The cells are intact after preembedding immunostaining (at least they appear to be under phase contrast). After a post immunostaining treatment with 1% glutaraldehyde, the cells are enrobed in agar (1:1 of 2% low-melting temperature agar) at 37C, dehydrated in a graded ethanol series (10, 30, 50, 75, 95, 100, 100 -- 15 minutes each) and then progressively infiltrated with LR White resin (all at room temp). Polymerization is at 55C for 24 hours.
Any help/ideas would be greatly appreciated.
Craig Lending Department of Biology SUNY Brockport Brockport, NY 14420 Phone: 716-395-5755 Fax: 716-395-2741 e-mail: clending-at-acs.brockport.edu
Damian - Sabatini (he made GA the EM fixative) did some experiments and declared that postfixation could be left for at least six months. I guess that is true for a few tissues. Lipids are not well fixed in GA and lipid rich tissues in particular suffer when post fixation is delayed. By how much - well how long is a piece of string? Postfix as soon as possible. Store in buffer refrigerated. What you do for SEM or LM matters very little, but for TEM this matters. Never store tissues in GA for TEM. GA crosslinks materials, overfixing with GA results in a coarser texture which obscures fine details at high resolution. Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au } } 1. How long can I store rat tissue in glutaraldehyde fixative before } conpleting the tissue preparation process and not experience tissue } degradation? Can I go as long as three weeks? At this point, I don't } know if I will be processing for SEM (CPD) or TEM - depends on the LM } results. } } 2. Should I store in buffer rather than the fixative? Some other } solution? } } 3. Would I use a different formulation of fixative for this kind of
} delayed processing of tissue? } } So much for quick questions and thanks so much for your help! Now, I } have to go answer someone else's question. } } Damian Neuberger } neuberd-at-baxter.com
Dear Richard, The only way I have removed 5-minute epoxy from something was by gently heating. The epoxy softens at heat-gun temps (70 deg.C?). This was on a leaky air-pressure valve, not a grid, so I just heated it until I could peel it off.. The other suggestion would be acetone, at least overnight. If you ask a embedding/biologist EM type, they may know a solvent to dissolve epoxy. You wrote:
} Hello all. } last week I stupidly stuck a Cu grid on top of the area of interest } of a TEM cross section. Having tried to remove it for a few days, I thought } I'd turn to the microscopy community for some help! } The sample is Si, polished to less than ten microns thick (orangey colour), } with Al and SiO2 on the top surface. It's the only one I have. I stuck the } 2x1mm slot grid onto the sample with 5-minute epoxy (devcon); I didn't realise } the grid was in the wrong place until a couple of hours later. Since then } I've soaked it in dmf (dimethylformamide) [3 days] acetone [a few hours] and } ashed it in nitrogen and oxygen [a couple of hours]. The sample is still in } one piece and stuck to the grid. I've tried pushing it about with a fine hair } but it's still well fixed. } So, while it is soaking for a little longer in dmf and being ashed } periodically, has anyone got any bright ideas how to rescue my sample? } } Many thanks in advance, } } Richard Beanland,
Good day everyone I would appreciate your comments on what permanent mounting media are available which meet the correct refractive index of glass and do not lead to fading of toluidene blue-stained sections?
Thank you.
James Wesley-Smith EM Unit University of Natal Durban, South Africa
Kodak Rapid Process Copy film (if it is still available) or Kodak Direct MP film (is available, e.g. from SPI or Ted Pella, etc) are excellent for inexpensive single-process preparation of B & W transparencies from EM prints.
Robin H Cross Director : EM Unit, Rhodes University, Grahamstown, South Africa eurc-at-giraffe.ru.ac.za - tel: +27 461 318168 - fax: +27 461 24377
Here is one addition (hopefully valuable) to the EDX-Spot Mode discussion. =20 If your SEM has a specimen current meter, you can use that successfully for checking the beam position very accurately before and during the EDX analysis of small particles.=20
The technique is very simple. You first position the spot mode beam at high magnification, and then by moving the beam in X-Y direction you either maximize or minimize the specimen current reading depending on whether the average atomic number (=3D Z) of the particle is lighter or=20 heavier that that of the matrix. The great advantage in the use of the=20 specimen current (=3D absorbed electrons) for positioning the beam is that= =20 you same time maximize the X-ray emission from the particle and minimize the possible X-ray contribution from the matrix. This is due to the fact that absorbed electrons "sense" the shape of the particle under the specimen surface.
If the specimen current stays constant during the measurement of the EDX spectrum, then you can be absolutely certain that the beam did not leave the particle during the measurement. However, normally there is a small increase ( { 1 %) in the specimen current due to the contamination build-up.
=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4= =A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4= =A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4 =A4 =A4 =A4 Seppo J. Sivonen e-mail: seppo.sivonen-at-oulu.fi =A4=20 =A4 University of Oulu =A4=20 =A4 Institute of Electron Optics tel: +358-8-553 3140 =A4 =A4 Box 400 fax: +358-8-553 3149 =A4=20 =A4 FIN-90571 Oulu =A4 =A4 FINLAND http://koivu.oulu.fi/~eolwww/welcome.html =A4 =A4 =A4 =A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4= =A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4= =A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4=A4
I don't know about the refractive index, but I've used Epoxy resin as a mountant quite successfully. It doesn't seem to fade Toluidine Blue semi-thins.
DePeX is not too bad but I have had some fading over long periods.
---------- } From: James Wesley-Smith {wesleysm-at-biology.und.ac.za} } To: 'Microscopy-at-sparc5.microscopy.com' {Microscopy-at-Sparc5.Microscopy.Com} } Subject: LM coverslip mounting medium } Date: 17 July 1997 09:08 } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Good day everyone } I would appreciate your comments on what permanent mounting media are } available which meet the correct refractive index of glass and do not lead } to fading of toluidene blue-stained sections? } } Thank you. } } } James Wesley-Smith } EM Unit } University of Natal } Durban, South Africa }
Perhaps I didn't make my original comments clear enough. But yes the unexposed negative goes in the enlarger film carrier in the enlarger. This has the big advantage that reduction is no problem. The disadvantage is that if your enlarger doesn't do one to one printing in the first place then you still can't do it - just the reductions. This is usually not a problem if you are doing this with e.m cut film because it is larger and so needs to be reduced. You could even do this with prints ie copying them from the baseboard of the enlarger onto the negative in the film carrier but the amounts of stray light needed for copying stand lights makes it more fiddly. I think someone else has already mentioned it is important to mask out stray light.
I stumbled upon this idea because in one lab, I worked, they had a large Durst Laborator 1000 enlarger with all the accessories. This allowed you to use it as a copy camera and gave you special large format film holders for the purpose. Another lab I worked in had a DeVere large format enlarger with full reduction facilities. It seemed reasonable therefore to use an ordinary Durst enlarger as a copy camera so that I could do the 'DeVere thing, in reverse. I just didn't have all of the clever light-tight film holders for masking unexposed film in the carrier but with care it works, anyway.
If I remember rightly many of the big Durst enlargers have a reversible mirror in the condensor system - our Durst 1200s look as if you can just take out the fitting so that it faces towards the operator rather than the lamp. You should then be able to view the image of your negative or whatever else is on the baseboard through the little window in the front of your enlarger and VOILA you have a simple camera which will do large format. I am sure it should be in the manual somewhere.
Bizarre it may seem, to use an enlarger to reduce, but isn't there a certain pleasing symmetry to it? Sorry to go on but I thought this might be of general interest.
DISCLAIMERS: I hasten to add you can only do this with some enlargers; it will disrupt normal printing in a one enlarger darkroom; you may damage the enlarger so be careful; if in doubt read the manual or ask the supplier/manufacturer; and I will refuse to re-imburse anyone for self-inflicted damage. I have no connections with Durst or De Vere other than as a satisfied user.
Malcolm Haswell ----------
Malcolm, This sounds bizarre. Are you talking about a contact print here on to film? Or do you mean leave the negative carrier in the carrier holder of the enlarger, and use it upside down? (sort of like using the enlarger as an upside down camera.)
Garry
} Garry } } we are drifting well away from the main theme, but you can still make slides } with your enlarger even if you haven't got close focussing on your lens. } It's more fiddly but if you have a light box you simply put the e.m. } negative on the light box under the enlarger and the film to be exposed in } the negative carrier. } The tricky bit is getting the position and focus right but you do most of } that by enlarging something of the right format first onto the light box. } Then of course when you're exposing the film in the enlarger you must } remember to turn on the light box and not the enlarger. } } I have used this a couple of times and it works fine in an emergency } although of course if you were doing a lot it would be easier to make the } slides with a close-up 35mm camera and a light box.. } } Malcolm Haswell } e.m. unit } University of Sunderland } UK } ---------- } From: Garry Burgess } To: 'Microscopy Society of America } Subject: RE: Rush Lab + Projection Slides } Date: 15 July 1997 18:12 } } {SNIP} } Neat method of making slides, by the way. I'm going to remember it for } possible future use. } } Yes, it works fine. But the biggest obstacle for someone doing this for the } first time is to make sure that they have a lens that is capable of making } such a small focused image. You also have to make sure that you use glass } slide holders, because negative film cannot stand up to the heat of a slide
} projector without glass protection, because it's not as tough as slide film. } } Garry
Tom I tend to agree with the reviewer. I have spent several years looking at the membrane cytroskeleton of muscle, and commonly the gold particles appear "extracellular" when we know for sure they are on the inside of the plasma membrane. The argument is that as an IgG is about 11nm long, tagged to a 5nm gold particle, and if an indirct tagging system is system is used this adds an another 11nm. this leads of a potential radius of 25nm from the label site. I am sure from quantitative studies it is less than this, though I am equally sure that the gold particle commonly does not reflect to epitope position exactly.
Simon
Tom Phillips wrote:
} ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } ----------------------------- } -----------------------------------------. } } We are having a mini-debate here over whether one can confidently say } what } side of the membrane an epitope on a membrane faces (e.g., towards the } } cytoplasm or lumen of the RER) based on the distribution of gold } labeling. } If most the labeling is on one side, would you be confident the } epitope is } solely on that side? One of my collaborators had a paper criticized } by a } reviewer who said this couldn't be done reliably. } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } 3 Tucker Hall } University of Missouri } Columbia, MO 65211 } (573)-882-4712 (voice) } (573)-882-0123 (fax)
-- Simon C. Watkins Ph.D. Associate Professor Director CBI University of Pittsburgh Pittsburgh PA 15261 tel:412-648-3051 Fax:412-648-2004 URL:http://sbic6.sbic.pitt.edu
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Reply to: RE} Au labeling of membranes - can you judge sideness?
Dear Thomas, Your discussion regarding which side of the membrane the labeling is on should include how to determine the resolution of the labeling technique. The resolution is determined by the size of the probe used for visualization as well as the antibody used. For example, an IgG molecule has an extended length of about 8-10nm and a 5nm gold particle would be expected to have a total diameter of 7-8nm of covered with protein-A. The resolution comes from the circular radius of this complex from the antigen outward and upwards. The antibody could fall anywhere within this radius, thus making it difficult to say if it is on the inside or outside of the membrane. You could confidently say it labels the membrane but to go further may be stretching what you see visually. Section thickness also plays a part in reducing the resolution. For further information you may want to pick up a book by Garreth Griffiths called "Fine structure immunocytochemistry" printed by Springer-Verlag ISBN 3-540-54805-X. It is a comprehensive look at a number of variables pertaining to immunocytochemistry.
Linda Chicoine Center for Cell Imaging Dept. of Cell Biology Yale University 203-785-3646 phone 203-785-7226 fax
--------------------------------------
We are having a mini-debate here over whether one can confidently say what side of the membrane an epitope on a membrane faces (e.g., towards the cytoplasm or lumen of the RER) based on the distribution of gold labeling. If most the labeling is on one side, would you be confident the epitope is solely on that side? One of my collaborators had a paper criticized by a reviewer who said this couldn't be done reliably.
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility 3 Tucker Hall University of Missouri Columbia, MO 65211 (573)-882-4712 (voice) (573)-882-0123 (fax)
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Could someone please tell me how to unsubscribe to the Microscopy List Server? I plan to be a way for the next couple of weeks and would like to turn the flood of messages off. Thanks Rob Willson Dept of Anatomy and Cell Biology Tufts University
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Tom Phillips wrote: ....."what side of the membrane an epitope on a membrane faces (e.g., towards thecytoplasm or lumen of the RER) based on the distribution of gold labeling."....
I would also agree that the normal gold label may be to large, because of the size of the coupled IgG moelcule. What about the Nanogold labels: 1.4 nm gold attched to FAB fragment? I don't know what the total size would be, but much smalelr than if coupled to igG.Would this be small enough? the silver enhancement, to enlarge the size for viewing on TEM, is done after label. Louisa Howard EM Facility, 6044 Gilman Dartmouth College Hanover, NH, 03755
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Well, i dont know if this is a dumb solution, but you could try to etch the copper away with somehting like dilute nitric acid. i dont know the dangers with silicon or the aluminum, but perhaps you could find an etchant that would attack only the copper.
Michael T. Marshall Research Engineer, Electron Microscopy University of Illinois at Urbana-Champaign Frederick Seitz Materials Research Laboratory 104 South Goodwin avenue Urbana, IL 61801-2985 (217) 244-8193 fax: (217) 244-2278
For what it's worth regarding the use of Permawash: I have negatives that I washed according to their (Permawash) instructions at least 25 years ago and they are still in perfectly good condition.
Thanks to all who replied to my question about storing rat tissue in fixative. Here is a cut and paste summary of replies from all over the world! for anyone who is interested.
Sabatini (he made GA the EM fixative) did some experiments and declared that postfixation could be left for at least six months. I guess that is true for a few tissues. Lipids are not well fixed in GA and lipid rich tissues in particular suffer when post fixation is delayed. By how much - well how long is a piece of string? Postfix as soon as possible. Store in buffer refrigerated. What you do for SEM or LM matters very little, but for TEM this matters. Never store tissues in GA for TEM. GA crosslinks materials, overfixing with GA results in a coarser texture which obscures fine details at high resolution.
I have stored samples in 2% glut. in 0.1M sodium cacodylate for 2 weeks because I forgot about them. These were cell cultures and they turned out OK, I embedded these into epon/araldite. I have also stored things in buffer after fixation for a few weeks and this turned out OK too.
Dysktra's book on biological EM has micrographs of mouse kidney stored in formaldehyde/glut fix for something like 2 years that looks pretty good.
I often have kept things in GA for as long as 3 weeks, provided that it is kept cold. (There may be some problem with microtubule disassembly, according to some). Over time, the GA will break down 3weeks).
Store in buffer: No. You stand the risk of fixative being washed out and the possibility of "de-fixing" the material. You also could encourage bacterial growth eventually.
My recommendation would be to fix, wash, and post-fix in OsO4. If you then dehydrate to 70% EtOH or Acetone,it will keep for YEARS!
Although we try not to store anything, we prefer to store our samples in Trumps fix. A buffered glute/form combo. We have stored for weeks at a time and had good results
There is some evidence to suggest that storage in Trumps or half Karnovsky rather than pure glut (i.e. glut plut some paraformaldehyde)
I have stored tissue in glutaraldehyde or Trump's fix(glut/para fix) for years before using it for TEM. I wouldn't store it in buffer.
I would probably do a normal fix and then switch to buffer and store cold. I don't believe you will see any degradation as long as you stay away from post-fixation with osmium before storage; I would leave that go until you were ready to proceed with prep.
Damian- I've always tried to avoid any long term storage in fix, however storing samples after fixation in a 0.2M buffer has seemed to work well for up to 1-2 weeks. And after reading the thread on lipid preservation, I think it wise to osmicate prior to the storage in buffer too. _mike
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} } Good day everyone } } I would appreciate your comments on what permanent mounting media are } } available which meet the correct refractive index of glass and do not } lead } } to fading of toluidene blue-stained sections? } } I learned a trick from a DuPont-Sorvall rep (that tells you how long ago that was!) that retards or eliminates ALL fading caused by oxidants in the mounting medium. Add 1-2% BHT (the preservative used in bologna, hot dogs, etc.) to any mounting medium. There should be a bottle of the stuff sitting on the shelf in the EML at U.C. Berkeley that has enough to supply every EM lab in the country...
Caroline Schooley Educational Outreach Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/
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} } Good day everyone } } I would appreciate your comments on what permanent mounting media are } } available which meet the correct refractive index of glass and do not } lead } } to fading of toluidene blue-stained sections? } } I learned a trick from a DuPont-Sorvall rep (that tells you how long ago that was!) that retards or eliminates ALL fading caused by oxidants in the mounting medium. Add 1-2% BHT (the preservative used in bologna, hot dogs, etc.) to any mounting medium. There should be a bottle of the stuff sitting on the shelf in the EML at U.C. Berkeley that has enough to supply every EM lab in the country...
Caroline Schooley Educational Outreach Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.MSA.microscopy.com/ProjectMICRO/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/
Richard Beanland +44 1327 356363 wrote: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Hello all. } last week I stupidly stuck a Cu grid on top of the area of interest } of a TEM cross section. Having tried to remove it for a few days, I thought } I'd turn to the microscopy community for some help! } The sample is Si, polished to less than ten microns thick (orangey colour), } with Al and SiO2 on the top surface. It's the only one I have. I stuck the } 2x1mm slot grid onto the sample with 5-minute epoxy (devcon); I didn't realise } the grid was in the wrong place until a couple of hours later. Since then } I've soaked it in dmf (dimethylformamide) [3 days] acetone [a few hours] and } ashed it in nitrogen and oxygen [a couple of hours]. The sample is still in } one piece and stuck to the grid. I've tried pushing it about with a fine hair } but it's still well fixed. } So, while it is soaking for a little longer in dmf and being ashed } periodically, has anyone got any bright ideas how to rescue my sample? } } Many thanks in advance, } } Richard Beanland, } Gmmt Ltd., } Caswell, } Towcester, } Northants NN12 8EQ } } Tel +44 1327 356363 } Fax +44 1327 356775 } e-mail richard.beanland-at-gecm.com Dear Mr. Beanland,
We have available an Epoxy Dissolver that is supposed to work and all 2 component epoxies. As I have not tried this product on all epoxies available, I do not know if it will work with this particular type.
If you can, visit your local pharmacy and ask them for some DMSO(Dimethylsulfoxide). This is the primary ingredient and it will need to be heated to operate effectively.
Please let me know if you have any other questions.
Sincerely,
Gary Liechty Allied High Tech Products, Inc. 2376 E. Pacifica Pl. Rancho Dominguez, Ca. 90220 310-635-2466 800-675-1118 310-762-6808 Fax
Products for Materialographic, SEM and TEM sample preparation
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From: Bernie Kestel-Argonne National Lab
I forgot to mention an important feature on the South Bay 550 series jet polishers in my previous message. The "open faced" specimen retainer has only a 0.0015" thick polyethylene diaphram with a central hole in it to hold the specimen down on a platinum tipped holder. This thin sheet offers almost NO flow resistance or bubble trapping area near the specimen. The all important electrolyte viscosity/polishing film can be thicker with this design, producing smoother finished surfaces while "bridging" across grain boundaries, precipitates and other features. That is why I do most polishing at -45 C. or so and add butyl cellosolve to increase electrolyte viscosity to 10-12 centipoises-ideal. (Like half & half from a refrigerator, approx.). Of course a PVC cap with a central hole positions the specimen laterally. I pass this along to hopefully ease someones prep. burden - -I'm NOT financially connected to South Bay! I feel this unit is like driving a modern auto compared to a hand cranked, manual shifted, no air conditioning machine. Take the easy route!
Our lab has been using Glutaraldehyde that expired in November of 1995, but has been kept refrigerated since then. Lately we have been noticing problems with our fixation, and I was wondering if perhaps it is because of this old Glutaraldehyde. Does anyone know how important these expiry dates really are with respect to its effect on fixation?????????
I don't mean to be hypercritical but doesn't everybody else see how to unsubscribe at the top of their message. I am a new subscriber (one week) and there must have been at least 3 messages similar to this one. Read the rules or does the old adage apply that:
OLD MICROSCOPISTS NEVER DIE THEY JUST LOSE THEIR RESOLUTION.
Mike Mead
RWILLSON-at-pearl.tufts.edu wrote:
} ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } ----------------------------- } -----------------------------------------. } } Could someone please tell me how to unsubscribe to the Microscopy } List Server? I plan to be a way for the next couple of weeks and } would like } to turn the flood of messages off. } Thanks } Rob Willson } Dept of Anatomy and Cell Biology } Tufts University
{P} I don't mean to be hypercritical but doesn't everybody else see how to unsubscribe at the top of their message. I am a new subscriber (one week) and there must have been at least 3 messages similar to this one. Read the rules or does the old adage apply that:
{P} {FONT SIZE=+2} OLD MICROSCOPISTS NEVER DIE {/FONT} {BR} {FONT SIZE=+2} THEY JUST LOSE THEIR RESOLUTION. {/FONT} {FONT SIZE=+2} {/FONT}
{P} RWILLSON-at-pearl.tufts.edu wrote: {BLOCKQUOTE TYPE=CITE} ------------------------------------------------------------------------ {BR} The Microscopy ListServer -- Sponsor: The Microscopy Society of America
{P} Could someone please tell me how to unsubscribe to the Microscopy {BR} List Server? I plan to be a way for the next couple of weeks and would like {BR} to turn the flood of messages off. {BR} Thanks {BR} Rob Willson {BR} Dept of Anatomy and Cell Biology {BR} Tufts University {/BLOCKQUOTE} {/HTML}
After reading the summary Damian posted I wanted to chip in my 2 cents worth.
One respondent recommeded storage in fix since long-term buffer storage might "unfix" the specimen. I find it hard to believe that buffer could undo the powerful crosslinking glut causes. I have heard of this possibility with formalin fixation but that is not nearly as powerful a fix as glut. Anyone know of any experiments in this area?
Another responsent advocated storage in ethanol or acetone after post-fixing in OsO4, stating the tissue would keep for years. It will keep but a lot of cytoplasm will be extracted. Hayat's book, Prin. and Tech. of EM (1981 edition) discusses this and gives references and an illustration on pp 150-155.
I would store in buffer, changing it often if I could not complete processing soon.
Geoff -- *************************************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane Piscataway, NJ 08854 voice: (732)-235-4583; fax -4029 e-mail: mcauliff-at-umdnj.edu ***************************************************************
I am in the market for a used clinical ultracentrifuge as well as an oven to be used during TEM specimen embedding. If anyone knows of a lab that is interested in selling these items, please contact me directly.
Thanks,
Dan Caruso c/o Eugene Gordon Biological Technician Medjet, Inc. 1090 King Georges Post Road Edison, NJ 08837 Phone: (732) 738-3990 Fax: (732) 738-3984 MEDJET-at-WORLDNET.ATT.NET
Garry Burgess wrote: ================================================== Our lab has been using Glutaraldehyde that expired in November of 1995, but has been kept refrigerated since then. Lately we have been noticing problems with our fixation, and I was wondering if perhaps it is because of this old Glutaraldehyde. Does anyone know how important these expiry dates really are with respect to its effect on fixation?????? ================================================== You are quite correct in that there can be some degree of arbitrariness in the statement of the expiration date. At least to us, the expiration date should correlate with some future point in time, after which, one could expect to see some deterioration of performance. Of course many of us know that film and paper, if properly stored don't turn into pumpkins on their expiration dates.
In the case of glutaraldehyde, the two most important factors influencing what will be the actual expiration date (as opposed to that stamped on the product), in the case of glut would be
a) starting purity of the ampouled product, since it is an autocatalytic reaction, and once the dimers and trimers reach some critical level, deterioration (e.g. polymerization) can proceed quite quickly. Hence a starting purity of the least amounts of the dimers and trimers, etc. relative to a glut with higher levels, would be expected to have longer shelf life. But we are talking about variations in starting purities that, when fresh, I would expect, would give any user good results.
b) thermal history during shipment. It is quite an education to follow the progress of a shipment and to see to what levels of heat exposure a particular shipment is exposed. Over the years I have myself "visited" UPS and FedEx trucks and have been quite surprised at how hot the inside of a truck can get on a hot summer day. Indeed some of the large warehouse type sorting rooms of the courier services are not air conditioned. Travelling on a highway for hours at a time with a hot summer sun beating down on the trailer leads to sometimes very hot temperatures. I have also been in institutional receiving departments that have been like a furnace in the middle of the summer. And I have been in receiving departments during the winter, and on the coldest of days, when supplemental heat is being provided by portable space heaters and the temperature of nearby boxes seem almost too hot to touch (well a slight exaggeration, because they were not breaking out into flames, but you get my point).
You would not want to know what a receiving department is like at Cairo University in June and July, where the temperatures are over 100 deg. in the shade!
So the answer is, you don't know what has been that thermal history. Have you been lucky or have you been unlucky? Most people would not want to leave the outcome of their important experiments to the chance that it was OK.
So at least in our case, and I suspect others too, supplier applied expiration dates are a best estimate to predict when one should start thinking about ordering fresh material and tossing the old material.
Chuck
PS: In the specific case at hand, if there is even the slightest trace of a precipitate in the ampoule, just write it off and don't even try using it. But in this case, even if there was not a precipitate, that fact we are approaching two years beyond the expiration date (e.g. 11/95), it would be good cause the do the same thing.
=================================================== 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 don't have any experience with using it on Devcon 5 minute epoxy, but I have found that DMSO will dissove several epoxies quite well. Observe the usual precautions about DMSO, i.e. keep it off your hands, especially when it may contain any toxic substance.
Sincerely, Andy Andy Buechele The Catholic University of America 409 Hannan Hall Washington, D.C. 20064 (202) 319-4995 FAX: (202) 319-4469
I don't have any experience with using it on Devcon 5 minute epoxy, but I have found that DMSO will dissove several epoxies quite well. Observe the usual precautions about DMSO, i.e. keep it off your hands, especially when it may contain any toxic substance.
Sincerely, Andy
Andy Buechele The Catholic University of America 409 Hannan Hall Washington, D.C. 20064 (202) 319-4995 FAX: (202) 319-4469
George & www: These things were published over twenty years ago. What concerns me about our microscopy forum is that many of us are re-iterating believes or quoting from memory. Looking things up is considerable work and books do not include all important knowledge previously published. Here is a bit of my memory: I recall a discussion between Sjostrand and Cosslett almost 30 years ago. Sjostrand had published biological sections and claimed 10 A resolution and told the group that he was going to reduce that substantially. Cosslett then got up and with a few figures proved (?) that 10 A was as well as could be done with fixed tissues in sections. Later there were publications showing that in monolayers, cells required only three minutes of 1% (?) GA fixation. Beyond that, over fixation caused artefact - meaningless granularity, which obscured the finest details. Nothing to affect the average x30k micrograph, but at 300k its a big factor. Over fixing is not a good practise. Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
---------- } From: George C. Ruben {George.C.Ruben-at-Dartmouth.EDU} } To: jim-at-proscitech.com.au } Subject: re: storing in fixative } Date: Thursday, 17 July 1997 21:39 } } --- You wrote: } Never store tissues in GA for TEM. GA crosslinks materials, overfixing with } GA results in a coarser texture which obscures fine details at high } resolution. } --- end of quote --- } } What embedding resolution do we we have if fixation is done correctly and does } overfixing effect resolution? Have you run standards or are you just talking } about the qualitative details in a picture after post fixing and staining-- } } --George C. Ruben } Dept . Biological Sciences
Could anyone tell me what to call a lens that is located between a galvano-scanner and objective lens in the light path of a scanning laser microscope?
I'm translating a manual of a scanning laser microscope from Japanese into English. (Is it *scanning laser* microscope or *laser scanning* microscope anyway?) And I can't find a proper English term for this lens which is called "pupil projection lens" whe n translated literally.
Any suggestion is welcomed.
Thanks in advance.
Chiba Atsushi [(Mr.) -- *Chiba* is my surname] Voice: (+81) 010-045-9451
Dear Microscopists, we plann to use TEM Philips 420 for Low-dose work. We will need Low-dose system for this microscope. Unfortunately Philips said us,that they didn't produce Low-dose for this kind of microscope,already. Can you help us please, how or where we could get this Low-dose unit or how to work in low-dose conditions /to prevent destroing samples during focusing/ without this system on TEM Philips 420 ? All your responces or opinions will wery useful for us.
Thank You very much Milos
Milos Motejl Lab. of Biomembranes South Bohemian University Ceske Budejovice Czech Republic
I tested EDTA versus Chromium Potossium Sulphate and found the latter much better for ultrastructural preservation (I was looking at bone-lining-cells) as long as the pieces of bone were tiny and the decalc short (few days) I have the original reference somewhere around if a medline search doesn't do the trick. Dunno about ascorbic acid as I didn't try that.
Amanda
Miss A.J.Wilson Electron Microscope Unit St George's Hospital Medical School Cranmer Terrace Tooting London SW17 ORE Tel: 0181 725 5220 awilson-at-sghms.ac.uk awilson-at-aw.u-net.com
We have two LKB 7800 series Knifemakers that still render outstanding service. Please have a look at our publication "further modification of the LKB 7800 series Knifemaker for improved reproducibility in breaking'cryo' knives.(ref Jnl of Microscopy Vol. 168, Pt 1. Nov 1992 pp 111 - 114). The simple modifications suggested in this paper transformed our knifemaker's such that any thought of replacing them with newer 'better' units vaporised. I remember that Jan Slot, on a visit to our lab, was very impressed with the modification and performance of the knifemakers some years back
Tony Bruton University of Natal Pietermaritzburg South Africa
} } } Linda Fox {lfox1-at-wpo.it.luc.edu} 2/July/1997 06:57pm } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Does anyone know where we can get our LKB 7800B knife breaker serviced? If possible, by someone in the Chicago area? It is making sporadically bad knives and we have adjusted all the knobs by the instruction booklet. If it needs to be replaced, can anyone recommend a good one? This one has been with us for over 20 years.
Thanks, Linda Fox, Loyola Univ. Medical Center, Chicago lfox1-at-wpo.it.luc.edu
Has anyone out there had first hand experience using the freeze fracture attachment for an Edwards 306 vacuum coating unit? Please reply directly to me.
Thank you very much.
James Wesley-Smith Electron Microscope Unit University of Natal Durban, South Africa
First of all, I'd like to thank all of those who cared to respond to my question, especially Dr. Garber from SPI who gave a detailed and thoughtful response. And OK, I get the message, it's just not worth the risk.
But I have another question that perhaps you people also might have some thoughts on. If I am understanding them properly, some of the Pathologists here claim that they can tell the difference between a specimen that was delayed before putting into Glut., resulting in artifact such as swollen mitochondria, vs. poor fixation as a result of outdated glutaraldehyde, such as damaged membranes in general. Is this sort of reasoning valid?
I agree with Dr. Garber, there is a good possibility the fix has gone off. If I were you, I would order a small replacement supply and test this to make sure your problem is the fix- that way you assure yourself of not wasting your supply. Order more and throw out the old glut. once your certain it's the problem.
My two cents,
Karen Pawlowski Lab. Tech. UT Southwestern Medical Center, PhD Student UT Dallas, Dallas TX
Are you by chance the Jim Martin I know with MA State Police?
I have been working this year to help bring the new Polaroid "DMC" Digital Microscope Camera to market. If we can confirm your location, I would recommend arranging a demo of this camera which should be EXCELLENT for all but the very lowest-level fluorescence work. Also, we will be featuring the DMC at the IAI meeting at Danvers, MA if you're going.
You'll need a WIN-95/Pentium system (Mac version drivers on the way in about 30-days). Recommend at least 32mb RAM and a hard drive large enough to handle your library of images (1.3mb or 5.5mb uncompressed, depending on resolution selected). The system plugs-in directly to any software that is TWAIN compatible (that's almost anything!).
Printers? Anything you want. Dye-sub is generally the best quality, but I have had outstanding success on the ink jet printers (like HP 870, et. al.) if the best media are used. Try the "premium glossy" papers, or even the "premium" clay-coated (matte finish) papers.
Get back to me with questions: corlb-at-polaroid.com Also there are specifications on the Polaroid Web Page: www.polaroid.com, look under "Polaroid at work".
______________________________ Reply Separator _________________________________ Subject: forensic/materials science, video-digital imaging Author: John D Warren at ~575ts2 Date: 7/16/97 6:35 PM
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I am planning to purchase a video or digital camera to document, analyze, and print images of paint and fiber samples using polarized light microscopy and fluorescence microscopy. Previous threads re video or digital cameras have focused largely on black and white images obtained using SEM or TEM and printed using high-DPI inkjets or dye-sub printers.
May I ask for recommendations or comments on purchasing a system (input through output) to deal with colored images of paint cross-sections, fibers, petrographic samples, etc.? This system would allow still images to be captured, analyzed/manipulated using image analysis software (including Photoshop), embedded in reports, and printed with near photographic quality in color.
I'll be pleased to provide more details, as requested.
First of all, I'd like to thank all of those who cared to respond to my question, especially Dr. Garber from SPI who gave a detailed and thoughtful response. And OK, I get the message, it's just not worth the risk.
But I have another question that perhaps you people also might have some thoughts on. If I am understanding them properly, some of the Pathologists here claim that they can tell the difference between a specimen that was delayed before putting into Glut., resulting in artifact such as swollen mitochondria, vs. poor fixation as a result of outdated glutaraldehyde, such as damaged membranes in general. Is this sort of reasoning valid?
We have two grades of glutaraldehyde - EM & BIO, and I believe you are talking about the former.
We have done some study on storage and the polymer peak at 230nm, and found material in a sealed ampule staying alright for up to two years. But since the customer will be breaking open an ampule, take out to lab, withdraw a little, and then put it back in the refrigerator, we figured these "in's and out's" will cut the stability to some extent and put an expiry date of one year.
Assuming that your lot with an expiration of November, 1995, was not taken out and then put back into the refrig too frequently, it could be stable for at least six more months; i.e., June, 1996, if not until the end of 1996. But now it is well beyond that period and so has possibly some polymer which causes it to be less effective in fixation. You should probably consider buying a new lot.
Good luck!
DR. PARASARAN POLYSCIENCES, INC.
---------- } From: Garry Burgess {GBurgess-at-exchange.hsc.mb.ca} } To: 'Microscopy Society of America - Mailing List' {microscopy-at-sparc5.microscopy.com} } Subject: Expired Glutaraldehyde } Date: Thursday, July 17, 1997 12:40 PM } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Our lab has been using Glutaraldehyde that expired in November of 1995, } but has been kept refrigerated since then. Lately we have been noticing } problems with our fixation, and I was wondering if perhaps it is because } of this old Glutaraldehyde. Does anyone know how important these expiry } dates really are with respect to its effect on fixation????????? } } Not properly fixed, } } Garry
Along the same lines, we use the brightness of the video signal to make sure we are on the feature of interest. Our JEOL 840A has LED meters for contrast and brightness that we can monitor even in spot mode. We even used to do this on our JEOL U3, but it has been gone so long now I cannot remember exactly how we watched for it.
At 12:27 PM 7/17/97 +0300, you wrote: } Here is one addition (hopefully valuable) to the EDX-Spot Mode discussion. } } If your SEM has a specimen current meter, you can use that successfully } for checking the beam position very accurately before and during the EDX } analysis of small particles. } } The technique is very simple. You first position the spot mode beam } at high magnification, and then by moving the beam in X-Y direction } you either maximize or minimize the specimen current reading depending on } whether the average atomic number (= Z) of the particle is lighter or } heavier that that of the matrix. The great advantage in the use of the } specimen current (= absorbed electrons) for positioning the beam is that } you same time maximize the X-ray emission from the particle and minimize } the possible X-ray contribution from the matrix. This is due to the fact } that absorbed electrons "sense" the shape of the particle under the specimen } surface. } } If the specimen current stays constant during the measurement of the EDX } spectrum, then you can be absolutely certain that the beam did not leave } the particle during the measurement. However, normally there is a small } increase ( { 1 %) in the specimen current due to the contamination build-up. ---------------------------------------------------- Warren E. Straszheim 270 Metals Development, Ames Lab/ISU, Ames IA, 50011 Phone: 515-294-8187 FAX: 515-294-3091
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Nora,
We examine similar samples in our lab on occasion. I would need more information on your samples to give a more specific procedure, but here are a few tips:
a) pigments in mineral oil: try diluting sample about 1:20 in hexane, heptane, or mineral spirits (i.e., a compatible solvent), let pigments settle out by gravity overnight (if they will), otherwise spin down gently in a centrifuge. Decant off the supernatant (i.e., the mineral oil in the solvent) without losing the pigments. Add more solvent to the container and try to redisperse the pigments. With some method development, you should be able to obtain a dispersion of the pigment in the solvent with only a little of the mineral oil left. Then put a droplet of this dispersion on a suitable polished substrate (carbon?), and wick off a little of the solvent with a kimwipe.
b) pigments in an oil/water emulsion. If you mean that the sample is an oil in water emulsion like a latex, dilute the sample in water 1:20, put a droplet on the substrate, and wick off the water. In this way you should be able to separate the 'oil' and the pigments enough to pick out the pigments and image or analyze them.
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, Dear friends from the list,
We have just received an enquiry about SEM analysis for: a) pigments particles dispersed in mineral oils b) pigments particles in oil-water emulsions
Does any of you have any experience in that subject? Unfortunately, we do not count with equipment for sample preparation other than the sputter-coater and carbon evaporator.
Any help will be very welcome.
Thanks in advance,
Nora Pratta Centro Regional de Investigacion y Desarrollo Santa Fe - Argentina
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Chiba Atsushi wrote: :Could anyone tell me what to call a lens that is located between a galvano-scanner and objective lens in the light path of a scanning laser microscope? : :I'm translating a manual of a scanning laser microscope from Japanese into English. (Is it *scanning laser* microscope or *laser scanning* microscope anyway?) And I can't find a proper English term for this lens which is called "pupil projection lens" wh e : :Any suggestion is welcomed. : :Thanks in advance.
There are several words that can be used, but "transfer lens" is very common. Also used are "telecentric lens" and "relay lens."
best regards mark
Mark W. Lund, PhD Director } } Soft X-ray Web page http://www.moxtek.com { { MOXTEK, Inc. 452 West 1260 North Orem UT 84057 801-225-0930 FAX 801-221-1121 lundm-at-xray.byu.edu
"The state is good at simple tasks, like killing people and seizing their wealth. It has far more trouble reaching inside individuals and making them good." Doug Bandow
In response to Garry Burgess's question regarding swollen mitochondria sometimes referred to as "popcorn" mitochondria, the only time I have seen them perfectly preserved was after whole body perfusion of fixative through the heart.
It is likely that pathologists do not have tissue preserved in this way from diagnosing human diseases. I guess for argument sake, Garry, if I were in your shoes I would ask the pathologist (tactfully) weather the swollen mitochondria could be the result of the pathology and not poor fixation.
For perspective, though, I should tell you that I once worked in the Anatomy Department at Loma Linda University and the professor there didn't believe in purified glutaraldehye. He used the 25% stuff and kept it under the lab bench. His belief was that the older it got the better....something to do with the ratio of dimer to trimer. If you want to check his publications, I believe his name was Robert Schultz.
{HTML} {FONT SIZE=+1} In response to Garry Burgess's question regarding swollen mitochondria sometimes referred to as "popcorn" mitochondria, the only time I have seen them perfectly preserved was after whole body perfusion of fixative through the heart. {/FONT} {FONT SIZE=+1} {/FONT}
{P} {FONT SIZE=+1} It is likely that pathologists do not have tissue preserved in this way from diagnosing human diseases. I guess for argument sake, Garry, if I were in your shoes I would ask the pathologist (tactfully) weather the swollen mitochondria could be the result of the pathology and not poor fixation. {/FONT} {FONT SIZE=+1} {/FONT}
{P} {FONT SIZE=+1} For perspective, though, I should tell you that I once worked in the Anatomy Department at Loma Linda University and the professor there didn't believe in purified glutaraldehye. He used the 25% stuff and kept it under the lab bench. His belief was that the older it got the better....something to do with the ratio of dimer to trimer. If you want to check his publications, I believe his name was Robert Schultz. {/FONT} {FONT SIZE=+1} {/FONT}
In response to Garry Burgess's question regarding swollen mitochondria sometimes referred to as "popcorn" mitochondria, the only time I have seen them perfectly preserved was after whole body perfusion of fixative through a living and pumping heart.
Obviously, it is not likely that pathologists have tissue preserved in this way from living humans. I guess for argument sake, I would ask the pathologist weather the swollen mitochondria and other membrane defects could be the
result of the pathology and not poor fixation.
On the other hand, dead cells such as cuticle and cortical cells in hair have well defined cell membrane complexes that look well preserved even if you don't fix the hair--although cell organelles like mitochondria are missing.
For perspective, I should tell you that I once worked in the Anatomy Department at Loma Linda University and the professor there didn't believe in purified glutaraldehye. He used the 25% stuff and kept it under the lab bench. His belief was that the older it got the better....something to do with the ratio of dimer to trimer. I'm not a chemist, so I don't know. I'm sure the Ph.D.'s at the companies who sell the "good" stuff have their point of view.
} ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } ----------------------------- } -----------------------------------------. } } First of all, I'd like to thank all of those who cared to respond to } my } question, especially Dr. Garber from SPI who gave a detailed and } thoughtful response. And OK, I get the message, it's just not worth } the } risk. } } But I have another question that perhaps you people also might have } some } thoughts on. If I am understanding them properly, some of the } Pathologists here claim that they can tell the difference between a } specimen that was delayed before putting into Glut., resulting in } artifact such as swollen mitochondria, vs. poor fixation as a result } of } outdated glutaraldehyde, such as damaged membranes in general. Is } this } sort of reasoning valid? } } Just curious, } Garry
{HTML} {FONT SIZE=+1} ----------------------------------------------------------------------- {/FONT} {BR} {FONT SIZE=+1} The Microscopy ListServer -- Sponsor: The Microscopy Society of America {/FONT} {BR} {FONT SIZE=+1} To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com {/FONT} {BR} {FONT SIZE=+1} -----------------------------------------------------------------------. {/FONT} {FONT SIZE=+1} {/FONT}
{P} {FONT SIZE=+1} In response to Garry Burgess's question regarding swollen mitochondria {/FONT} {BR} {FONT SIZE=+1} sometimes referred to as "popcorn" mitochondria, the only time I have seen {/FONT} {BR} {FONT SIZE=+1} them perfectly preserved was after whole body perfusion of fixative through {/FONT} {BR} {FONT SIZE=+1} a living and pumping heart. {/FONT} {FONT SIZE=+1} {/FONT}
{P} {FONT SIZE=+1} Obviously, it is not likely that pathologists have tissue preserved in this way {/FONT} {BR} {FONT SIZE=+1} from living humans. I guess for argument sake, I would ask the pathologist {/FONT} {BR} {FONT SIZE=+1} weather the swollen mitochondria and other membrane defects could be the {/FONT} {BR} {FONT SIZE=+1} result of the pathology and not poor fixation. {/FONT} {FONT SIZE=+1} {/FONT}
{P} {FONT SIZE=+1} On the other hand, dead cells such as cuticle and cortical cells in hair have {/FONT} {BR} {FONT SIZE=+1} well defined cell membrane complexes that look well preserved even if you {/FONT} {BR} {FONT SIZE=+1} don't fix the hair--although cell organelles like mitochondria are missing. {/FONT} {FONT SIZE=+1} {/FONT}
{P} {FONT SIZE=+1} For perspective, I should tell you that I once worked in the Anatomy {/FONT} {BR} {FONT SIZE=+1} Department at Loma Linda University and the professor there didn't believe {/FONT} {BR} {FONT SIZE=+1} in purified glutaraldehye. He used the 25% stuff and kept it under the lab bench. {/FONT} {BR} {FONT SIZE=+1} His belief was that the older it got the better....something to do with the ratio of {/FONT} {BR} {FONT SIZE=+1} dimer to trimer. I'm not a chemist, so I don't know. I'm sure the Ph.D.'s at the {/FONT} {BR} {FONT SIZE=+1} companies who sell the "good" stuff have their point of view. {/FONT}
{P} {BR} {FONT SIZE=+1} Michael Mead {/FONT}
{P} __________________________________________________________________________________ {BR} Garry Burgess wrote: {BLOCKQUOTE TYPE=CITE} ------------------------------------------------------------------------ {BR} The Microscopy ListServer -- Sponsor: The Microscopy Society of America
{P} First of all, I'd like to thank all of those who cared to respond to my {BR} question, especially Dr. Garber from SPI who gave a detailed and {BR} thoughtful response. And OK, I get the message, it's just not worth the {BR} risk.
{P} But I have another question that perhaps you people also might have some {BR} thoughts on. If I am understanding them properly, some of the {BR} Pathologists here claim that they can tell the difference between a {BR} specimen that was delayed before putting into Glut., resulting in {BR} artifact such as swollen mitochondria, vs. poor fixation as a result of {BR} outdated glutaraldehyde, such as damaged membranes in general. Is this {BR} sort of reasoning valid?
{P} Just curious, {BR} Garry {/BLOCKQUOTE} {/HTML}
Does anybody out there have experience with the {bold} Pixera digital camera system {/bold} . Would appreciate your comments on the image quality, the possibility of converting the images to publication quality prints on a good printer and the level of technical support provided by the company.
My apologies to the list. I sent a private email that seems to have gotten posted to the MDS mail server. (And Eudora is configured not to do that ... the net gremlins at work?)
Phil
****be famous! send in a tech tip or question*** Philip Oshel Technical Editor, Microscopy Today Station A PO Box 5037 Champaign, IL 61825-5037 oshel-at-ux1.cso.uiuc.edu
Would anyone be interested in a used LKB Knifemaker (7800) with cover + six boxes of LKB glass? Fred says it is in excellent condition and that the price would be in the $2500-$3000 range.
If interested, please contact Fred directly at fredl-at-awod.com (Fred G. Lightfoot) or call Fred at (803) 856-8613.
Thanks and regards, Don Cox, Goldmark Biologicals, goldmarker-at-aol.com
Would anyone have an interest in purchasing a used LKB Knifemaker (7800) with six boxes of LKB glass? I think the price will be in the $2500 range. Fred says it is in excellent condition.
If interested, please contact Fred Lightfoot at
fredl-at-awod.com (Fred G. Lightfoot)
or call Fred at (803) 856-8613
Regards, Don Cox Goldmark Biologicals goldmarker-at-aol.com
We section hard tissues containing synthetic biomaterials. While we usually follow a conventional EDTA decal. route, we also sometimes apply a little EDTA to the block face (having embedded in LR White) to decal. "in situ" while sectioning. This alternative seems to work, but as we are using diamond knives it may be purely psychological! I have never seen it written up.
Best wishes, Paul
Dr Paul V. Hatton Lecturer in Biomaterials School of Clinical Dentistry University of Sheffield Claremont Crescent SHEFFIELD S10 2TA
Tel. (0114) 271 7938 Fax. (0114) 2665326 or 2797050
First of all, I'd like to thank all of those who cared to respond to my question, especially Dr. Garber from SPI who gave a detailed and thoughtful response. And OK, I get the message, it's just not worth the risk.
But I have another question that perhaps you people also might have some thoughts on. If I am understanding them properly, some of the Pathologists here claim that they can tell the difference between a specimen that was delayed before putting into Glut., resulting in artifact such as swollen mitochondria, vs. poor fixation as a result of outdated glutaraldehyde, such as damaged membranes in general. Is this sort of reasoning valid?
For an inexpensive target: Obtain a piece of OFHC Cu foil, cut circle slightly larger than target, unscrew the existing target, fold/crimp the foil over the existing target. Even if you have to get the foil from Alfa/Aesar or the like, it should cost only about $100, compared to the several hundred+ vendors want for a "real" target.
Will be curious how well you are able to sputter Cu with the Polaron.
In my previous life doing TEM we would develop and wash negatives for about 5 minutes, then rinse in methanol. Air dry or even a blow dryer on low got us a dry negative in 15-20 minutes. I seem to remember that there was a little clouding of the negative and you can't get too anxious with the hair dryer.
Now, with negative, flatbed scanners one could scan the negative (10 minutes) and print an image on transparency paper with an inkjet (Epson 1440 dpi or a dyesub printer). Haven't tried this with TEM negatives but I know people who do.
Seems like about one hour would do it.
What about a positive TEM film that you could project directly.
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Randy,
Sometimes when I want to make a transparency for seminars, or workshops or whatever, I put the EM negative into the enlarger and project it to a small 2X2" size on to yet another piece of EM film that I've cut in half for this purpose. (you need a special enlarger lens to get this small!). Then I cut it to size and mount it in a glass slide mount, and voila, I have supersize black and white slides, with good resolution and contrast. (at least better than 35mm projection slides) But sometimes in the past, if I tried to rush things, I notice that the image turned brown. To fix this problem though, I simply re-fix and re-wash this image, and the brown discoloration (which is probably some residual silver halide and fix) is removed, and all is well again.
Garry
} ---------- } From: rtind-at-siu.edu[SMTP:rtind-at-siu.edu] } Sent: 14 July, 1997 14:51 } To: Garry Burgess } Subject: RE: Rush Lab } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thanks ever so much for all those comments on "burn" marks. As a newbie, I'm grateful for your help. Now I've got a question on sample preparation.
We're currently doing many different types of sample preparation in an "inherited" Edwards Auto 306, which works. But because we do so many different things in it, we're always having to rearrange its "innards" and this is turning out to be a bottleneck. So I was wondering about doing some of the preparations in a separate unit.
In particular, I'm interested in purchasing equipment for plasma etching and carbon coating. (IF--or WHEN-- we can afford it, that is!) So I'm interested in hearing peoples' recommendations, good and bad experiences and so forth. I also have no idea (yet) how much these things cost.
We do the etching to get a better look at those small inorganic particles embedded in a polymer matrix that I mentioned in the EDX spot mode conversation.
(And if vendors would like to contact me, you are welcome to do so! Please do this directly and not via the listserver.)
Title: Optical Microscopy and Imaging in the Biomedical Sciences
When: October 8 - October 16, 1997
Where: Marine Biology Laboratory, Woods Hole, MA, USA
Tuition: $1950 (Includes room and board)
Application Deadline: August 5, 1997
Admission application and information: Carol Harnel, Admissions Coordinator Marine Biological Laboratory 7 MBL Street Woods Hole, MA 02543-1015 (508) 289-7401 Internet: admissions-at-mbl.edu WWW: http://www.mbl.edu
Course Director: Colin S. Izzard, State University of New York -at- Albany Phone: [518] 442 - 4367 EMail: csizzard-at-csc.albany.edu
Course Description:
For Whom: Designed primarily for research scientists, physicians, postdoctoral trainees and advanced graduate students in animal, plant, medical and material sciences. Non-biologists seeking a comprehensive introduction to microscopy and video-imaging will benefit greatly from this course as well. There are no specific prerequisites, but an understanding of the basic principles of optics is desirable. Limited to 24 students.
The eight day course consists of lectures, laboratory demonstrations, exercises and discussions that will enable the participant to obtain and interpret microscope images of high quality, to perform quantitative optical measurements, and to produce photographic and video records for documentation and analysis.
Topics to be covered include: principles of microscope design and image formation bright field, dark field, phase contrast, differential interference contrast, interference reflection, and fluorescence microscopy confocal scanning microscopy and image deconvolution digital image restoration and 3-D reconstruction video imaging, recording, enhancement, and intensification analog and digital image processing and analysis fluorescent probes and ratio-imaging laser tweezers and laser scissors
Applications to live cells will be emphasized; other specimens will be covered as well.
Students will have direct hands-on experience with state-of-the-art microscopes, video cameras, recorders and image processing equipment provided by major optical and electronics companies. Instruction will be provided by experienced staff from universities and industry.
Students are encouraged to bring their own biological (primary cultures, cell lines, etc.) and material specimens and to discuss individual research problems with the faculty.
I've seen someone use this technique to decalcify and section tissue in epon years ago. It worked fine. I also don't know where any written info can be found about it.
By epon, I mean specifically medcast from Ted Pella Inc., I understand this particular media has been discontinued, but I don't think the EDTA decalcification is limited by the media. It only decalcified a few mm-s of surface tissue.
Karen Pawlowski Lab Tech UT Southwestern Med. Ctr. Student/ UT Dallas
On Sun, 20 Jul 1997, P.V.Hatton wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } We section hard tissues containing synthetic biomaterials. While we } usually follow a conventional EDTA decal. route, we also sometimes } apply a little EDTA to the block face (having embedded in LR White) } to decal. "in situ" while sectioning. This alternative seems to } work, but as we are using diamond knives it may be purely } psychological! I have never seen it written up. } } Best wishes, Paul } } } Dr Paul V. Hatton } Lecturer in Biomaterials } School of Clinical Dentistry } University of Sheffield } Claremont Crescent } SHEFFIELD S10 2TA } } Tel. (0114) 271 7938 } Fax. (0114) 2665326 } or 2797050 }
This is prompted by Cynthia Bennett's recent request in regard to plasma etching units. A few years ago, we were briefly interested in such things, particularly in how to remove the surface from a polymeric material without damaging the underlying substructure. However, for our purposes we found that plasma etching, ion mills, etc., would be far too destructive - not only would there be heating problems, but also all those ions running wild would tend to cause a lot of chemical damage.
We had just got round to trying out ATOMIC OXYGEN, generated in a radio frequency discharge, and the first result or two seemed promising, and then with a great bureaucratic reshuffle the owner of the equipmment pulled up sticks and went elsewhere. Does anyone know where such equipment might be obtainable at reasonable cost?
If anyone has experience with this sort of thing, I would be pleased to hear from them.
Thanks in advance,
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
Now that this whole question of expired Glut. and fixation has come up, I'm wondering if perhaps my 2.5% solution that we use here is perhaps suboptimal, and whether or not we might be better off using a strong glutaraldehyde concentration such as 4% for our routine fixation.
Is there anyone else out there fixing human tissue routinely? I would be interested in what concentration of glutaraldehyde that you people are using.
Now that this whole question of expired Glut. and fixation has come up, I'm wondering if perhaps my 2.5% solution that we use here is perhaps suboptimal, and whether or not we might be better off using a strong glutaraldehyde concentration such as 4% for our routine fixation.
Is there anyone else out there fixing human tissue routinely? I would be interested in what concentration of glutaraldehyde that you people are using.
Hello all, I was just asked for answers regarding Canada Balsam, which I hope some member of the group could assist me with, they are..
Questions regarding Canada Balsam, 1) When was it first used for microscopy? 2) How long does it last before degrading? 3) What if any are the aging effects? 4) Does it interfere or affect the sample in any way?
The question was actually asked for a different application other than microscopy, but I thought it would be an interesting discussion none the less.
Thanks
David Dr. David C. Bell Room 13-1018 E-Mail: dcb-at-MIT.EDU Center for Mat. Sci. and Eng. PH: (617) 253-3317 Massachusetts Institute of Technology FAX: (617) 258-6478 77 Massachusetts Ave, Cambridge, MA 02139-4307
We are looking for a used High-Voltage Tank for a JEOL l00. Can anyone help us? Please answer us direct. Thank you. Peter Stolzenberg PESTO INC. pesto-at-erols.com
To all: Can anyone help us getting a used Jeol l00 tank. We would appreciate any leads. Please E-Mail us direct. Thank you! Peter Stolzenberg,Pesto Inc. P.O. Box 648, GWYNEDD VALLEY ,PA 19437 215-699-6160 FAX215-699-5275 E-Mail: pesto-at-erols.com
} Dear Microscopists, } we plann to use TEM Philips 420 for Low-dose work. We will need Low-dose } system for this microscope. Unfortunately Philips said us,that they didn't } produce Low-dose for this kind of microscope,already. } Can you help us please, how or where we could get this Low-dose unit or } how to work in low-dose conditions /to prevent destroing samples during } focusing/ without this system on TEM Philips 420 ? } All your responces or opinions will wery useful for us. } } Thank You very much } Milos } } Milos Motejl } Lab. of Biomembranes } South Bohemian University } Ceske Budejovice } Czech Republic } } motejl-at-paru.cas.cz } tel. 042-038/7775485 } ................................
Philips used to produce free-standing Low Dose modules for their 400 series TEMs - off hand, I don't recall the part number, but if you check around you might find a used one available.
Actually, the easiest approach to low dose is the simplest - work at low magnification. I used to be an applications specialist for Philips, and had the time to experiment with different ways of using a TEM. With a little pre-calibration and experiment, I could repeatedly, with a single micrograph, record the 0.9 nm lattice spacing of crocidolite with the TEM at 10,000 x magnification - the point being that you can record quite high resolution images at low magnification, which gives you sufficient intensity at the film to record the image but considerably reduces the dose to specimen, compared with operating at high mag. Note, that this approach fails if you take the mag into the LM mag range, as the whole optics of the microscope changes at this point, one consequence being a drastic drop in resolution.
You can further reduce the dose to the specimen if you have a STEM unit. Basically the idea is to operate the TEM column in a high mag/high res mode and very low beam intensity but use the STEM unit to image the specimen at low mag/low res, so as to locate a the region of interest.
With a standard STEM set up, you can only image a small part of the field of view. However, if you have a computer connection to allow you to control beam position, it is straightforward to produce a short routine that will step the illuminated area across the whole area covered by the film. With this approach, I have 'easily' recorded diffraction patterns and images from parafin wax.
With the same sort of computer control set up, it is not much more difficult to duplicate most of the functions of the low dose unit, except that you can't change the illumination focus - but that can be done manually at the appropriate step in the sequence.
Garry Burgess wrote: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Now that this whole question of expired Glut. and fixation has come up, } I'm wondering if perhaps my 2.5% solution that we use here is perhaps } suboptimal, and whether or not we might be better off using a strong } glutaraldehyde concentration such as 4% for our routine fixation. } } Is there anyone else out there fixing human tissue routinely? I would } be interested in what concentration of glutaraldehyde that you people } are using. } } Garry
Here in the EM Facility at Boston Medical Center, we use 2.5% glut in 0.2M cacodylate buffer for all human specimens; the fix lasts for several months in the fridge and yields very high quality results if the tissue is fixed promptly at the biopsy site. If a surgical specimen has been held overnight in the cold and is then sampled for EM in the morning, the preservation suffers but is still suitable for diagnosis.
Dr. Tom Christensen Director, EM Facility Boston Medical Center Boston, Mass
I do not know if you have gotten any other replies on this. We do failure analysis on integrated circuits. There are two types of "short circuits". One is a resistive short, and the other is a leakage path where hole-electron recombination occurs. Resistive shorts can be located with liquid crystal methods that sense the thermal dissipation. (There are other, more complex, methods, but we use liquid crystal extensively) Leakage paths can be located with systems based on "night vision" technology that was developed for the military. Hole-electron recom- bination releases the excess energy as photons, and the light amplification allows the imaging of the emission site.
Darrell Miles IBM Microelectronics Test and Analytical Services http://www.chips.ibm.com/services/asg
A tentative okay has been given for a session on corrosion casting for the 1998 meeting in Atlanta. We would urge any of our Mercox users who are interested in presenting a paper to contact us at Ladd Research or Dr. Fred Hossler at:
Dr. Fred Hossler Professor of Anatomy East Tennesse State University Johnson City, TN 37614 e-mail SEMTEMman-at-aol.com
Methadology is of particular interest.
We would also like to have some opinions on the advantages/disadvantages of using clear, blue or red mercox for casting.
In March 1997 we requested protocols and references for staining semi-thin sections of vertebrate tissues embedded in plastic; our original standard stain was toluidine blue.
We received 14 responses to this request, which we have edited to a ~7 page file. Responses included stains such as hematoxalin, alcian blue, eosin, Mayer's mucicarmine, methyl green, methylene blue/azure II, basic fuschin, and Stevenel's blue. A copy of the file is available upon request by e-mail. We have begun using a polychrome stain (see Van Reempts and Borgers, 1975, Stain Tech. 50:19-23) with pleasing results.
Christine Roy and David Hall Albert Einstein College of Medicine Bronx, NY 10461
Dr. Steven Barlow EM Facility/Biology Department 5500 Campanile Drive San Diego CA 92182-4614 phone: (619)594-4523 fax: (619) 594-5676 email: sbarlow-at-sunstroke.sdsu.edu website: http://www.sci.sdsu.edu/EM_Facility
I am interested in finding out what people are using for 4"x5" SEM film. I have used Polaroid Type 55 P/N and Kodak type 4427 Commercial Film. At $1.50 to $2.00 per exposure, both are a little too expensive for student use (i.e. low good photo to bad photo ratio). Can anyone recommend a less expensive alternative? My predecessor used to buy 200 foot rolls of surplus aerial photography film (4 or 5 inches wide) which he would cut to the proper length. Final cost was less than $0.05 per sheet. Any similar suggestions?
Bob
Dr. Robert R. Wise Department of Biology University of Wisconsin-Oshkosh Oshkosh, WI 54901
(414) 424-3404 tel (414) 424-1101 fax wise-at-uwosh.edu
Nora Pratta Centro Regional de Investigacion y Desarrollo Santa Fe - Argentina Inquired about doing SEM/BSE/EDX on particulates in oil and oil/water mixture.
If the oil or oil water mixture can be 'frozen' at liquid nitrogen temperatures by rapid freezing in a metal block or propane jet freezer th= en transferred to a cryo ultramicrotome where you could prepare frozen sections for examination on an SEM cold stage. =
This is a very expensive but technically superb method. An alternate woul= d be to freeze dry the particulates out of the section onto a carbon substrate or other suitable substrate. You will lose some of the distribution information in the X-Y plane but should resolve the Z distribution represented by your sections themselves.
There are some commercial labs with this equipment and some private companies that may collaborate if the subject interests them.
You may contact me off line or visit the Web Site: =
http://www.RMC-Scientific.com/microtomes/ =
We are a commercial manufacturer of all of the instruments listed above.=
Steve Miller Director of Sales RMC 3450 S. Broadmont, Suite 100 Tucson, AZ 85713 Tel 520-903-9366 Fax 520-903-0132
"Inter/Micro" is a meeting, now in its 49th year, held annually in Chicago and hosted by the McCrone Research Institute (McRI). I am attending the meeting as I have almost every year for the past 25 or so. However it is a meeting little known to those without regular contact with McRI. As a way of introducing this under-attended meeting to a wider audience I thought the readers of this list might find it informative to have a summary of the technical highlights and other goings-on at Inter/Micro. If I can, I'll write a daily summary every evening and post it to this list. Otherwise I'll just summarize and post as time permits. I'll mention two or three of those papers I found most interesting each day.
This year approximately 60 technical presentations are scheduled. A single meeting room is used and there are no parallel sessions. One has an opportunity to hear all of the papers and I find it quite valuable to sit in on papers which might not be in an area of my own personal focus. This "cross-fertilization" of ideas and techniques is often the most enlightening thing that I experience at technical meetings and I am glad they've continued the tradition at Inter/Micro. There will also be an exhibition of products and equipment, as is usual at such meetings. A full program is available at McCrone's web page, http://www.mcri.org. I won't duplicate that here but I'll just mention that the themes for the various sessions for the week are as follows: Monday: General Microscopy Tuesday am: Instrumentation Tuesday pm: Techniques Wednesday: History and Art Wednesday Eve: Dinner in Conjunction with the State Microscopical Society of Illinois (SMSI) Thursday: Forensic Microscopy Friday: A Tutorial on Dispersion Staining
The highlights (for me) of Monday's program included the following.
The program was opened with a talk by Brian Ford on "Crytosporidium, a New Threat from Water Supplies." Brian discussed how Cryptosporidium is representative of a "new" class of microorganisms threatening our modern society. Of course, it is not new at all however a combination of factors is working together to make many existing organisms newly hazardous. These factors include new practices and factors coming with technological advances such as the wearing of contact lenses which provide a previously non-existing environment in which certain organisms can thrive. Another factor is the emerging resistance of some organisms to existing treatments. Previously "eradicated" threats are reemerging as "new" threats again. In the case of Cryptosporidium, we have an organism causing severe but usually non-fatal intestinal distress that we are immune from after the initial exposure and bout of sickness. But it has become so "rare" as a contaminant that few of us received the immunizing infection early in our lives, leaving large segments of the population subject to infection when water treatments fail or other sources of the protozoa present to the population.
Another highlight today was a paper by John Wuepper of Whirlpool entitled "Problem Solving via Analytical Microscopy: What is it Really Worth?" At Inter/Micro we have on many occations over the years lamented the under-appreciated and under-valued status of the work we do on behalf of industry and society at large. John suggested an excellent technique that we can use if we revise our thinking and the presentation of our work to the corporations or agencies we serve. He demonstrated with several examples that the "leverage" obtained from an investment in microscopical problem solving is often enormous, in the range of 25 to 5,000 in the examples he gave. By "leverage" he meant the ratio of the cost of the process to the savings enjoyed by the company as a result of the work done. The "investments" may range from a few hundred dollars to a hundred thousand or more, but the return and leverage is enormous. In one instance he cited microscopy saved a company more than $500,000,000 dollars. (Those figures will catch the eye of even the most myopic bean-counter - my editorial, not John's!) A lively discussion followed John's presentation and it is clear that his approach of communicating with our host-employers in terms they understand and respond to is critical to the success and growth, sometimes just the survival, of a microscopy laboratory.
Jan Hinsch or Leica gave one of the most beautifully illustrated as well as educational talks of the day when he spoke on "What Pleurosigma can Tell the Microscopist." Pleurosigma Angulatum, a species of diatom, has long been used as a microscope test object for evaluating the quality of higher numerical aperture objectives. Jan's talk was one of those wonderful half-hours where an audience gets to enjoy not only an aestheticly beautiful talk but one that, through the instructional insight of the author, makes crystal clear some technically difficult fundamental principles. In the case of today's talk, those principles dealt with the theoretical resolution limits of the light microscope and I, along with many others in the audience I think, came away with with a vastly better understanding of what had heretofor been a baffling subject.
Another fascinating talk today illustrated the diversity of topics we regularly enjoy at Inter/Micro. Ryan D. Tweney of the Department of Psychology at Bowling Green State University spoke on "Cognition and the Microscope." Ryan discussed and illustrated many of those murky processes that stand between knowledge or observation on the one hand and understanding on the other. It was apparent that he was only able to scratch the surface of a subject which we would all benefit from knowing much more about and I, for one, hope we'll see him back regularly in the future.
I'll try to come back tomorrow with another update! Right now I've got friendships to rekindle and think I hear the calling of a lonely brew with my name on it.
Thank you to those of you who replied regarding LM mounting medium. It appears as if oxidants in the medium are the main culprits. I am listing these replies below.
One point worth mentioning about the common use of epoxy resins as a coverslip mountant is that its refractive index is not 1.5 (I'm not sure of its exact value). One may be able to get away with it working at low numerical apertures, but it will take its toll at 0.65 and above.
Thanks again!
James Wesley-Smith EM Unit University of Natal Durban, South Africa
I learned a trick from a DuPont-Sorvall rep (that tells you how long ago that was!) that retards or eliminates ALL fading caused by oxidants in the mounting medium. Add 1-2% BHT (the preservative used in bologna, hot dogs, etc.) to any mounting medium. -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- I have been using ENTELLAN which has ND20 1.49-1.5 and gives excellent preservation of toluidine blue stained plant tissue for at least 4 years. I get it from Electron Microscopy Sciences who have a good list of mounting media with refractive indicies in their catalogue. -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- Hello fading is the problem. We use these toluidine blue sections for histology courses and we haven't find yet the no-fade mounting medium. Our best choice is DEPEX, manufactured by GURR. Avoid EUKITT, fading occurrs in a matter of hours. Some collegues have used cured epon, but it is a time consuming process and fading does occur. -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- I think the fading is caused by oxygen (which very easily diffuses through hydrophobic media like resins). We routinely leave such preparations uncovered, and add a drop of immersion oil and coverslip to photograph. This prep is easily soaked off in xylene to restain if necessary. -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- I don't know about the refractive index, but I've used Epoxy resin as a mountant quite successfully. It doesn't seem to fade Toluidine Blue semi-thins.
DePeX is not too bad but I have had some fading over long periods. -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-
I have been working with plant material embedded in Spurr resin for many, many years. One micron sections were stained with toulidin blue or other specific stains and permanent mounted with Permount, Fisher Scientific, and no fading for decades. I have not seen any fading using DePeX, but Spurr resin sections usually get very wrinkeled.
-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- We use the following protocol for toluidine blue (TB) staining and permanent mounting: Frozen or dewaxed abd hydrated paraffin sections 0.1% TB in acetate or phosphate buffer (generally at pH 2-3 for sulfated glycosaminoglycans) 5 min Rinsing in buffer Precipitation with a 6:1 mixture of 2% aqueous KI and Kferricyanide 2-3 min Mount with a drop of 25% aqueous gum arabic containing 2% fructose without coverslip! Form a layer of te mounting medium using a glass rod. Let the gum arabic layer dry at room temperature in horizontal position (it takes generally one night). The refractive index of the dried gum arabic is practically identical to that of the glass. Mount in DPX or Canada using a coverslip. This procedure is good to produce permanent metachromatic staining. Dimethylmethylene blue (DMMB) is a better metachromatic dye (Aldrich Co, or SERVA) The protocol is similar, except the poststaining stabilization. For this purpose, 2% aqueous ammoniummolybdenate is used. DMMB is a very strong metachromatic staining. It is very useful for mast cells, cartilage, sulfomucins. In many cases, we use it successfully in 0.05 or 0.01% aqueous solution for 5-10 min.
For more inforations, see Modis, L.: Organization os the Extracellular Matrix: A Polarization Microscopic Approach. CRC Press, Boca Raton, 1991. Chapter 12.
Kindly forgive me if these references have already been posted, but the answers to many of these processing protocol queries can also be found in a series of articles by Coetzee and van der Merwe, viz.
J Coetzee and CFvan der Merwe (1984) Extraction of substances during glutaraldehyde fixation of plant cells. Journal of Microscopy 135, Pt2, pp 147-158.
J Coetzee and CFvan der Merwe (1985) Penetration rate of glutaraldehyde in various buffers into plant tissue and gelatin gels. Journal of Microscopy 137, Pt2, pp 129-136.
J Coetzee and CFvan der Merwe (1986) The influence of processing protocol on the ultrastructure of bean leaf cells. South African Journal of Botany, 52, pp 95-99.
These articles are compulsory reading for our trainee microscopists, since they dispell many processing 'myths'.
James Wesley-Smith EM Unit University of Natal Durban, South Africa
We want to analyse the intensity of ED-patterns of organic specimen. For this we used ELD, a program which is included in the CRISP packages from Calidris. Now we want to compare the intensity data from ELD with the intensity getting from other programms. Does anybody knows some programs, which we can used for intensity estimation? Can you tell something about prices and the possibility to get such programms!
Hans Kothe Working group Dr. Voigt-Martin Universit=E4t Mainz =20
25th Scottish Microscopy Group Symposium (First Circular)
Stakis Dunblane Hotel, Dunblane. Wednesday 12 November 1997.
This the SILVER Scottish Microscopy Symposium will take place at the=20 above venue and the Organising Committee have arranged a Scientific=20 Programme which we hope will appeal to as many microscopists as=20 possible. Also a celebratory meal will mark this anniversary.
The following topics have been selected:
Environmental Scanning Electron Microscopy - Dirk van der Vall,=20 Eindhoven, Holland.
Advances in Confocal Microscopy - Tony Wilson, Oxford, England
Stereology - Vyvyan Howard, Liverpool, England
Cryo/Immunocytochemistry - Jeremy Skepper, Cambridge, England
These invited talks will be interspersed with short presentations. We=20 would welcome offers of short (10-15 minute) talks which deal with any=20 aspect of microscopy and in particular electron microscopy. There is an=20 abundance of useful techniques and protocols in daily use; if you think=20 that you have something which others could adopt or benefit from,=20 please send us your name and a brief title for your presentation to Ian=20 Roberts at {irober-at-scri.sari.ac.uk}
These meetings are enjoyable, interesting and useful and an=20 opportunity to meet and share ideas with fellow microscopists. The cost=20 will be =A320 (pounds).
Celebratory Dinner (evening)
To mark the 25th Anniversary of these meetings, we hope to arrange an=20 evening dinner to which all delegates and partners are invited to attend.= =20 The separate cost of this will be =A325.00/head, and a favourable rate of= =20 =A3100 per night/ per couple for dinner, bed and breakfast at the hotel has= =20 been negotiated. If you wish to attend this evening function, please=20 contact Martin Maxwell at {MARTIN.MAXWELL-at-BBSRC.AC.UK}
A second circular will be sent in the future given details of all talks. Al= so=20 there is a web page at http://www.abdn.ac.uk/~nhi691/smg97.htm that=20 gives more information.
Kevin Mackenzie Tillydrone E.M. Unit University of Aberdeen Tillydrone Avenue Aberdeen AB9 2NT
Tel 01224-272847 Fax 01224-272396 Web site- http://www.abdn.ac.uk/~nhi691/
---------------------- Kevin Mackenzie k.s.mackenzie-at-abdn.ac.uk
We used to commonly use a Kodak film called Ektapan in 4x5 size. It is developed in D-76. usually, although other standard film developers could also be used. The advantage is that it's much cheaper than Polaroid PN 55, at about $20 per 25-sheet box (last time we checked the price). The disadvantages, of course, are that a developing set-up and darkroom are required and you don't get an automatic study print.
Actually, it would seem that any common 4x5-inch film could be adapted to SEM use, depending upon contrast requirements. T-Max 100 or 400, Plus-X Pan, Ilford, or Agfa films, etc., all should work. All of these films should be readily available and all use a variety of common developers.
Hope this helps.
Randy Tindall Center for Electron Microscopy Southern Illinois University at Carbondale
Since my initial request for the x-ray wavelengths file I have since found a Microsoft Access database file which was originally provided by John Donovan (UC Berkeley). I have also received request for posting the file for FTP (... altho one reply indicated the original "Fiori" file was available at ftp://www.anc.anl.gov ...), and the John has since indicated his database which also includes higher order lines is generally available. I can make John's MDB file and my Excel (XLS v2.1) file available via "anonymous" FTP. My XLS file is only slightly different, i.e., it has been sorted by "Z" and "N" ... Lastly, a word about the FTP site. It is actually a magneto-optical drive and used for archiving image files. As a MO drive it will offer these specific files only until this cartridge fills and I have to put in another (approximately 1 week). I can make these files available in the future upon request.
You can point your browser at ftp://whitewater.uoregon.edu/share/cameca/
or FTP anonymous to whitewater.uoregon.edu
and look into the "/share/cameca/" directory
and find John's original Access file (xray.mdb, 704kb) ... you can also find the Excel file I created with it, but it is 2Mb. I couldn't get Access to save the file as XLS ... it created the file but the cells were empty (?) ... I copied and pasted instead ... you may not be able to C&P if you don't have enuf memory. Let me know if you have any problems ...
TIA & cheerios, shAf {\/} /\ {\/} /\ {\/} /\ {\/} cogito, ergo zZOooOM {\/} /\ {\/} /\ {\/} /\ {\/} Michael Shaffer, R.A. - University of Oregon Electron Probe Facility mshaf-at-oregon.uoregon.edu -or- mshaf-at-darkwing.uoregon.edu http://darkwing.uoregon.edu/~mshaf/epmahome/
Long storage in glut is not good - the membranes are still permeable since glut does not fix lipid. There will be an exodous of material and changes. But - prefix, wash, postfix with osmium stabilizing the membranes and the lipids, and store in buffer (not in alcohol - alcohol removes osmium) for as long as needed. There is some movement of osmium, but unless the storage is in a solvent, it is neglible. Bye, Hildy
I need some help identifying clays using SEM. Do you know of references, textbooks, etc. with pictures that would me identify common clays i.e. illite, kaolinite, smectite, chlorite, etc.
The knifemaker available to me is a LKB Type 7801A from LKB Instruments Inc. out of Rockville Maryland, but the company has since been taken over by Leica/Leo. The knifemaker has two wheel type gauges that adjust the tension from opposing sides on the glass piece. I can make glass squares, but when I try to cut the square into the two knives, I don't get the correct shapes, even though I have tried systematically changing the settings on the two gauges many different ways. Typically, the "sharp" edge may be chipped, the reflection line in the glass is going in the opposite direction to what it should and the opposite edge to the sharp edge may be either sharp as well or too thick a blunt edge. That's just one of the two knives; its' counterpart often has two "sharp" edges. Can anyone with any direct experience with this model of knifemaker provide me with any advice on how to adjust the settings to produce two glass knives?
Thanks in advance for any assistance that can be provided.
The knifemaker available to me is a LKB Type 7801A from LKB Instruments Inc. out of Rockville Maryland, but the company has since been taken over by Leica/Leo. The knifemaker has two wheel type gauges that adjust the tension from opposing sides on the glass piece. I can make glass squares, but when I try to cut the square into the two knives, I don't get the correct shapes, even though I have tried systematically changing the settings on the two gauges many different ways. Typically, the "sharp" edge may be chipped, the reflection line in the glass is going in the opposite direction to what it should and the opposite edge to the sharp edge may be either sharp as well or too thick a blunt edge. That's just one of the two knives; its' counterpart often has two "sharp" edges. Can anyone with any direct experience with this model of knifemaker provide me with any advice on how to adjust the settings to produce two glass knives?
Thanks in advance for any assistance that can be provided.
People, Does anyone know where to get uranyl formate? My previous suppliers no longer make it-why? I do have the acetate, but the formate works better with microfilaments. Any leads are welcome.
Last winter when I was making some changes to our tissue processing/embedding protocol (which has been around here forever it seems), I began asking questions about EDTA decalcification. One of our professors told me that the EDTA decalcification was found to produce the fewest if any artefacts IF (1) you use the disodium salt (not the tetrasodium), (2) it is done at 4 C (not room temp), AND (3) decalcification is complete in 7-8 days. He claims that tissue pieces which take longer than that begin to loose their membrane integrity. I have no references for this but he has always been a reliable source of information in the past.
Pat Hales McGill University Dept. of Anatomy & Cell Biology hales-at-hippo.medcor.mcgill.ca
If some of you EDS practitioners (yes, I'm a WDS practitioner) could help me with this I would be most appreciative:
1. presently I am involved in a project in which I have to examine zinc arsenides - the process by which these are made can, at times, also produce elemental Zn, ZnO and elemental As (yuk!, and then some)
2. I mainly characterize these beasties by x-ray diffraction, but in this particular inspection I am doing some SEM work too
3. the samples I am looking at are suspected to be mainly the zinc arsenide phase(s), however, I get EDS spectra with highly variable Zn/As - in fact, more wide-ranging than expected for various of the possible zinc arsenides
4. I suspect that the Zn/As variation has as much to do with loss (diminishment) of the As signal (due to absorption?) as it does with variation in the ZnAs
5. to that end (#4), I performed a test yesterday in which I examined just one crystal type (based on morphology) in areas of the sample where these crystals were at the "surface" of the sample and at various "depths" within holes and/or depressions.....basically, the deeper in the "hole" the lower the As signal
6. concurrent with the dimished As, I also observed diminishment of Zn L-alpha line (also a low-energy x-ray) such that Zn-K/Zn-L was also highly variable - would you consider the latter a further indication of absorption?
In the interim, as I await your response, I intend to run x-ray diffraction on those samples with "apparently" low As (some are even As "absent") to "confirm" whether or not the Zn or ZnO phases could be present. Together (my XRD and preliminary SEM, plus your sage advice) I hope we can resolve this.
Thanks, in advance!!
Winton
P.S. if this message "surfaces" twice, please forgive me...the first time I sent it I directed it to the listserver.....once discovered, some kind soul might take it on himself/herself to forward it back to this list
Dr. Winton Cornell Senior Research Associate & Supervisor, Microanalysis Laboratory Department of Geosciences The University of Tulsa 600 South College Tulsa, OK 74104-3189
Hi All: I recently put a message on the listserver requesting information on a LogEtronics enlarger. Some responses I received suggested that I should think about using a digital camera instead to capture images from negatives and then process the images and then send them to a printer. I would like some feedback from users. Is the quality close to that of film? Does anyone have a system that they are extremely happy with? Are there any commercial systems available? Please let me hear your experiences.
Thanks,
Michael Coviello EM Lab Manager The University of Texas -at- Arlington Arlington, TX E-mail coviello-at-mae.uta.edu 817-272-5496
Since my last post I have been working with the XLS spreadsheet file and have added absorption edges, font highlighting, and Chuck's and John's references. The new file name is xray_MS.XLS and is zipped as xray_MS.ZIP ... Like I said before this will be a temporary FTP location for this file ... if Nestor is watching this thread, maybe he'll put the zipped file on the MAS FTP site. For those interested, I'm in the process of adding Cameca spectrometer sine-theta values and creating a PDF file for the purpose of having this data immediately available for on-line browsing ...
You can point your browser at ftp://whitewater.uoregon.edu/share/cameca/
or FTP anonymous to whitewater.uoregon.edu
and look into the "/share/cameca/" directory
Let me know if you have any problems ... and PLEASE let me know if you find any errors ...
TIA & cheerios, shAf {\/} /\ {\/} /\ {\/} /\ {\/} cogito, ergo zZOooOM {\/} /\ {\/} /\ {\/} /\ {\/} Michael Shaffer, R.A. - University of Oregon Electron Probe Facility mshaf-at-oregon.uoregon.edu -or- mshaf-at-darkwing.uoregon.edu http://darkwing.uoregon.edu/~mshaf/epmahome/
Sharon, I would replace the cutting wheel first and then see what happens. The systematic approach should work.
cheers
Edward J. Basgall, PhD The Pennsylvania State University Surface Chemistry Group ejb11-at-psu.edu Materials Research Institute Building Ph: 814-865-0493 University Park, PA 16802-7003 FAX: 814-863-0618
{The knifemaker available to me is a LKB Type 7801A from LKB Instruments Inc. {out of Rockville Maryland, but the company has since been taken over by {Leica/Leo. The knifemaker has two wheel type gauges that adjust the tension {from opposing sides on the glass piece. I can make glass squares, but when {I try to cut the square into the two knives, I don't get the correct {shapes, even {though I have tried systematically changing the settings on the two gauges {many different ways. Typically, the "sharp" edge may be chipped, the {reflection line in the glass is going in the opposite direction to what it {should {and the opposite edge to the sharp edge may be either sharp as well or too {thick a blunt edge. That's just one of the two knives; its' counterpart {often has {two "sharp" edges. Can anyone with any direct experience with this model of {knifemaker provide me with any advice on how to adjust the settings to produce {two glass knives?
{Thanks in advance for any assistance that can be provided.
You will get lots of opinions on this. Here's mine. After spending countless hours in the darkroom over the years, I've converted enthusiastically to digital imaging, and now would prepare any serious final labeled images with Photoshop, making the final prints on a photographic quality printer. Does that mean that all original EM pictures must be taken digitally? I don't think so. If you think about it, only a small percentage of the EM images you take are apt to end up in publications, projection slides or other serious uses. I find it much easier to store numerous EM negatives as 3-1/4x4" sheets of film in glassine envelopes than to fiddle with the multiple zip disks that are necessary to store all the EM digital images in files large enough to allow the resolution that may be necessary later on (the resolution level that we take for granted in film negatives). So I would take them initially on film (even though our department has a Philips EM100 fitted with a CCD camera for digital imaging), and would observe the negatives on a viewer (or on study prints) to decide what will be used for the final product (publication figures, projection slides, etc.). For the final pictures, I would generate digital images by scanning the chosen negatives (with Leafscan 45 scanner) or scanning carefully-prepared prints (with a Hewlett-Packard ScanJet II CX scanner), then crop, arrange, label and otherwise fine tune the pictures with Photoshop 4 on a Power Mac 7600. For photographic quality printing, we use a Kodak XLS 8600 PS printer.
For example, a very complex figure was prepared from multiple darkroom prints, from which numerous small rectangles were cut out and mounted, each showing polysomes at 100 kX mag. To convert to digital image, I scanned the complex figure with the HP ScanJet, enlarging (exaggerating) the image size so the file was 10-15 MB (to provide good resolution). The scan was done without scanner sharpening (sharpening would be done in Photoshop). The file was saved as a TIFF file, and taken up in Photoshop 4. Crop properly (rotate slightly, if necessary). The brightness/contrast of the rectangles varied somewhat, so each was selected (marquee), and Image} Adjust} AutoLevel was used to normalize (be sure white=~5-7% and black=95% in Image} Adjust} Levels [or Curves] eyedroppers). Sharpen whole figure with Filter} Sharpen} Unsharp Mask with amount ~150-200%, radius 1, threshold 0. Dust or other small blemishes (that are clearly of no scientific interest) can be removed with Filter} Noise} Dust & Scratches (after minimal selection with marquee), using radius 1-5 (no more than necessary to remove). Size (Image} Image Size) the figure to width (or height) required by the journal, and set resolution to 400-600 ppi. Put in figure number, size bar, and labels, using layers (not directly on EM image). Print on Kodak XLS printer. The image quality and detail compares well with work done in the darkroom. More and more journals now allow you to submit such digital image files for the final reproductions after a paper has been accepted.
Kent (A. Kent Christensen, University of Michigan, akc-at-umich.edu)
-------------------------------------------------
On Tue, 22 Jul 1997, Mike Coviello wrote:
} I recently put a message on the listserver requesting information on a } LogEtronics enlarger. Some responses I received suggested that I should } think about using a digital camera instead to capture images from negatives } and then process the images and then send them to a printer. I would like } some feedback from users. Is the quality close to that of film? Does anyone } have a system that they are extremely happy with? Are there any commercial } systems available? Please let me hear your experiences. } } Michael Coviello } EM Lab Manager } The University of Texas -at- Arlington } Arlington, TX } E-mail coviello-at-mae.uta.edu } 817-272-5496
The Microstructure and Microanalysis Program in the Characterization and Environmental Laboratory at GE Corporate Research and Development, Schenectady, NY, has an opening for a Polymer Microscopist at the Lead Professional level. The primary duties associated with this position involve the execution of research projects using Transmission Electron Microscopy (TEM) to determine the structure of polymer blends and coatings. Additional duties may involve research conducted using TEM to study other materials, including metals, ceramics, or composites, or using other microscopy techniques, including Acoustic Microscopy and Atomic Force Microscopy.
The Characterization and Environmental Technology Laboratory is involved in research into the structure and composition of materials in support of development programs both at GE CRD and at GE businesses. Staff members are expected to work independently with a high level of expertise, and to become involved with a number of major project teams. Good communication skills, both written and oral, are extremely important.
The minimum requirements for this position are a MS in Materials Science or a closely-related field and some prior experience with Transmission Electron Microscopy. Demonstrated experience in polymer materials science and characterization is also highly desirable.
Resumes and other information can be sent to:
Ernest L. Hall Manager, Microstructure and Microanalysis Program Room K1-2C12 GE Corporate Research and Development PO Box 8 Schenectady, NY 12301 Fax: 518-387-6972 E-mail: hallel-at-crd.ge.com
I'm going to a new position (PPG Industries) and I'm planning to implement a digital darkroom instead of a conventional darkroom. The advantages are tremendous.
At the Materials Directorate at Wright Lab, WPAFB, we have been using a Leaf 45 negative scanner. They are hard to find, but there is a company that has updated it and is now selling it:
I sent out a message several weeks ago on a web site that has information on a collection of negative scanners, flatbed scanners, and some drum scanners:
http://www.foto.unibas.ch/scanners.html
Scitex used to make the Leaf, but now they have their flatbed smartscanners:
http://www.scitex.com/
Here's a site for information on doing it digitally:
A site for drum scanners: http://www.budde.com/products.htm http://www.budde.com/print/magic.htm http://www.budde.com/print/scanview.htm http://www.budde.com/print/sm11000.htm
Our Leaf system is hooked into a MAC system with a lot of RAM, disk space, MO drive, ZIP drive, and access to our LAN. This gives a lot of flexibilty with different users. We bring in the images with a 16 bit format into Photoshop, adjust the levels, convert the image to 8 bit grayscale with appropriate gamma processing, invert the image into a positive print and save. If the intermediate 4 x 5 setting of the Leaf 45 is used, a TEM negative can be digitized with about 1200 dpi. If the image is printed to a 300 dpi grayscale printer (i.e. sub-dye), then that gives an enlargement of 4x at the printer. The Leaf system is capable of much higher pixel resolution and can be used to digitize a very small area on the negative which can be printed with the printer's 300 dpi setting providing a very high enlargement factor. Photoshop has all the tools that could be done in the darkroom but are rather tedious to do such as unsharp masking. Fixing scratches on negatives (mine never have them), dodging, burning an other things that are required for printing a good TEM negative can all be done in a few minutes. In addition, it can be used on both MAC and PC platforms. John Russ's Image Processing Toolkit provides plugins for Photoshop that can provide image processing and stereology: http://members.aol.com/imagproctk/index.htm
I put the scale markers into the print in layers, thus preserving the original scanned image and saving in a TIF format provides the image in a compatible format for other programs. I particularly like printing from Powerpoint onto our Kodak 8650 printer. With Powerpoint, I copy several images of the Tif file on one page (or several differnt files) and print. Output quality if very good.
You need a fast computer with large RAM and large hard drives, a good size monitor (minimum 17"), the scanner, pub-quality printer, and a portable-disk large format disk drive. A 600 dpi laser printer is quite useful also. I also use a flatbed scanner to digitize 6 TEM negatives in a Neg-a-file sheet and digitize the page at 150 dpi. I also digitize my datasheet notes in lineart format. These images are then put into my ThumbsPlus Image database (shareware Program ~$50 for PC and Macs) I then have instant access to my images. It sure beats making proof sheets and processing them.
All of this would probably be cheaper than going with a LogEtronics.
-Hopes this helps.
-Scott
} ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com -----------------------------------------------------------------------. } } Hi All: } I recently put a message on the listserver requesting information on a LogEtronics enlarger. Some responses I received suggested that I should think about using a digital camera instead to capture images from negatives and then process the images and then send them to a printer. I would like some feedback from users. Is the quality close to that of film? Does anyone have a system that they are extremely happy with? Are there any commercial systems available? Please let me hear your experiences. } } Thanks, } } Michael Coviello EM Lab Manager The University of Texas -at- Arlington Arlington, TX E-mail coviello-at-mae.uta.edu 817-272-5496 }
I would like to summarize some of the answers I received concerning the labelling of actin at EM level in non-muscle cells, namely chondrocytes in the rabbit growth plate.
Rosemary White labelled actin in plants using monoclonal C4 anti-actin from ICN on LR White sections (Protoplasma 131:153-165 and 150:72-74). This antibody is mouse IgG against human actin and recognizes a common actin epitope in many species. Kirk Czymmek was successful using the antibody N.350 from Amersham to label actin at the EM level in fungi (J Microscopy Vol 181, Feb 1996 pp 153-161; Protoplasma 163, pp. 199-202). This is a mouse IgM against chicken gizzard actin and has been shown to label human, monkey, chicken, rat, etc actin, therefore binding to a highly conserved region of actin. BTW, we ordered this antibody today and hope to have some results shortly!
Since we got nice labelling with phalloidin-BODIPY FL in the confocal, we wanted to use rabbit anti-BODIPY FL antibody (Molecular Probes) followed by anti-rabbit-gold to reveal the actin at EM level. This was not successful. Molecular Probes could not give us any references of anybody using this antibody for immunohistochemistry. Tamara Howard commented, that though she has not used the anti-BODIPY, she had a dismal luck with antibodies to FITC and Texas Red from various sources at the EM level.
Thanks to everybody who took the time and responded to my questions. This list is such a wonderful way of sharing experience! I would never think of looking in Protoplasma, since Medline does not bring it up!
Sarka Lhotak EM Facility, McMaster University Hamilton, Ontario, Canada
Dear Winton, If you look at particles in holes or depressions, the edges of the hole will preferentially absorb the lower energy x-rays while the higher energy ones can penetrate the edges of the hole. If you look at the As Ka line with the SEM at 25 or 30 kV it will be less affected than the As L line. You will also get better quantification. As with WDS, a flat, smooth sample would also help. You wrote: } Micro-colleagues: } } If some of you EDS practitioners (yes, I'm a WDS practitioner) could help me } with this I would be most appreciative: } } 1. presently I am involved in a project in which I have to examine zinc } arsenides - the process by which these are made can, at times, also produce } elemental Zn, ZnO and elemental As (yuk!, and then some) } } 2. I mainly characterize these beasties by x-ray diffraction, but in this } particular inspection I am doing some SEM work too } } 3. the samples I am looking at are suspected to be mainly the zinc arsenide } phase(s), however, I get EDS spectra with highly variable Zn/As - in fact, } more wide-ranging than expected for various of the possible zinc arsenides } } 4. I suspect that the Zn/As variation has as much to do with loss } (diminishment) of the As signal (due to absorption?) as it does with } variation in the ZnAs } } 5. to that end (#4), I performed a test yesterday in which I examined just } one crystal type (based on morphology) in areas of the sample where these } crystals were at the "surface" of the sample and at various "depths" within } holes and/or depressions.....basically, the deeper in the "hole" the lower } the As signal } } 6. concurrent with the dimished As, I also observed diminishment of Zn } L-alpha line (also a low-energy x-ray) such that Zn-K/Zn-L was also highly } variable - would you consider the latter a further indication of } absorption? } } In the interim, as I await your response, I intend to run x-ray diffraction } on those samples with "apparently" low As (some are even As "absent") to } "confirm" whether or not the Zn or ZnO phases could be present. Together } (my XRD and preliminary SEM, plus your sage advice) I hope we can resolve this. } } Thanks, in advance!! } } Winton } } } P.S. if this message "surfaces" twice, please forgive me...the first time I } sent it I directed it to the listserver.....once discovered, some kind soul } might take it on himself/herself to forward it back to this list } } } Dr. Winton Cornell } Senior Research Associate & Supervisor, Microanalysis Laboratory } Department of Geosciences } The University of Tulsa } 600 South College } Tulsa, OK 74104-3189 } } phone: 918-631-3248 } fax: 918-631-2091 } e-mail: wcornell-at-centum.utulsa.edu } } } } Mary Mager Electron Microscopist Metals and Materials Eng., UBC 6350 Stores Rd. Vancouver, B.C. V6T 1Z4 CANADA tel:604-822-5648, fax:604-822-3619 e-mail: mager-at-unixg.ubc.ca
Continuing my reports on Inter/Micro 97: Today's sessions focused on Instrumentation (AM) and Techniques (PM).
In the morning session, James M. Landrigan III of Polaroid discussed their new Digital Microscope Camera ("DMC"). I have been anxious to see what Polaroid has been up to and I am not surprised to report that their new system will mark a significant addition to the development of digital imaging. I won't risk mis-quoting the specs here since I didn't yet get their literature, but I'll just mention that the system includes a dedicated digital camera with a standard C-mount lens attachment which should make it readily attachable to a range of microscopes (and other equipment). The system also incorporates a larger (12.5mm?) pixel array than competing systems which they claim results in an improved signal-to-noise ratio. The camera includes more than 1,000,000 pixels and produces images of either 800X600 or 1600X1200 pixels (other formats too?) Part of the available resolution is from software interpolation, which I'm not wild about, but I couldn't get clear just what the actual pixel array was so you'll want to get more details from them. Also, one factor which concerns me at first hearing is that they use a rectangular pixel, not a square one. I'll want to ask them why that choice was made. The price is about $6,000 for the camera and software system which, while not cheap, certainly seems reasonable. All-in-all, I'm happy to see Polaroid step into this field. Competition among the giants can only help us users of the technology and this appears to be a good entry into the fray.
Wayne Niemeyer of the McCrone Group showed some applications and results of "Low Voltage Scanning Electron Microscopy." Those of you who know the capabilities of these systems won't need "preaching to the choir" but I must say that it really is impressive what can be done with these systems. Wayne showed some beautiful images of exquisitely fine structural features, some in complex, deeply three-dimensional surfaces, all without any trace of charging, all in sharp, clear detail. I've seen 'em before but I was impressed again.
John Reffner of Nicolet/Spectra-Tech reminded us that "There is Microscopy in Infrared Microspectroscopy." He pointed out the complimentary nature of microscopy and microspectroscopy, that a scientist should be not a microscopist _or_ a spectroscopist, but both if he or she wants to fully exploit the capabilities of these instruments. It is appropriate that one from the company which actually emphasizes the importance of good microscopy (and good microscopes) as a part of microspectroscopy should make this point. Spectra-Tech is, in my opinion, to microspectroscopy what Zeiss used to be to microscopy. Yes, they may be the most expensive, but "you get what you pay for" here as elsewhere. I think that we should advocate retaining the capability to do good work with our equipment, even if we need to learn a bit more to take advantage of that capability, rather than accept instruments which have been "dumbed-down" to the least common denominator of the people who use them. So far, Spectra-Tech has not succumbed to any pressure they might feel to adopt that trend and I hope you'll join me in encouraging them to continue to hold the high ground. Someone needs to!
(Ok, ok, so I jamb in a bit of editorializing too. I'm not related to any of the companies I'm mentioning, so I think I can use a bit of license here.)
In the afternoon, Theodore M. Clarke of J.I. Case Corporation discussed "High Magnification Photomacrography Using the Kodak 1.6i/AB MegaPlus (TM) Camera." This provided a nice complement to the Polaroid talk, but this by an independant technical evaluator who put the Kodak product through some serious resolution tests, finding that it performed well if you accepted Ted's recommended 500 X NA rule of thumb for maximum useful magnification. This is 1/2 of the conventional wisdom but Ted made (and illustrated) a good argument for the more conservative standard. The Kodak product performed quite well in Ted's tests and I suggest all interested parties watch for the publication of his work in The Microscope in coming months. The technical detail of his work was too much to repeat here, but watch for the publication as it will be well worth using not only for its evaluation of the Kodak product but as a model for how digital cameras can be well and truely tested.
Also in the afternoon, Allen Whiteside of the McCrone Group presented two back-to-back papers on "Preparing Holey Carbon Films" and "AEM Preparations Using Holey Carbon Film." (Note that "The McCrone Group" is a different organization than the meeting hosts, the McCrone Research Institute. They separated years ago.) Once again, the folks at the McCrone Group demonstrate that they are masters of specimen manipulation (i.e., particle handling) and specimen preparation as well as analytical microscopy. I think they have probably originated more useful techniques than any other single organization. Allen showed their in-house technique for the preparation of holey carbon films which can and should be tailored to the specific needs of the analysis at hand. A film of one thickness, pore density, and web structure will be appropriate for one analysis, something different for another. They drop a solution of formvar in ethylene dichloride onto a glass slide which is rotated on a turntable to spin out the applied fluid while the solvent evaporates. The film is then lifted and applied to grids in a more-or-less conventional manner but the secrets of the technique lie in the speed of rotation and the formulation of the solvent/formvar solution. Again, the idea is to experiment and determine the best film preparation system for a particular kind of analysis, not adopt a single procedure. There is more, or course, and this was the subject of Allen's second talk. There are numerous techniques for applying the particles of interest to the holey film, again chosen to best suit the needs of the analysis.
I should close, but I just want to mention one more thing. There are many more papers being presented than I'm mentioning here. I'm only hitting on a few that seem particulary interesting to me for one odd reason or another. One should not infer anything from my failing to mention some of the other fine papers!
More to come, stay tuned!
Steve Shaffer MicroDataware
p.s. Yes, we are till publishing the Particle Atlas Electronic Edition (PAEE). Please contact me via EMail if you would like further information.
p.p.s. I know I'm a terrible speller and I'm preparing these things quickly while on the road and without access to my regular array of make-me-look-good software, so please forgive the rough nature of these communications. ;-)
I've dealt with both 35mm (immunofluorescence) and 3 1/4 X 4 1/4 inch (EM) negatives in both ways. Using a high quality light box and a very high quality macro lens on a fairly standard sort of CCD camera (Dage 72, 640X480, 8bits). I've gotten good digital images, printed on a Codonics dye-sub. But (my humble opinion) the quality doesn't approach what you can do in the darkroom. It is, however, WAY faster and easier. A fancier camera may help, but it all depends on what you want to do with the images. If the images are already on film, then I would tend to stick with silver grains over pixels. If you really need to go digital and have some money, scanners are probably the way to go for digitizing negatives -- check the list's archives, there has been much discussion of scanners.
Greg Martin Dept. of Cell Biology and Anatomy Johns Hopkins School of Medicine
On Tue, 22 Jul 1997, Mike Coviello wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Hi All: } I recently put a message on the listserver requesting information on a } LogEtronics enlarger. Some responses I received suggested that I should } think about using a digital camera instead to capture images from negatives } and then process the images and then send them to a printer. I would like } some feedback from users. Is the quality close to that of film? Does anyone } have a system that they are extremely happy with? Are there any commercial } systems available? Please let me hear your experiences. } } Thanks, } } Michael Coviello } EM Lab Manager } The University of Texas -at- Arlington } Arlington, TX } E-mail coviello-at-mae.uta.edu } 817-272-5496 } } }
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Mike,
I would buy a Durst 1200 point source enlarger if you are doing high resolution electron microscopy ---- the logEtronics enlarger uses a scanning spot to equalize large contrast variations within an image and this enlarger prints like a diffuse source enlarger not a point source enlarger. We have found that lines are sharper and narrower with the point source enlarger. Our work requires darkroom printing of reversal negatives made from thin vertically Pt-C replicated specimens in which we want to visualize molecular details on the structural highlights. Usually these features only become visible after printing the reversal negative on fiber based paper and drying the print on glossy paper---- For TEM publications that do not need this detail as part of their story, it is easy to scan in images at a hardware resolution of 600 dpi and label the image with Adobe photoshop---- I find that I need access to a good darkroom as well as the appropriate digital darkroom equipment.
George Ruben Dept. Biological Sciences Dartmouth College Hanover, NH 03755
Hi everyone, for many years I have been embedding in Spurr's resin but have recently decided to give glycol methacrylate a whirl for thicker LM sections of plant material. Many years have passed since my last work with GMA. I remember that we made a plexigalss container which we could evacuate the oxygen from by flushing with nitrogen and that polymerization was then carried out below a longwave UV light placed in the container. Does this sound correct? If so does anyone have any plans or tricks that might assist me in constructing such a device? Cheers, John
================= C. John Runions, Ph.D Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
We've gone digital TEM and everyone loves it for ease of capture and speediness to the investigator. Our 2Kx2K images look great on a large monitor and printed with on a Tektronix dye-sub printer. Heck, even an Epson Stylus printer with 1440 dpi makes darn good prints (on photo-quality paper)! In a side-by-side comparison, I'll take a high contrast print from our Durst EM1200 over the dye-sub. However, dollar for dollar, one alternative to consider is to transfer TEM negatives to Kodak PhotoCD and print only select images, when you need publication quality, to the Fujix Pictrograph 3000 digital printer (which costs less than the Durst with all the electronic bells and whistles.) Thereby you have your negative, a high-resolution digital image, a digital print, and you can still make a print if you really have to.
There's my humble opinion. Thank you.
Walter F. Bobrowski Subcellular Pathology Parke-Davis Pharmaceutical Research Ann Arbor, MI 48105
To all who would advocate a totally digital darkroom,
While on the whole I have to agree that the direction that science/microscopy is taking is a digital one, I must point out that there are some serious shortcomings to digital imaging. The most serious, in my mind, is the issue of archivability. While I could launch into this myself, I'd prefer to bring to everyone's attention the following article (which addresses this issue far better than I can):
"Ensuring the Longevity of Digital Documents" Jeff Rothenberg, Scientific American, January 1995
If we can't honestly answer the question of how we, or anyone else, will be able to use (or even access) our digital images 30 years from now, we need to re-evaluate the speed with which we are going digital. If properly cared for, film can last for decades. The uncertain aging of digital storage media, the often rapid obsolescence of drive mechanisms & media types and the ongoing changes in image file formats are cause for concern.
Yours, Doug Cromey ..................................................................... : Douglas W. Cromey, M.S. Dept. of Cell Biology & Anatomy : : Sr. Research Specialist University of Arizona : : (office: AHSC 4212A) P.O. Box 245044 : : (voice: 520-626-2824) Tucson, AZ 85724-5044 USA : : (FAX: 520-626-2097) (email: doug-cromey-at-ns.arizona.edu) : :...................................................................: http://www.pharmacy.arizona.edu/exp_path.html
Our multi-user facility is currently archiving our confocal and LM digital images on Panasonic optical disks (re-writable, very stable -at- about $125 for 1 GB). The disadvantage is that few of our users have their own Panasonic drives so most people simply archive the images at our core and then move the ones they want by FTP as needed. I would like to switch to a more universal medium - namely CD ROM's. My understanding is that CD's can now be written to in multiple sessions so you don't need to fill an entire disk at once. Furthermore, it is my understanding that a disk of TIFF images should be readable by both IBM/WINTEL and Mac/PowerPC types computers. Is anybody actually doing this? Comments on how reliable are the recorders, which ones are best, pitfalls, etc would be appreciated. Before I get a dozen advocates of ZIP/Jazz drives, I don't want to go that route since that they are not as ubiquitous as CD drives. Thanks in advance.
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility 3 Tucker Hall University of Missouri Columbia, MO 65211 (573)-882-4712 (voice) (573)-882-0123 (fax)
Mike, your needs for image quality may exceed ours (industrial research laboratory, not as interested in publishing outside as academic), but we recently removed all of our darkrooms because we had not processed a negative or print in two years.
All of our microscopy (AFM, TEM, SEM, and Optical) is done with digital image capture. The speed to get a hardcopy, the flexibility of sharing images worldwide via electronic transfer, and the cost per print/image all strongly favor total digital imaging. Although the quality of a typical digital image does not equal that which can be obtained with photographic film they meet our needs completely. For the highest quality digital imaging in optical microscopy we use a Kodak 460 digital camera or a Leaf MicroLumina (3500 X 2400 pixels). Storage of images is done on CD-ROM for archival capability and erasable Magneto-Optical CD (40GB jukebox on a network server) is used for day-to-day image storage. Photoshop is used for image "retouching". Reports are composed in MicroSoft Word and hardcopy of images are obtained with networked Kodak 8650 printers (one dedicated to B/W and one for color).
As a final note, we've been using video and digital images for 9 years and would never think of going back to a photographic imaging process.
********************************************************* Dr. Dennis B. Barr Research Laboratories Eastman Chemical Company Kingsport, TN 37664-5150 Phone:423-229-2188 Fax:423-229-4558 Email: dennbarr-at-eastman.com ********************************************************
} -----Original Message----- } From: Mike Coviello [SMTP:Coviello-at-mae.uta.edu] } Sent: Tuesday, July 22, 1997 4:24 PM } To: Microscopy-at-Sparc5.Microscopy.Com } Subject: Digital Darkrooms } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
If you spend $30,000--$50,000 for hardware and software (which will be obsolete in less than five years) you can produce very good hardcopy images comparable to conventional photography. The digital process requires a highly skilled system caretaker and the learning period for neophytes is much longer. For every year that you can delay the transition to digital you will reduce the problems significantly. These statements are based on my experiences with photography for 32 years and digital graphics for 10 years. Larry D. Ackerman (415) 476-8751 Howard Hughes Medical Institute FAX (415) 476-5774 UCSF, Box 0724, Rm U426 533 Parnassus Ave. mishot-at-itsa.ucsf.edu San Francisco, CA 94143
On the other hand... I can send digital images to any of my JIT-based manufacturing customers in a matter of moments via e-mail. Do that with film! I can store images with and without annotation in an inexpensive database that is continually being upgraded by its developers... and is forward and backward compatible with our standard desktop applications. I have reduced my Polaroid film expenditures by at least five thousand dollars per year (with the attendant costs to order, deliver, bill, etc.) My image acquisition system is never out of stock. Images don't get "lost in the mail" I can extract a variety of numbers from a digital image easier than using primitive tools like an ASTM grain size overlay.
It should be noted that the medical industry is a driving force for improvements in digital imaging technologies and electronic collaboration precisely because of the failure of film to fulfill the needs of today's customers.
} ------------------------------------------------ } Opinions or statements expressed herein, rational or otherwise, do not } necessarily reflect those of my employer. } } Harold J. Crossman } OSRAM SYLVANIA INC. } Lighting Research Center } 71 Cherry Hill Dr. } Beverly, MA 01915 } Phone: (508) 750-1717 } E-mail: crossman-at-osi.sylvania.com } } Our web sites: www.sylvania.com } www.siemens.com } -- } } "Crossman, Harold" {crossman-at-osi.SYLVANIA.com}
We use CD-ROMs for archiving images, data, etc. If you make the CD-ROM in ISO-9660 format it can be read by PC's, Macs, and Unix machines. ISO-9660 does not allow long file names. There are other formats - Joliet system - that allow long file names but these can only be read in Win95 machines. I recommend that you have a dedicated machine for making CD-ROMS. Partition the hard drive so that your system files are on C-drive and leave the D-drive for files to be archived. Two of the major manufacturs are Yamaha and Pinnacle Micro. If you look up their web-sites and read the FAQ's related to installation and troubleshooting, you will get some idea of the important issues in setting up a system (There are certain hard drive specifications, etc.) Multisession is possible, but you need software that will read a multisession disk (usually the software package that was used to make the CD). If you make a multisession disk, and place it in a computer without the proper software - the computer will only see the last session. This drawback may be changing (changed?) with the next generation of machines and software. Overall, I think CD-ROM is the current best method for archiving data.
Regards,
John J. Turek, Ph.D. Associate Professor Director, Electron Microscopy Laboratory and Core Laboratory for Image Analysis and Multidimensional Applications (CRISTAL) Department of Basic Medical Sciences 1246 Lynn Hall, G193C Purdue University W. Lafayette, IN 47907-1246 Phone: 765-494-5854 Fax: 765-494-0781 Email: jjt-at-vet.purdue.edu
We have installed writable CD-ROM on a UNIX system using software manufactured by Microson, called GEAR 32.
There have been serious problems arising from incompatibility of UNIX with CD-ROM technology (asynchronous versus synchronous - The Unix machines deliver information when they are ready but the CD writing process requires information at a constant rate which is determined by the disc speed). I don't believe this is a problem with PC's, but watch out if you want to use CD-R on a UNIX platform. We had to buy a new external hard drive, directly connected to the CD-R device, in order to get everything to work reliably.
Also, and this may apply to you as well, the claims of the software manual concerning various options like multisession backing up were not actually implementable. We must write an entire CD at once. This is is not so bad actually as we deal with large quantities of image data, and the disks themselves are now only around $4-$5, which you really can't beat for 650MB of space. The software packages available when working in the PC world may be better, but beware, and make sure you get what is advertised. I think this technology has a future and that the microscopy world can benifit. However, the software for running it, at least on a UNIX platform, has some progress still ahead of it.
Wharton
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ Wharton Sinkler PhD Department of Materials Science and Engineering Northwestern University 2225 North Campus Drive Evanston, IL 60208-3108 tel: (847) 491-7809 fax: (847) 491-7820 email: sinkler-at-apollo.numis.nwu.edu
On Wed, 23 Jul 1997, Tom Phillips wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Our multi-user facility is currently archiving our confocal and LM digital } images on Panasonic optical disks (re-writable, very stable -at- about $125 } for 1 GB). The disadvantage is that few of our users have their own } Panasonic drives so most people simply archive the images at our core and } then move the ones they want by FTP as needed. I would like to switch to a } more universal medium - namely CD ROM's. My understanding is that CD's can } now be written to in multiple sessions so you don't need to fill an entire } disk at once. Furthermore, it is my understanding that a disk of TIFF } images should be readable by both IBM/WINTEL and Mac/PowerPC types } computers. Is anybody actually doing this? Comments on how reliable are } the recorders, which ones are best, pitfalls, etc would be appreciated. } Before I get a dozen advocates of ZIP/Jazz drives, I don't want to go that } route since that they are not as ubiquitous as CD drives. Thanks in } advance. } } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } 3 Tucker Hall } University of Missouri } Columbia, MO 65211 } (573)-882-4712 (voice) } (573)-882-0123 (fax) } } }
We have installed writable CD-ROM on a UNIX system using software manufactured by Microson, called GEAR 32.
There have been serious problems arising from incompatibility of UNIX with CD-ROM technology (asynchronous versus synchronous - The Unix machines deliver information when they are ready but the CD writing process requires information at a constant rate which is determined by the disc speed). I don't believe this is a problem with PC's, but watch out if you want to use CD-R on a UNIX platform. We had to buy a new external hard drive, directly connected to the CD-R device, in order to get everything to work reliably.
Also, and this may apply to you as well, the claims of the software manual concerning various options like multisession backing up were not actually implementable. We must write an entire CD at once. This is is not so bad actually as we deal with large quantities of image data, and the disks themselves are now only around $4-$5, which you really can't beat for 650MB of space. The software packages available when working in the PC world may be better, but beware, and make sure you get what is advertised. I think this technology has a future and that the microscopy world can benifit. However, the software for running it, at least on a UNIX platform, has some progress still ahead of it.
Wharton
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ Wharton Sinkler PhD Department of Materials Science and Engineering Northwestern University 2225 North Campus Drive Evanston, IL 60208-3108 tel: (847) 491-7809 fax: (847) 491-7820 email: sinkler-at-apollo.numis.nwu.edu
On Wed, 23 Jul 1997, Tom Phillips wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Our multi-user facility is currently archiving our confocal and LM digital } images on Panasonic optical disks (re-writable, very stable -at- about $125 } for 1 GB). The disadvantage is that few of our users have their own } Panasonic drives so most people simply archive the images at our core and } then move the ones they want by FTP as needed. I would like to switch to a } more universal medium - namely CD ROM's. My understanding is that CD's can } now be written to in multiple sessions so you don't need to fill an entire } disk at once. Furthermore, it is my understanding that a disk of TIFF } images should be readable by both IBM/WINTEL and Mac/PowerPC types } computers. Is anybody actually doing this? Comments on how reliable are } the recorders, which ones are best, pitfalls, etc would be appreciated. } Before I get a dozen advocates of ZIP/Jazz drives, I don't want to go that } route since that they are not as ubiquitous as CD drives. Thanks in } advance. } } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } 3 Tucker Hall } University of Missouri } Columbia, MO 65211 } (573)-882-4712 (voice) } (573)-882-0123 (fax) } } }
} ... } } . . . The most serious, in my mind, is the issue of archivability. } ... } } "Ensuring the Longevity of Digital Documents" } Jeff Rothenberg, Scientific American, January 1995 } } If we can't honestly answer the question of how we, or anyone else, } will be } able to use (or even access) our digital images 30 years from now, we } need } to re-evaluate the speed with which we are going digital. If properly } } cared for, film can last for decades. The uncertain aging of digital } storage media, the often rapid obsolescence of drive mechanisms & } media } types and the ongoing changes in image file formats are cause for } concern. } } ...
I agree ... however, while there exists good reasoning to avoid magnetic media for long-term archiving, there seems to be little concern in this regard for magneto-optical or CD ROM media. Also regarding archival quality ... digital hardcopy will not hold up to long-term storage of preperly washed photographic papers (... I think it has already been mentioned that the resolution and the quality of the grayscale for even the best digital printers doesn't even come close to chemica darkroom printing ...). Still ... even as an ex-photographic art student ... I can appreciate the new capabilities (in general) that a digital darkroom has over the traditional for producing *lots of work* and its being "publication quality" ... if not "art for the purist" quality ...
cheerios, shAf -- {\/} /\ {\/} /\ {\/} /\ {\/} cogito, ergo zZOooOM {\/} /\ {\/} /\ {\/} /\ {\/} Michael Shaffer, R.A. - University of Oregon Electron Probe Facility mshaf-at-oregon.uoregon.edu -or- mshaf-at-darkwing.uoregon.edu http://darkwing.uoregon.edu/~mshaf/
Hi Tom. I have been burning CD's for our SEM users for about a year and a half now. We have an HP 4020i and I am using the Easy CD software package (supposedly the best). Of the 50 or so users who have archived onto CD only 2 have not been able to get images I wrote and another says he can't but I think in his case it is the operator. Both of the users who are having trouble are able to read the data from the first write, but not from successive write sessions. Thay have brought their disks back into the lab and I am able to read ALL the data from ALL write sessions on each of our four cd readers. Our computer guy here asked how old the drives they were using to read the disks were cause if they are too old they may not support the multi-session standards. Only one user seems to have such an aged cd rom. The other is baffeling. Heck, my cd-rom at the house is old as the hills (and the cheapest I could find) and it does fine. HP was no help in this matter either so if anyone has any ideas I would sure appreciate hearing from you. I can still say I reccommend the writer and feel it is probably the best deal. I don't feel bad making the users buy an $8-9 disk, even the guys who can only read the first session. You are correct that both MAC & PC's will read the files as long as the MAC is running system 7 or better. SGI will not nor will OS2. Good luck
At 10:32 AM 7/23/97 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} { GO GATORS Scott D. Whittaker 218 Carr Hall Research Assistant Gainesville, FL 32610 University Of Florida ph 352-392-1295 ICBR EM Core Lab fax 352-846-0251 sdw-at-biotech.ufl.edu http://www.biotech.ufl.edu/~emcl/ The home of " Tips & Tricks "
We have been archiving to CDs with a Philips 2000 for almost two years with no problems. Much cheaper than opticals. $9.00 for 600 MB.
Bob Morphology Core Seattle
On Wed, 23 Jul 1997, Tom Phillips wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Our multi-user facility is currently archiving our confocal and LM digital } images on Panasonic optical disks (re-writable, very stable -at- about $125 } for 1 GB). The disadvantage is that few of our users have their own } Panasonic drives so most people simply archive the images at our core and } then move the ones they want by FTP as needed. I would like to switch to a } more universal medium - namely CD ROM's. My understanding is that CD's can } now be written to in multiple sessions so you don't need to fill an entire } disk at once. Furthermore, it is my understanding that a disk of TIFF } images should be readable by both IBM/WINTEL and Mac/PowerPC types } computers. Is anybody actually doing this? Comments on how reliable are } the recorders, which ones are best, pitfalls, etc would be appreciated. } Before I get a dozen advocates of ZIP/Jazz drives, I don't want to go that } route since that they are not as ubiquitous as CD drives. Thanks in } advance. } } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } 3 Tucker Hall } University of Missouri } Columbia, MO 65211 } (573)-882-4712 (voice) } (573)-882-0123 (fax) } } }
} ... } } } } Our multi-user facility is currently archiving our confocal and LM } digital } } images on Panasonic optical disks (re-writable, very stable -at- about } $125 } } for 1 GB). The disadvantage is that few of our users have their own } } } Panasonic drives so most people simply archive the images at our } core and } } then move the ones they want by FTP as needed. I would like to } switch to a } } more universal medium - namely CD ROM's. My understanding is that } CD's can } } now be written to in multiple sessions so you don't need to fill an } entire } } disk at once. Furthermore, it is my understanding that a disk of } TIFF } } images should be readable by both IBM/WINTEL and Mac/PowerPC types } } computers. Is anybody actually doing this? Comments on how } reliable are } } the recorders, which ones are best, pitfalls, etc would be } appreciated. } } Before I get a dozen advocates of ZIP/Jazz drives, I don't want to } go that } } route since that they are not as ubiquitous as CD drives. Thanks } in } } advance. ...
Whereas I would have thought CDROM should be the best method (... considering the cost of the media ...), judging from the responses we've seen I'm glad I went with a Fujitsu 640 magneto-optical. It has a solid SCSI interface (... I use Adaptec ...) and it is re-writable. I had some early issues when it first came out (... we had been using 230Mb for 1.5 years ...) but it works flawlessly now. The cost per 640Mb cartridge is (however) $35 ... and (... of course ...) the drives aren't as ubiquitous. We use it in a intranet configuration, and all image files and data from the SEM and Cameca archive directly to it. Non-WIntel users download from it via "anonymous" FTP ...
... another point of view ...
cheerios, shAf -- {\/} /\ {\/} /\ {\/} /\ {\/} cogito, ergo zZOooOM {\/} /\ {\/} /\ {\/} /\ {\/} Michael Shaffer, R.A. - University of Oregon Electron Probe Facility mshaf-at-oregon.uoregon.edu -or- mshaf-at-darkwing.uoregon.edu http://darkwing.uoregon.edu/~mshaf/
If some of you EDS practioners (yes, I'm a WDS practioner) could help me with this I would be most appreciative:
1. presently I am involved in a project for which I have to examine zinc arsenides - the process by which these are made can, at times, also produce elemental Zn, ZnO and elemental As (yuk!, and then some)
2. I mainly characterize these beasties by x-ray diffraction, but in this particular inspection I am doing some SEM work too
3. the samples I am looking at are suspected to be mainly the zinc arsenide phase(s), however, I get EDS spectra with highly variable Zn/As - in fact, more wide-ranging than expected for various of the possible zinc arsenides
4. I suspect that the Zn/As variation has as much to do with loss (diminishment) of the As signal (due to absorption?) as it does with variation in the ZnAs
5. to that end (#4), I performed a test yesterday in which I examined just one crystal type (based on morphology) in areas of the sample where these crystals were at the "surface" of the sample and at various "depths" within holes and/or depressions.....basically, the deeper in the "hole" the lower the As signal
6. concurrent with the dimished As, I also observed diminishment of Zn L-alpha line (also a low-energy x-ray) such that Zn-K/Zn-L was also highly variable - would you consider the latter a further indication of absorption?
In the interim as, I await your response, I intend to run x-ray diffraction on the samples with "apparently" low As (some are even As "absent") to "confirm" whether or not the Zn or ZnO phases could be present. Together (my XRD and preliminary SEM, plus your sage advice) I hope to resolve this.
Thanks, in advance!!
Winton
Dr. Winton Cornell Senior Research Associate & Supervisor, Microanalysis Laboratory Department of Geosciences The University of Tulsa 600 South College Tulsa, OK 74104-3189
Sharon et.al. That knife maker is an everlasting instrument but the adjustments must be right, otherwise its a big waste of glass and time. 1 Adjust the back holder of the glass, so that the cutting stroke across the diagonal of the glass square stops one or two mm before running off the glass. 2 Loosen the front holder and press the moving part to just touch the corner of the glass. Increase the push against the spring tension by two scale divisions and then tighten the knurled knob. 3 The front and back lateral adjustments must now be set. Centre both of these, check with a glass square inserted and a piece of paper as a straight edge that the centre line would be corner to corner on the glass. 4 Now adjust the back lateral adjustment so that the centre line would be 1 to 2mm to the left of the back corner. Tighten the lock-nut. 5 Repeat for the front adjustment but to the right of the front corner. 6 Break a couple of test squares, without the front damper, which pushes a bit of rubber against the glass, applied. Those knives should have their edges close, but not across the corners. If required adjust the lateral controls. 7 Once well set, the laterals never need adjustment. Draw a marker pen line across the dials to indicate settings or tighten them just beyond finger strengths.
Never touch the clamping head when it is lowered; it determines the clamping pressure and over-tightening its locking device is pointless.
The rubber damping device slows down the moment of fracture of the front knife only. This result in a wider stress free area for that knife. Apply the damper after clamping the head. Mover rubber front to just touch glass plus two scale divisions. The glass should not move back. Score, break and then, always before lifting the clamping head, release the damper.
I am writing this from memory but I've taught that operation a few hundred times. Ask me if you still have problems with the knife maker. Disclaimer: ProSciTech and most other EM suppliers supply microtomy glass. We should have an interest in maladjusted knife-makers. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 77 740 370 Fax: +61 77 892 313 Great microscopy catalogue, 400+ Links, MSDS ************************ http://www.proscitech.com.au
---------- } From: Sharon C. Thomas {sthomas-at-lanl.gov} } To: Microscopy-at-sparc5.microscopy.com } Subject: Help requested for maladjusted glass knifemaker } Date: Wednesday, 23 July 1997 3:44 } The knifemaker available to me is a LKB Type 7801A from LKB Instruments Inc. } out of Rockville Maryland, but the company has since been taken over by } Leica/Leo. The knifemaker has two wheel type gauges that adjust the tension } from opposing sides on the glass piece. I can make glass squares, but when } I try to cut the square into the two knives, I don't get the correct } shapes, even } though I have tried systematically changing the settings on the two gauges } many different ways. Typically, the "sharp" edge may be chipped, the } reflection line in the glass is going in the opposite direction to what it } should } and the opposite edge to the sharp edge may be either sharp as well or too } thick a blunt edge. That's just one of the two knives; its' counterpart } often has } two "sharp" edges. Can anyone with any direct experience with this model of } knifemaker provide me with any advice on how to adjust the settings to produce } two glass knives? } } Thanks in advance for any assistance that can be provided. } }
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Tom:
While we have do many Zip drives in use in our multi-user facility, they are used for purposes other than archive storage. We archive all of our digital images (from 6 major instruments) onto CD-ROMS. We have more than 60 Gbytes of active, primary storage on on-line servers, which is good enough for 3-4 months of image storage, after which the oldest images are downloaded onto CDs. This has worked nicely for us for the last 3-4 years. We store all our images in their original formats (DigitalMicrograph, Adobe PhotoShop etc.) so they can be handled ultimately just as if they had just been recorded.
Hope this helps.
Larry
} } Our multi-user facility is currently archiving our confocal and LM digital } images on Panasonic optical disks (re-writable, very stable -at- about $125 } for 1 GB). The disadvantage is that few of our users have their own } Panasonic drives so most people simply archive the images at our core and } then move the ones they want by FTP as needed. I would like to switch to a } more universal medium - namely CD ROM's. My understanding is that CD's can } now be written to in multiple sessions so you don't need to fill an entire } disk at once. Furthermore, it is my understanding that a disk of TIFF } images should be readable by both IBM/WINTEL and Mac/PowerPC types } computers. Is anybody actually doing this? Comments on how reliable are } the recorders, which ones are best, pitfalls, etc would be appreciated. } Before I get a dozen advocates of ZIP/Jazz drives, I don't want to go that } route since that they are not as ubiquitous as CD drives. Thanks in } advance. } } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } 3 Tucker Hall } University of Missouri } Columbia, MO 65211 } (573)-882-4712 (voice) } (573)-882-0123 (fax)
Dr. Lawrence F. Allard Senior Research Staff Member High Temperature Materials Laboratory Oak Ridge National Laboratory 1 Bethel Valley Road Bldg. 4515, MS 6064 PO Box 2008 Oak Ridge, TN 37831-6064
I am a biophysics graduate student. One of my exam question is how to correct CTF in image reconstruction. After finishing the exam, I have more questions than answers!
What kinds of contrast exists in electron microscopic image for weak-phase object? Is amplitude contrast the same as aperture contrast?
What is the contribution of inelastic scattering on the image contrast? How to account for it?
How does partial spatial coherence and temporal coherence influence the image and electron diffraction?
Is it always sufficient to use just first order approximation ( the linear theory of phase and amplitude contrast image formation ) for CTF consideration?
Is it true for low spatial frequencies ignoring the CTF was better than compensating for phase contrast alone?
I guess compensation for the CTF was necessary and sufficient to accurately reconstruct molecular densities. Could anyone tell me the current ways for accurate determination of CTF?
By improving the different components in CTF, what is the optimal contrast could be achieved in image?
Is it possible to put EM reconstruction on the same scale with the structures derived from X-ray crystallography and NMR after deliberate correction of CTF, solvent effects and differences between atomic scattering factors for electron and X-ray?
To what extend we could use the insights obtained by building models and comparison of the models with EM reconstruction?
Answers to any parts or aspects of these confusions I have will be greatly appreciated!
I am a biophysics graduate student. One of my exam question is how to correct CTF in image reconstruction. After finishing the exam, I have more questions than answers!
What kinds of contrast exists in electron microscopic image for weak-phase object? Is amplitude contrast the same as aperture contrast?
What is the contribution of inelastic scattering on the image contrast? How to account for it?
How does partial spatial coherence and temporal coherence influence the image and electron diffraction?
Is it always sufficient to use just first order approximation ( the linear theory of phase and amplitude contrast image formation ) for CTF consideration?
Is it true for low spatial frequencies ignoring the CTF was better than compensating for phase contrast alone?
I guess compensation for the CTF was necessary and sufficient to accurately reconstruct molecular densities. Could anyone tell me the current ways for accurate determination of CTF?
By improving the different components in CTF, what is the optimal contrast could be achieved in image?
Is it possible to put EM reconstruction on the same scale with the structures derived from X-ray crystallography and NMR after deliberate correction of CTF, solvent effects and differences between atomic scattering factors for electron and X-ray?
To what extend we could use the insights obtained by building models and comparison of the models with EM reconstruction?
Answers to any parts or aspects of these confusions I have will be greatly appreciated!
} I am a biophysics graduate student. One of my exam question } is how to correct CTF in image reconstruction. After finishing } the exam, I have more questions than answers! } (snip - long list of questions deleted...)
You ask several good questions that space (and time) does not permit me to answer in detail. May I suggest that you check out the Ph. D. Thesis of Xiadong Zou (Electron Crystallography of Inorganic Structures - Theory and Practice.) This was published in the Chemical Communications of Stockholm University (1995 No. 5.) You may be able to get a copy from your library. If not, write the Department of Structural Chemistry, Arrhenius Laboratory, Stockholm University, S-106 91 Stockholm, Sweden. Xiadong's thesis advisor was Professor Sven Hovmoller.
In the first part of her thesis, Xiadong does a great job of summarizing the theory of electron imaging and diffraction in a unified fashion. One of the problems that we all face is that the theory was developed by several communities and the terminology and notation is often conflicting and confusing. For example, one needs to be careful to decide whether the author chooses underfocus to be negative or positive. Xiadong shows several examples of the effect of correction for CTF on image reconstruction.
-- Best Regards, John Minter
Eastman Kodak Company Phone: (716) 722-3407 Analytical Technology Division FAX: (716) 477-3029 Room 2112 Bldg 49 Kodak Park Site email: minter-at-kodak.com Rochester, NY 14562-3712 calendar: via PROFS
Haven't yet convinced the guys at work to spend the whopping {g} $400-600 to implement a CD-R, but do have one on the home system which works great for back-up/archiving. Is SCSI2 (aren't they all?) running EzCD Pro software.
My HP 6020 does not support "packet mode" so it can't be treated like a floppy, but does support "multi-session". It is not necessary to write "disk at once". The caveat is that for each writing "session" something like 13 to 20 Mb of CD space is consumed as "overhead". ...But at $6 for 650Mb, do we care?
As to compatibility... some old CD players (under W3.1?) read the first session and quit. When those systems were made, multisession was not commonly available.... I haven't yet found a system old enough that it would not read the CDs I burned.
Who really knows how long the CD-Rs will last (and will the hardware be around to read them?). TDK rates their disks for 100 years....
If your only goal is to get thicker, larger sections with GMA, I have a proceedure that doesn't require a special chamber or UV light. I have used this procedure for 15 years with very good results. The blocks stain well with toluidine blue, but little else, though. I use a modified embedding procedure for JB-4, developed by Ann Klinker and Marge Hukee 15 years ago. It does require the blocks to be set without air, we use parafilm sealed to the top of the mold. E-mail me if you are interested in the details. kna101 utdallas.edu
Karen Pawlowski
On Wed, 23 Jul 1997, C. John Runions wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Hi everyone, for many years I have been embedding in Spurr's resin but } have recently decided to give glycol methacrylate a whirl for thicker LM } sections of plant material. Many years have passed since my last work with } GMA. I remember that we made a plexigalss container which we could } evacuate the oxygen from by flushing with nitrogen and that polymerization } was then carried out below a longwave UV light placed in the container. } Does this sound correct? If so does anyone have any plans or tricks that } might assist me in constructing such a device? Cheers, John } } ================= } C. John Runions, Ph.D } Section of Ecology and Systematics } Corson Hall } Cornell University } Ithaca, New York } USA 14853 } } email cjr14-at-cornell.edu [ie. cjr(fourteen)-at-...] } phone (607) 254-4282 } Fax (607) 255-8088 } } }
John Hunt writes: } From: John Hunt {hunt-at-msc.cornell.edu} } Subject: Re: CD-ROM's for archiving - any experience? (fwd) } To: rick-at-warf.msc.cornell.edu (Rick Cochran) } Date: Thu, 24 Jul 1997 09:29:12 -0400 (EDT) } } I am forwarding this exchange from the microscopy listserver for } your interest.
Which OS is best to run your CDR under depends on what environment your date is being acquired in. The three environments which pop to mind are Unix, Windows, and Mac. CDR solutions are available for all three environments.
For our Unix environment, we have recently succeeded in setting up CDR under Linux using freeware utilities. You need a SCSI system with a suitably large disk, 'mkisofs' for mastering the CD images, and 'cdwrite' or 'cdrecord' for actually writing the CD. The only problem we ran into was that most CDR drives require the 'SCSI disconnect' feature which was disabled by default in the Linux SCSI driver for our SCSI adapter.
mkisofs is available at tsx-11.mit.edu:/pub/linux/packages/mkisofs
cdwrite and cdrecord are available at sunsite.unc.edu:/pub/Linux/utils/disk-management
Since there is no industry standard for the SCSI commands for CDR drives, support for each drive must be explicitly written into the burning utility. You should determine which drive cdwrite and cdrecord support before purchasing a drive.
-- |Rick Cochran phone: 607-255-7223| |Cornell Materials Science Center FAX: 607-255-3957| |E20 Clark Hall, Ithaca, N.Y. 14853 email: rick-at-msc.cornell.edu| | "The Founding Fathers did not establish the United States as a | | democratic republic so that elected officials would decide trivia, | | while all great questions would be decided by the judiciary." | | Judge Andrew Kleinfeld |
At 11:24 PM 7/23/97 -0400, you wrote: } My question is: Has anyone *ever* gone back to access and use original } image data that is 30 years old? I personally do not remember ever using } original images that were even 5 years old....
Actually, I used to work in a Pathology diagnostic EM lab and we certainly went back 20 years sometimes (rare diseases often require an entire career to accumulate enough examples to publish about). I imaging the need for archiving images is different for each field, but I still think its a bit short sighted to not consider the future while living in the present. Although this is off the science track, I wonder how our grandkids will manage to view our digital snapshots when we're pretty sure that dye-sub prints don't last that long and they need to find a viewer/reader for our family album CD-ROM disk.
} Perhaps we should realistically not place too great an emphasis on the } usefulness of saving all images for 30 years. CD-ROMs supposedly have that } capability, but who believes that today's CDs will be readable by any } technology available in 30 years? Anybody keep their old 8-track tape } players in good service? I think most digital image formats will have to } periodically be upgraded to the latest storage technologies, just as people } take 16 mm home movies and convert them over to video tapes. Of course, } this does not have to be done with film, and negatives probably will always } be able to be converted easily to hard copy many years in the future....but } I still never make images on film any more...
I may not have an 8-track (anymore), but how could we have the current crop of re-releases of old musical works (originally recorded as analog) if the music industry hadn't had a long lasting standard that they could still work with today? Remember too that movie to video transfer results in a lower resolution image (loss of information) stored on a tape format (I hope you meant VHS, not the now obsolete BetaMax) that probably has a 5 year lifetime.
Actually, I'm of the mindset that of the mass storage technology currently available today, CD-ROMs are the only ones likely to be readable in 30 years. That "Amazing Kreskin" like prediction (opinion) is based primarily on the large market presence of CD readers (one industry analyst believes that the installed base of CD devices will reach 150 million by the end of 1997, source: Advanced Imaging Magazine), compared with almost everything else.
Don't get me wrong, I like digital imaging for most of the same reasons everyone else likes it. I use it here at work and I try to teach students and staff about it because I think its the way science is going. I'm just concerned that we, as microscopists, are headed into this without realising all the issues related to archiving of and longevity of images.
Yours, Doug ..................................................................... : Douglas W. Cromey, M.S. Dept. of Cell Biology & Anatomy : : Sr. Research Specialist University of Arizona : : (office: AHSC 4212A) P.O. Box 245044 : : (voice: 520-626-2824) Tucson, AZ 85724-5044 USA : : (FAX: 520-626-2097) (email: doug-cromey-at-ns.arizona.edu) : :...................................................................: http://www.pharmacy.arizona.edu/exp_path.html
Damian Neuberger mentioned that he had been told that ZIP drives were being discontinued. I think his source of information must have been wrong. Iomega recently introduced an internal SCSI version in addition to already existing external parallel port and external SCSI. A number of computer companies are now offering ZIP drives as standard equipment. We have used ZIP drives for several years on this campus for storing images from TEMs, SEMs, AFM/STMs, and Laser Confocal microscopes and we have been extremely pleased with them. The disks are sold by our local Best Buy Store as well as at most computer stores and Iomega has licensed other manufactures to produce the disks. We also have the option of storing images on a Panasonic Read/Write Magneto-Optical drive (1 GigaByte disks)on our SEM, but most users prefer the ZIP. I for one don't like the idea of having a thousand or so images on one disk. I much prefer to have several ZIP disks with a hundred or so images on each. Stanley L. Flegler, Assistant Director Center for Electron Optics Michigan State University
I can add little to what Jim Darley has said about setting up the knifemaker but I was wondering whether you used to get good knives and now get bad or have just started using the LKB.
Do you have the instruction charts for the knifemaker? Our LKB 7801A has a glossy card 4 page manual and a condensed instruction laminated chart which are very useful, when you understand them. They are normally hidden under the machine on a little purpose built tray.
When you say that the reflection line is the wrong way do you mean that the 'meniscus' in the glass curves the wrong way? This might depend on which way up you score your rhomboid/squares to make the triangles. We make 50 degree knives from 100/80 degree rhomboids by first producing the rhomboids by scoring on the roughened edge side of the glass strips. Then to bisect the rhomboid we turn it over so all the rough edges are at the bottom. (I hope you follow my meaning).
Finally what's your glass like - it hasn't been dropped or damaged. Is it the right stuff for e.m. - we use 6mm thick glass from a reputable e.m. supplier.
I hope this is of help but I'm sure that if you combine all of the replies you will solve your problem.
Malcolm Haswell e.m. unit University of Sunderland U.K. ----------
Sharon et.al. That knife maker is an everlasting instrument but the adjustments must be right, otherwise its a big waste of glass and time. {SNIP} ---------------------------------------------------------------------------- ------------------------- } From: Sharon C. Thomas {sthomas-at-lanl.gov} } To: Microscopy-at-sparc5.microscopy.com } Subject: Help requested for maladjusted glass knifemaker } Date: Wednesday, 23 July 1997 3:44 } The knifemaker available to me is a LKB Type 7801A from LKB Instruments Inc. } out of Rockville Maryland, but the company has since been taken over by } Leica/Leo. The knifemaker has two wheel type gauges that adjust the tension } from opposing sides on the glass piece. I can make glass squares, but when } I try to cut the square into the two knives, I don't get the correct shapes, even } though I have tried systematically changing the settings on the two gauges } many different ways. Typically, the "sharp" edge may be chipped, the } reflection line in the glass is going in the opposite direction to what it should } and the opposite edge to the sharp edge may be either sharp as well or too } thick a blunt edge. That's just one of the two knives; its' counterpart often has } two "sharp" edges. Can anyone with any direct experience with this model of } knifemaker provide me with any advice on how to adjust the settings to produce } two glass knives? } } Thanks in advance for any assistance that can be provided.
I got a message from G. Arentieri asking for help. There was no return address, so I cannot address him directly. I can do a discussion of long term storage of tissue, but it is not of general interest to the group, so I would like to send it to G.Arentieri directly. Greg, Could you try and contact me again? This time type your e-mail address just in case of another snafu. Bye, Hildy
I have been transfecting mammalian cells with a dopamine receptor and trying to detect, via fluorecence microscopy, its expression with a polycolonal antibody that I made to one of this dopamine receptor's intracellular loops. (The antibody detects the dopamine receptor epitope, to which the antibody was made, on western blots, when expressed by E. coli in the context of a fusion protein, so I know the antibody does recognize the epitope against which it was made.)
When I express the receptor in transfected HEK293 cells, fix the cells by methanol:acetone, and do fluorescence microscopy, I get no detection of receptor expression. I engineered a " FLAG tag" into the receptor and stained for FLAG labeling;the FLAG epiope gave intense staining, so I know the dopamine receptor is being expressed.
Does anyone have any thoughts on what factors I might to alter in my immunofluorescence assay in order to make the dopamine receptor antibody recognize the dopamine receptor in the transfected cells? It seems as if the receptor is folded in a conformation such that the epitope that the antibody recognizes is buried / not exposed. Are there better/more appropriate fixation methods? Does anyone have information on live staining or microwave heating/steaming of the cells?
Hello there, MAS Members and other interested parties, The latest issue of The Microbeam Analysis Society Newsletter, MicroNews (Summer 97), is now available on the Web at:
John Mansfield North Campus Electron Microbeam Analysis Laboratory 417 SRB, University of Michigan 2455 Hayward, Ann Arbor MI 48109-2143 Phone: (313) 936-3352 FAX (313) 936-3352 Cellular Phone: (313) 715-2510 (Leaving a phone message at 936-3352 is preferable to 715-2510) Email: jfmjfm-at-engin.umich.edu URL: http://emalwww.engin.umich.edu/people/jfmjfm/jfmjfm.html
Dear all, I need some information. I have a Hummer VI-a sputter coater and I am having trouble finding where to get targets. EMCORP which sold the instrument seems to be out of business. Their phone numbers do not work. Any help will be appreciated.
TIA Greg Rudomen University microscopy Imaging Center S.U.N.Y. -at- Stony Brook
I got a message from G. Arentieri asking for help. There was no return address, so I cannot address him directly. I can do a discussion of long term storage of tissue, but it is not of general interest to the group, so I would like to send it to G.Arentieri directly. Greg, Could you try and contact me again? This time type your e-mail address just in case of another snafu. Bye, Hildy
I for one would be very interested in Hildy Crowley's opinions on long term tissue storage. Others ...?
Geoff -- *************************************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane Piscataway, NJ 08854 voice: (732)-235-4583; fax -4029 e-mail: mcauliff-at-umdnj.edu ***************************************************************
The first thing I was wondering: Is the epitope extra or intra cellular? I guess the most success I've had is by manipulating the fixation. Trying the solvents vs. 2-4% paraformaldehyde or something like Zambonies or Bouins so the picric acid can go in fast and stablize the protien then the paraformaldehyde lightly crosslinks. Then if you need to open up the cell I use .1% Tween or to open epitopes a very short 1- 3min .01% Trypsin.
Bob Morphology Core
On Thu, 24 Jul 1997, kelly dowhower wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } I have been transfecting mammalian cells with a dopamine receptor and } trying to detect, via fluorecence microscopy, its expression with a } polycolonal antibody that I made to one of this dopamine receptor's } intracellular loops. (The antibody detects the dopamine receptor epitope, } to which the antibody was made, on western blots, when expressed by E. coli } in the context of a fusion protein, so I know the antibody does recognize } the epitope against which it was made.) } } When I express the receptor in transfected HEK293 cells, fix the cells } by methanol:acetone, and do fluorescence microscopy, I get no detection of } receptor expression. I engineered a " FLAG tag" into the receptor and } stained for FLAG labeling;the FLAG epiope gave intense staining, so I know } the dopamine receptor is being expressed. } } Does anyone have any thoughts on what factors I might to alter in my } immunofluorescence assay in order to make the dopamine receptor antibody } recognize the dopamine receptor in the transfected cells? It seems as if } the receptor is folded in a conformation such that the epitope that the } antibody recognizes is buried / not exposed. Are there better/more } appropriate fixation methods? Does anyone have information on live } staining or microwave heating/steaming of the cells? } } Thanks for your input. } } Kelly Karpa } kjd136-at-psu.edu } } }
I have an undergraduate student working on the design of a homemade backscatter diffraction pattern for the SEM. We are trying to get patterns using a JEOL T300. We tried using an accelerating voltage of up to 30keV, turned up the spot size, turned down the gun bias, and we get a "bright" screen but no kikuchi lines. We tried different specimen tilts, specimen working distances, and screen to specimen distances, and the specimen is a good quality Si wafer, but nothing seems to produce a bright enough signal to generate the Kikuchi lines. Are we limited by the SEM we are using, or are we just missing something?
Thanks. Lucille Giannuzzi
************************************************************************* Lucille A. Giannuzzi, Ph.D. Assistant Professor
Dept. of Mechanical, Materials, and Aerospace Eng.
University of Central Florida phone (407) 823-5770 PO Box 162450 fax (407) 823-0208 4000 Central Florida Blvd. email lag-at-pegasus.cc.ucf.edu Orlando, FL 32816-2450 USA *************************************************************************
Zip drives are not yet discontinued...but they probably have a shorter market life-span than CD-R's (hard to predict the future though).
Optical drives are neat for archiving...but who has an optical drive on thier home/office system?? They are rather pricy.
CD-R's are the way to go I believe. Almost all computers these days have CD devices. The Disk and the drives are rather inexpensive. The storage life of a CD (not in vacuum) is over 60 years (and far longer in Vacuum storage). We store Tiff files and use HTML frontends so that almost ANYONE with ANY SYSTEM with ANY SOFTWARE can use them (OK...maybe not...but I have heard of no problems so far). They just make sence. We do not use a dedicated machine...But I would suggest to anyone that they at least use a dedicated h-drive for writing to (helps alot). I guess if a lab does not allow alot of access to archives then maybe an Optical drive would be better (less need for compatability).
On the subject of archiving medium going out of date:
At first thought one might think this is a horrible problem. We have about 200 old 8 inch floppies and no drive to read them. Further...I am not so sure how many of them are any good. They contain alot of years of data.
Yet, and I know this may not apply to everyone, what is the data worth?? I have found that the archiving Medium and devices seem to out last the Data. Any Data around here that are more than about 10 years old are almost worth-less. The machines used to collect them are long since replaced with better machines...we would have to recollect any archived data that old. Further, almost all older data worth note has been published and exists in hard copy some place.
Its neat to think that 100 years from now, someone some place will want to see some of my data, or a perfect BSE image I took. When that thought comes to mind I get a bit excited...then suddenly a new thought appears, "Yeah Right Christopher!! Keep Dreaming!". Think about the machines that will be in use 100 years from now.
I'll start with some notes on Wednesday's sessions since I neglected to summarize them last night. I was, er, a bit slow by the time I got back. :-P The session focus on Wednesday was "History and Art," although that was only loosely held to. Several of the papers dealt with artistic subjects but not art conservation per se.
Gary Laughlin spoke about "A Unique Metallurgical Process From the Early Bronze Age" in which he described the findings at, and significance of, a site excavated at the Kestel Mine in the Taurus Mountains of Turkey. The site dates from the third millenium BC. The examinations indicate that cassiterite ore was mined and refined at the site to yield a black magnetic oxide. This would have been more readily separated, due to its magnetism, than could be accomplished by other means.
Dr. McCrone gave an "Update on the Turin Shroud" in which he reviewed several letters he received from Father Rinaldi prior to the Father's death in 1993. In the letters, Father Rinaldi effectively acknowledged the proof that the Shroud was painted and actually dated from much later than the time of Christ's crucifiction, thus was not the true article.
(For those of you who may not know, Dr. McCrone concluded early on in the Shroud investigations, and from microscopic observations alone, that the shroud was a painting. He stood nearly alone in this view and was vilified for nearly two decades before Carbon dating ultimately confirmed his conclusions.)
John Delly gave one of his typical, amazing presentations, this one on "Hand-colored Microscopical Illustrations." In it John took his audience on a delightful stroll through 19th century microscopy publications illuminated with hand-colored illustrations. He demonstrated the evolution and later de-evolution of the quality of such illustrations, the variation that one can see from different illustrators of the same work, and the differences that are seen edition-to-edition of the same work. Of course, when he became interested in the subject, John felt compelled to master the art himself. Through his own study and practice he gained the insignt necessary to understand and explain the techniques and variations seen in these historical works. The illustrations are, indeed, beautiful and many of us are fortunate to own examples of these illustrations in early works on microscopy. Because of the vast number of color illustrations necessary to address his subject, it is unlikely that this presentation will ever be recorded fully in print. (How about a book, John?) Those of us fortunate enough to be in the room may be the only ones ever to enjoy this particular product of John's efforts. Thank you, John, once again.
Have you ever stood befor an audience wondering why on earth you found yourself presenting in the particular session where you were? Wayne Moorehead must have when he spoke on "A Tale of Two Danas; Influences in Mineralogy" but he soldiered on and did a fine job chronicaling the lives of two remarkable men. The mineralogists in the audience will need no introduction to the Danas but I'll just mention for the others that the elder Dana published his first edition of the System of Mineralogy in 1835 at the tender age of 24. It was the first such major scientific work of classification written in English and he and his son went on to publish or edit a vast array of classic works in Mineralogy. Together or individually, they edited the prestigious American Journal of Science continuously for an astounding 95 years, from 1840 to 1935. One of the most noteable achievements that Wayne mentioned, in my mind anyway, was when James Dana, in the introduction to a revised edition of his classic System, abruptly abandoned his entire earlier classification system as outdated!. Believing that system no longer consistant with emerging thought, he just as abruptly adopted and described a newer system which largely stays with us today. I find such honest self-appraisal and the ability to continue to move forward without missing stride quite refreshing.
Wednesday afternoon was occupied by two sessions which would be unusual at other meetings. Using video microscopy, Anna Teetsov of McCrone Associates demonstrated some micromanipulation techniques within the context of creating artistic works on microscope slides by arranging butterfly scales of various colors into micro-images. Anna and a few others continue to develop this art form which is particulary unique to the community of microscopists. One has to have a microscope and micro-related knowledge and skills in order to produce these beautiful little creations, then one has to have a microscope to view them as well. Kind of nice, don't you think? Something we can hold purely for the aesthetic pleasure and uniquely our own.
The afternoon was closed with a demonstration by Alan Shin on how one can construct a working replica of Leeuwenhoek's single lens microscopes. I did not attend this demonstration as I have on a previous occasion taken a longer version from Alan in which we got to actually construct our own microscopes. Comments from those who did attend and look through the instrument Alan made reflected surprise at how much one could see and pleasure at the experience of seeing an insturment of such historical significance actually fabricated.
Today's sessions were on Forensic Microscopy. Jose Almirall told us about "Developments in Glass Examination: Automated Microscopy Techniques and Composition Analysis." Jose's talk was very interesting and perhaps somewhat troubling to practicing forensic scientists as he told us of (among other things) a remarkable consistancy in the optical properties of some glasses, especially window glass manufactured by the "float" process. The new information for me was the time over which the products of these plants will remain indistinguishable under conventional forensic examination techniques. I am not aware of other time-dependant studies of glass properties but Jose showed data collected over at least 18 months, during which the product of a float glass plant was entirely uniform in refractive index. He did however, offer a remedy for this disturbing finding. He showed that glass samples which are indistinguishable by refractive index can often be distinguished by elemental analysis of Fe, Mg, Al, and Zr using ICP/AES. Now all the forensic people have to do is get themselves one of these and... ;-)
Wayne Moorehead gave another excellent paper on Thursday, this one on "An Introduction to Microscopical Feather Identification." Wayne told us that the flight and tail feathers of birds are not always useful for identification but that the down or contour (breast) feathers can be distinctive, at least down to the order of birds, occasionally to the family, but virtually never to the genus or species. Still, identification at this level may prove very useful in a forensic case. Wayne illustrated how appropriate preparations can be made, what features of the feather barbule to examine and how they can vary. He also showed and described the identifying characteristic of numerous feather types.
Thom Hopen gave an interesting talk on "Teaching Forensic Microscopy in Countries Formerly Known as the Soviet Union." Thom responded to a State Department request that he make numerous trips to various countries of the former Soviet Union. He has taught courses of fiber and paint comparison, explosives residue analysis, and basic microscopy. He found his students to be highly motivated and dedicated people, anxious for quality instruction in basic forensic microscopy techniques. Often they are at least adequately equiped though sometimes have little or no idea how to fully exploit the equipment they have. (Unfortunately, when it comes to microscopy, this is too often true here also! My comment, not Thom's.) One can only immagine the difficulty of teaching in a completely and literally foreign environment, working through a translator, and using instrumentation previously never seen. Often, Thom had to set the equipment in proper working order prior to beginning instruction. But apparently all has worked out for him and his students and several more trips are planned to continue the education.
Well folks, I think I'll stop there. Of course, there were many more fine presentations and, once again, I'll mention that my choice of topics covered here in no way reflects badly on the other papers. All of the presentations were excellent.
Tomorrow is given over to a tutorial workshop on the Dispersion Staining technique. It will be given at McCrone Research Institute by Dr. McCrone and will be attended by twenty-odd students, all that can reasonably be accomodated in such a hands-on session. For the rest of us, this afternoon marked the end of another educational, enlightening, and just plain fun Inter/Micro.
Special thanks, as usual, to Nancy Daerr who coordinates all arrangements for these meetings and who, as usual, did an exceptional job of taking care of us and making our stay wholly enjoyable.
To all of those interested in these meetings, please note: Next year marks the Golden Anniversary of Inter/Micro, the fiftieth anniversary. (Wow!) Plan on attending what promises to be an excellent meeting. Contact Nancy Daerr for further information, to be put on a mailing list, etc. She can be reached at McCrone Research Institute, 2820 S. Michigan Avenue, Chicago, IL, 60616 or simply as ndaerr-at-mcri.org. The phone numbers at McRI are 312-842-7100 (voice) and 312-842-1078 (fax).
It's been a pleasure being your ears at Inter/Micro 97. But tomorrow... Ahhh, Chicago! The architecture, the museums, the Art Institute! I feel like a nice walk! Happy trails to all, and to all, Good Night.
Steve Shaffer MicroDataware sshaffer-at-microdataware.com
Please post or pass this on as appropriate. Thank you.
Regards,
John Vetrano
----------
Electron Microscopy Positions in Materials Characterization
Pacific Northwest National Laboratory
The Structural Materials Research Group at Pacific Northwest National Laboratory is seeking applicants for a staff position and a post-doctoral appointment, both specializing in the characterization of materials using transmission electron microscopy (TEM). The Electron Microscopist staff position requires a minimum of a 2-year specialized degree with expertise and training in the operation of analytical TEM instruments as well as in the preparation of electron transparent materials for examination. This individual will conduct research on a wide variety of metallic, ceramic and composite materials in support of scientists and engineers in the Structural Materials Research Group. Current activities include both basic and applied research in the areas of deformation mechanisms in metals, interfacial segregation and precipitation, radiation effects in metals, ceramics and composites, and enviromental degradation. In addition, the position will help oversee the maintainance of the Group microscopy facilities which include three conventional TEMs, a field-emission-gun (FEG) TEM and a scanning TEM, SEMs and a scanning Auger microprobe.
A post-doctoral position is also available focussed on the high-resolution characterization of grain boundaries in metallic alloys by analytical TEM. Energy dispersive x-ray and electron energy loss spectroscopies will be used with the FEG-TEM to elucidate segregation and precipitation mechanisms in aluminum and stainless steel alloys. The initial post-doctoral position would be for one-year and a second-year renewal is possible. The position requires a PhD degree in material science, physics or a related discipline and experience in analytical TEM for quantitative compositional analysis.
Pacific Northwest National Laboratory (PNNL) is operated by Battelle Memorial Institute for the U.S. Department of Energy. PNNL is a multiprogram laboratory located at the Hanford site near the junction of the Columbia, Snake and Yakima rivers in SE Washington state.
For consideration, please submit a resume (including references) and publication list by Oct. 1, 1997 to: Dr. Stephen M. Bruemmer, Pacific Northwest National Laboratory, Battelle Boulevard, P.O. Box 999, Richland, WA 99352. Phone: (509) 376-0636; Fax: (509) 376-6308; e-mail: sm_bruemmer-at-pnl.gov.
Using different Apple Macintosh Workstations and one mobile Apple Powerbook 1400cs (which has an 8xCD-ROM drive), the CD-R Backup solution proved perfect for Software and Data Backup, and taking complete backup with the Powerbook still leaves me very mobile.
I use the Philips 2660 CD-Burner with Astarte Toast CD Software. This not only allows for different formats, but also for creation of CD-ROM image files on harddisk for easy preparation of a 650M partition, or smaller. If the Macintosh is setup well, and the driver software is properly installed (I use the FWB-CD-driver although not recommended by Astarte), no problems are expected to occur.
To deal with the fact, that some older machines do not read multi session CD-ROMs, I installed a 1G-Harddisk to collect the data to be burned on the CD on a 650M image file. As soon as it is full, I burn 2 CDs (one for regular use, one for backup), and throw the image file away. This proved itself useful, because at the time I burn the CD, some files are no longer needed and can be thrown away before burning.
A known fact with Floppy Disks is the time they keep the data until they begin to loose it (1-2 years? depending on brand?). This issue might be important and also brand-dependent in CDs as well, but the extent, the expected timerange and hence significane is unknown to me.
Wolf Schweitzer, MD Research Fellow, VIFM, Melbourne, Australia wschweitzer-at-access.ch
This listserver is a great help to me as an amature microsopist. I enjoy the information obtained by reading the daily "chat". I work 8 hours a day in the computer business. My suggestion is never trust any technology of today to last very long. The abiliity to store files for a long time is worthless if the technology on which it is stored is antiquated by as much as 3 - 5 years.
Zip drives are hot technology right now, but becauase of their limitations of size, at this point, they are being passed by with upcoming technology. File storage is good for the time being, but in less than a year look for the most modern and up to date file storage technology to change. Computers and their technology are always changing and becoming less expensive as the R&D is paid off and manufacture of newer ideas comes into being.
Do not hitch your wagon to anything as a forever thing. If silver grain photographs are s "thing of the past", whatever we use today will be also, very soon. CDRom drives and platters are fantastic, but technology to handle 1000 times the information storage in a smaller package via opti-digital technology is somewhere close around the corner.
I am not a proponent of either silver grain or digital file storage. Know that your will have to provide the latest thing for your business client there is and that will always change. Two years ago 4X CDRom drives were all the rage. Today you have to invest in a 16X or 20X drive that is less expensive than the 4X was. 4 gigbit hard drives retail for less than half the cost of a 540 megabit drive 5 years ago.
Sometimes it dosen't matter if the data will actually ever be accessed. It must just be available. Certain regulations (QA type stuff) can require data archiving for 30 years or more.
Any target the correct diameter will work. What I have done is clean-up the spent target assembly, then silver-epoxy a new (generic) disk to the target assembly face. Most EM suppliers sell disks. Material can also be purchased from suppliers like Alfa Aesar (Johnson Matthey) or Goodfellow. There is a supplier in Nevada (I think) who is reputed to be less expensive, but I don't have any more info available...Maybe someone else knows that name/address.
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear all, I need some information. I have a Hummer VI-a sputter coater and I am having trouble finding where to get targets. EMCORP which sold the instrument seems to be out of business. Their phone numbers do not work. Any help will be appreciated.
TIA Greg Rudomen University microscopy Imaging Center S.U.N.Y. -at- Stony Brook
Greg wrote: } } ------------------------------------------------------------------------} The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------.} } Dear all, } I need some information. I have a Hummer VI-a sputter coater and I am } having trouble finding where to get targets. EMCORP which sold the } instrument seems to be out of business. Their phone numbers do not work. } Any help will be appreciated. } } TIA } Greg Rudomen } University microscopy Imaging Center } S.U.N.Y. -at- Stony Brook
Greg, We supply targets for all models of Hummer sputter coaters and almost any other type you might need. Please let me know what material and I will send you a quote. Thanks, JD Arnott Ladd Research
Greg wrote: } } ------------------------------------------------------------------------} The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------.} } Dear all, } I need some information. I have a Hummer VI-a sputter coater and I am } having trouble finding where to get targets. EMCORP which sold the } instrument seems to be out of business. Their phone numbers do not work. } Any help will be appreciated. } } TIA } Greg Rudomen } University microscopy Imaging Center } S.U.N.Y. -at- Stony Brook
Greg, We supply targets for all models of Hummer sputter coaters and almost any other type you might need. Please let me know what material and I will send you a quote. Thanks, JD Arnott
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear all, I need some information. I have a Hummer VI-a sputter coater and I am having trouble finding where to get targets. EMCORP which sold the instrument seems to be out of business. Their phone numbers do not work. Any help will be appreciated.
TIA Greg Rudomen University microscopy Imaging Center S.U.N.Y. -at- Stony Brook
Message-Id: {3.0.1.32.19970725104540.007717d4-at-mmserver.mm} X-Sender: opmills-at-mmserver.mm X-Mailer: Windows Eudora Pro Version 3.0.1 (32)
Hello,
I'm posting this query for a colleague here at MTU. Please contact him directly at qchorn-at-mtu.edu. Thanks.
Owen
++++++++++++++++++++++++
While characterizing the microstructure of a commercial ferrosilicon alloy, an interesting contrast event was observed within the silicon phase. The center of the silicon dendrites appears brighter than the edges when secondary electron imaging is used. This contrast is not observed using backscattered electron imaging. The following web site has images showing this contrast as well as other information about the alloy being examined:
http://www.mm.mtu.edu/~qchorn/siliconquestion.htm
Any insight as to what could be causing this contrast would be greatly appreciated.
Thanks,
Quinn C. Horn ++++++++++++++++++++++++++ Owen P. Mills Michigan Technological University Metallurgical & Materials Engineering Rm 512 MME Building Houghton, MI 49931 906-487-2002 906-487-2934 FAX opmills-at-mtu.edu
I agree with your comments that we sometimes try to save "all" of the data we generate, and much of it is of little future value. But having worked in the medical device industry for 20 years now, I would like to inject my two cents, just to provide another viewpoint.
In industry, some data (regardless of its age) is absolutely invaluable. An example would be data that is generated to support studies of the safety/efficacy of an implantable device. (Silicone breast implants are a good example). If 20 years from now, litigation arises with regard to one of our products, raw data may be required to lend credence to our studies. In fact, the FDA requires that we maintain certain data "forever", and some of this data is electronic. With this in mind, I have wrestled with this topic of data archiving and integrity for several years now. I try to ensure future compatibility of whatever software and hardware upgrades I make in our data collection equipment, but it's not always possible. Consequently, I have made it a point to also archive some of the older hardware and software, just in case.
Best Regards,
Bob ********************************** Bob Citron Chiron Vision Claremont, CA USA (909)399-1311 Bob_Citron-at-cc.chiron.com **********************************
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On the subject of archiving:
Zip drives are not yet discontinued...but they probably have a shorter market life-span than CD-R's (hard to predict the future though).
Optical drives are neat for archiving...but who has an optical drive on thier home/office system?? They are rather pricy.
CD-R's are the way to go I believe. Almost all computers these days have CD devices. The Disk and the drives are rather inexpensive. The storage life of a CD (not in vacuum) is over 60 years (and far longer in Vacuum storage). We store Tiff files and use HTML frontends so that almost ANYONE with ANY SYSTEM with ANY SOFTWARE can use them (OK...maybe not...but I have heard of no problems so far). They just make sence. We do not use a dedicated machine...But I would suggest to anyone that they at least use a dedicated h-drive for writing to (helps alot). I guess if a lab does not allow alot of access to archives then maybe an Optical drive would be better (less need for compatability).
On the subject of archiving medium going out of date:
At first thought one might think this is a horrible problem. We have about 200 old 8 inch floppies and no drive to read them. Further...I am not so sure how many of them are any good. They contain alot of years of data.
Yet, and I know this may not apply to everyone, what is the data worth?? I have found that the archiving Medium and devices seem to out last the Data. Any Data around here that are more than about 10 years old are almost worth-less. The machines used to collect them are long since replaced with better machines...we would have to recollect any archived data that old. Further, almost all older data worth note has been published and exists in hard copy some place.
Its neat to think that 100 years from now, someone some place will want to see some of my data, or a perfect BSE image I took. When that thought comes to mind I get a bit excited...then suddenly a new thought appears, "Yeah Right Christopher!! Keep Dreaming!". Think about the machines that will be in use 100 years from now.
I just want to publicly thank Steve Shaffer for the interesting summaries of the talks at Inter/Micro. I hope I speak for all when I say that those of us who (for whatever reason, be it time, money, other committments) could not attend, really appreciated it. Hope you enjoyed the "off" time as well!
Best Regards,
Bob ************************** Bob Citron Chiron Vision Claremont, CA USA Bob_Citron-at-cc.chiron.com **************************
Has anyone thought of something useful to do with all the leftovers from dye sub printers? I have boxes of used ribbons and other things that seem too good just to toss out. Could they be used for something, or maybe recycled somehow?
Jonathan Krupp Microscopy and Imaging Lab University of California Santa Cruz, CA 95064 (408) 459-2477 FAX (408) 429-0146 jmkrupp-at-cats.ucsc.edu
Hi everyone, I have a couple of announcements about tickets to the Cleveland Indians game on Tues evening. One - If you have reseved tickets, I need to see the money!! Please send a check made out to 'MSA' for $20/ticket (cheap seats, but includes a picnic) to me here at:
Ferro Corp. 7500 E. Pleasant Valley Rd. Independence, OH 44131
Two - If you plan on canceling your reservations, I've had a number of requests for more tickets...So - I'm starting a waiting list. First come, first on the list. I have had a few cancelations, so far - so please give me a call at (216)641-8585 x6613. E-mail address is: vbauer-at-ferro.geis.com.
I know this isn't "Microscopy" but I don't have current e-mail addresses for everyone, so please bear with me.
One woman that used to work in our lab thought the ribbons would be good for Halloween--go to a party as the "Primary Colors"
Thomas Moninger moninger-at-emiris.iaf.uiowa.edu University of Iowa Central Microscopy Research Facility http://www.uiowa.edu/~cemrf Views expressed are mine. On Fri, 25 Jul 1997, Jon Krupp wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Hi: } } Has anyone thought of something useful to do with all the leftovers from } dye sub printers? I have boxes of used ribbons and other things that seem } too good just to toss out. Could they be used for something, or maybe } recycled somehow? } } Jonathan Krupp } Microscopy and Imaging Lab } University of California } Santa Cruz, CA 95064 } (408) 459-2477 } FAX (408) 429-0146 } jmkrupp-at-cats.ucsc.edu } }
Subject: Time:12:45 PM OFFICE MEMO Philips 201 TEM Date:7/25/97
If anyone is interested in obtaining a Philips 201 (24 years old, on service contact continually) for a nominal cost, please contact me immediately. Ann E. Rushing Department of Biology Baylor University Waco, TX 76798 Ann_Rushing-at-baylor.edu (254) 755-2911
Bob Citron wrote: ================================================== I agree with your comments that we sometimes try to save "all" of the data we generate, and much of it is of little future value. But having worked in the medical device industry for 20 years now, I would like to inject my two cents, just to provide another viewpoint. ================================================== Actually, it is not just the medical device industry that has these concerns . The entire industry of analytical and testing laboratories have these concerns as well. You see, in our litigious society in America, there is no such thing as a "statute of limitations" for professional liability exposure (negligence as some would say). The "clock" starts ticking, not when the alleged error occurred, but when the alleged error is discovered. This means, in our case, we have to maintain complete records of all work done going back to the inception of our business in 1970 with all of the associated costs.
When these records are not maintained or are not kept available in retrievable form, then someone who is trying to make it seem like you have made some error years ago, really is in the driver's seat, that is, they can say whatever they want to say and you have nothing in your files that says otherwise. Worse yet, the people who did the work might not remember what really did go on, say twenty years ago, or can be either retired, no longer alive or just not anywhere to be found.
And what has to be kept is not only the original data from specific samples, but also the ancient records showing service history, instrument calibrations, and other test and measurement procedures that document that the instrumentation was operating the way it was claimed to have been operating.
Are there really instances where old archival information of this type is actually needed? You bet. Do laboratories or individual consultants get sued? You bet. And how do they fare without such supporting data and information? Not very well.
Disclaimer: From 9/76-3/97 served as shareholder and director of ILAC Ltd., Hamilton, Bermuda, an insurance company that insures analytical and testing laboratories for professional liability risk.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI Structure Probe, Inc. FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
Dear All, The supplier of sputter coater targets in Nevada is Abe Dayani of Refining Systems Inc. P.O. Box 72466 Las Vegas, NV 89170 phone: 702-368-057 fax: 702-368-0933 He is a buyer of estate jewelry and will make discs of any precious metal or alloy for much closer to the list price of the metal than most suppliers. I believe that if you send him a target he will stick the new disc on it. Woody wrote: } Any target the correct diameter will work. What I have done is clean-up } the } spent target assembly, then silver-epoxy a new (generic) disk to the target } assembly face. Most EM suppliers sell disks. Material can also be } purchased } from suppliers like Alfa Aesar (Johnson Matthey) or Goodfellow. There is a } supplier in Nevada (I think) who is reputed to be less expensive, but I } don't } have any more info available...Maybe someone else knows that name/address. } } Woody White } Mcdermott Technology Inc. } } Alt: } woody.white-at-worldnet.att.net } http://www.geocities.com/capecanaveral/3722 } ______________________________ Reply Separator } _________________________________ } Subject: Hummer VI-a sputter coater } Author: greg-at-umic.sunysb.edu_at_internet at X400post } Date: 7/24/97 2:59 PM } } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
For those of you who are not already familiar with it, I would like to introduce the Society for Luminescence Microscopy and Spectroscopy (SLMS). Members of this society are involved in cathodoluminescence (CL) and UV-excited fluorescence microscopy. Most of the members are involved with the earth sciences or the material sciences and ceramics. There are also some applications in forensics, archaeology, and other fields. The society has been in existence for about 10 years. There is a semi-annual newsletter, edited by Professor Kopp of the U. Tennessee. Dues are only $10.00 for full members and $5.00 for students. The instrumentation used includes the familiar SEMs and EMPAs and a significant number of the members are using relatively simple and inexpensive cold cathode based electron beam systems which are small enough in size that they can be mounted directly on the stage of a conventional monocular or binocular transmitted light microscope. The CL of the mineral samples can then be seen directly, in real time, with a minimum of sample preparation and the information revealed on impurity distributions, etc, is otherwise very difficult or impossible to obtain with other existing techniques.
The SLMS has a standards program which has dealt, so far, with the comparison of photographic results on a complexly zoned carbonate and with the comparison of the CL emission spectra from a Dy-doped zircon.
For more information on the SLMS, please visit our web site at http://zephyr.rice.edu/SLMS/SLMS.html This web site is maintained by Jinny Sisson at Rice University.
If you have any questions regarding the SLMS or the applications of CL, please feel free to contact me or one of the addresses on the web page.
Disclaimer: I am one of the many charter members of the SLMS and presently chairperson of the Standards Committee. I have a commercial interest in furnishing cold cathode based ebeam systems for CL investigations.
Donald J. Marshall
Donald J. Marshall Relion Industries P.O. Box 12 Bedford, MA 01730 Ph: 617-275-4695 FAX: 617-271-0252
To further comments from Robert H. Olley and Cynthia Bennett regarding plasma etching, I would like to share a little insight. Ion damage and "burn marks" are related to the type of plasma generation system. In the past several years there has been a significant amount of effort expended to reduce the ion energies present in the plasma. The greatest drive has been in the semiconductor industry which requires absolute cleaning or etching without any uncontrolled altering of the material being processed.
Two types of low energy plasma creation have been developed as a result, namely, inductively coupled and electron cyclotron resonance (ECR) which create ion energies of 10-15eV and 3-5eV, respectively. At these ion potentials, insufficient energy exists to either heat or alter the specimen. Organics are removed from the surface, or material is selectively removed from the specimen/substrate through the application of reactive gas species which are generated by the plasma. This process is purely chemical and does not rely on the forces of ion impingement to remove material.
To remove organics, oxygen is the ideal process gas which chemically converts hydrocarbons to CO, CO2 and H2O. To selectively etch materials, various process gases can be applied. This technology can be readily utilized without any concern over damage to the specimen. Robert Olley's application of the atomic oxygen generated in a radio frequency discharge began to touch on an appropriate application of plasma.
I hope that this clarifies some of the concerns over the use of plasmas.
Regards,
Paul
Paul E. Fischione E.A. Fischione Instruments, Inc. 9003 Corporate Circle Export, PA 15632 USA Phone (412)325-5444 FAX (412)325-5443 Web-site: www.fischione.com
Does anyone have a source for plastic slides/coverslips/dishes that can be used for tissue culture and won't autofluoresce? Some of the catalogues list products, but I haven't tried any of them and would like some feedback from the experts :)
Thanks to all who responded to my Hummer Sputter target question. Many of you have an adress for Anatech that is old. The new address is: Anatech LTD. 6621-F Electronic Dr. Springfield, Va 22151-4303 1-800-752-7629, Fax 703-941-8077
Greg Rudomen S.U.N.Y. at Stony Brook University Microscopy Imaging Center greg-at-umic.sunysb.edu
Greetings All, I'm curious about how often everyone aligns the beam on their 'scopes. I use a JEOL - JEM 100CX II. In the past, the alignment procedure was done every morning. Now, we only do it once a week. We don't seem to have any problems, and the photos are crisp. What are others doing?
Sharron G. Chism HT (ASCP) Electron Microscopy Lab Harris Methodist Hospital Fort Worth, Texas
} I'm curious about how often everyone aligns the beam on their 'scopes. } I use a JEOL - JEM 100CX II. In the past, the alignment procedure } was done every morning. Now, we only do it once a week. We don't seem } to have any problems, and the photos are crisp. What are others doing? }
I check alignment every morning. Hopefully any other problems will also show up.
Greg-at-unic.sunysb.edu University Microscopy Imkaging Center S.U.N.Y. Stony Brook
} ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com -----------------------------------------------------------------------. } } Greetings All, } I'm curious about how often everyone aligns the beam on their 'scopes. I use a JEOL - JEM 100CX II. In the past, the alignment procedure was done every morning. Now, we only do it once a week. We don't seem to have any problems, and the photos are crisp. What are others doing? } } Sharron G. Chism HT (ASCP) Electron Microscopy Lab Harris Methodist Hospital Fort Worth, Texas
If you have a multi-user environment with users of different experience levels, every user should perform the quick alignment procedures. When there is trouble and the machine can't be aligned with the simple procedures, then the manager or service engineer of the TEM should perform a complete one (which includes the mechanical alignment). In addition, the microscope should be left in a standard startup condition for the next user. (Aperture out, low mag, beam spread to uniformly light the screen, text on negative info screen erased, and the plate number and info changed to some default, e.g. 0001.)
I have recently used 10 micron frozen sections of muscle fibers for immunolabelling using an antibody produced in our lab that is specific for a protein on the myosin filaments. Visualization was achieved by a flourescent secondary antibody. The problem is that we are not able to obtain the resolution necessary with the flourescent antibody in order to access fiber to fiber variation in labelling within the sarcomere or between sarcomeres.
I am thinking of using a biotinylated secondary antibody such that a biotin/avidin horseradish peroxidase visualization system can be used. This would allow us to continue this study at the light microscope level. Eventually EM wil be utilized, but light microscopy will allow for broader asessment of this large muscle. Does anyone know of a dehydration and counterstaining protocol that can be used for final viewing of the muscle tissue?
As some of you may know, Alwyn Eades is leaving the position as Director of the Center for Microanalysis of Materials in the MRL for a faculty position at Lehigh University. We are very sorry to lose Alwyn, but he would like to affect a career change at this time and we wish him the best of luck.
We will be instituting a search for a new Director of the CMM. At this time, I am writing to you to inform you of this search and to ask that you notify any colleagues you feel might be qualified to apply. A description of the position is attached below. Please feel free to distribute it.
Alwyn has been an excellent Director of the CMM and to replace him we will need all the help we can get. I do hope you can help us.
Thank you in advance.
Howard Birnbaum
********************************************
PRINCIPAL RESEARCH SCIENTIST (DIRECTOR OF THE CENTER FOR MICROANALYSIS OF MATERIALS) FREDERICK SEITZ MATERIALS RESEARCH LABORATORY UNIVERSITY OF ILLINOIS AT URBANA-CHAMPAIGN
The Frederick Seitz Materials Research Laboratory (MRL) at the University of Illinois at Urbana-Champaign is seeking a Director for its Center for Microanalysis of Materials (CMM). The CMM is a major analytic instrumentation center encompassing the techniques of electron microscopy (TEM, STEM, LEEM and SEM), microchemical and surface analysis (SIMS, Scanning Auger, XPS, UPS), ion beam methods (RBS, channeling, PIXE), probe microscopies (STM and AFM), and x-ray methods. It functions to support the research activities of faculty, Research Associates and students at the University of Illinois and at other universities, and for researchers at Federal Laboratories and in industry. The present staff of the CMM consists of thirteen professional analysts who are expert in the various analytic methods.
The Director of the CMM reports to the Director of the MRL and the CMM is supported by the MRL as one of a number of instrumentation centers. The CMM Director works with the Director of the MRL in planning the development of the CMM, in the development of new analytic methods, in the purchase of new and replacement instruments, and in making the CMM increasingly important in the research endeavor of the MRL. He/she is responsible for the management of the CMM staff, for providing scientific and technical expertise to the CMM, and with the CMM staff members, to the users of the CMM. The Director coordinates staff development and outreach activities to researchers at the University of Illinois and nationwide. She/he represents the MRL at appropriate national activities involving instrumentation facilities. The person selected for the position of Director is expected to carry out an appropriate research program (for which support is provided) and to encourage appropriate research activities on the part of the CMM staff.
Candidates should have a Ph.D. in Physics, Chemistry or Materials Science and have an established research record in an appropriate field involving the use of modern analytical methods. The candidates should have strong technical expertise in at least one of the areas of analysis covered by the CMM and should be capable of providing technical leadership in all of the analytic areas. He/she should have an established record of management of scientific personnel and the ability to develop and manage budgets.
The salary for this position will be commensurate with the experience of the candidate chosen and full benefits of the University of Illinois will apply. Please submit applications to: Howard Birnbaum, Director; c/o Ms Donna Jacobs; Frederick Seitz Materials Research Laboratory; University of Illinois at Urbana-Champaign; 104 South Goodwin Avenue; Urbana, IL 61801. The application material should include a resume', a statement of your views on the operation of a microanalysis facility within a large university research establishment such as the MRL, the names and contact information for five references who are familiar with aspects of your career important to your qualifications for the position. Applications received by November 1, 1997 will be fully considered but acceptance of applications and screening of applicants will continue until the position is filled.
The University of Illinois is an equal opportunity / affirmative action employer,
Howard K. Birnbaum (217) 333-1370 Materials Research Laboratory FAX (217) 244-2278 University of Illinois e mail: birnbaum-at-uimrl7.mrl.uiuc.edu Urbana, IL 61801
} I'm curious about how often everyone aligns the beam on their 'scopes. } I use a JEOL - JEM 100CX II. In the past, the alignment procedure } was done every morning. Now, we only do it once a week. We don't seem } to have any problems, and the photos are crisp. What are others doing? } Dear Sharron, We go through an extensive procedure daily. In addition for dif- fraction we also check the condenser aperture position and do further align- ment in selected area and diffraction modes. About once a month we check the mechanical alignment of the objective upper pole piece, and once a year we disassemble and clean the entire lens column and do a complete mechanical alignment. I don't think the normal user--doing imaging of thick biological specimens at ~10k mag--would notice the difference in pictures if we didn't do the alignment as often, but there are uses for which it is critical, and it tells us a lot about the machine. Since we do all the maintenance--no service contract for the HVEM--the information we get from doing frequent alignments is quite useful. Yours, Bill Tivol
} I'm curious about how often everyone aligns the beam on their 'scopes. } I use a JEOL - JEM 100CX II. In the past, the alignment procedure } was done every morning. Now, we only do it once a week. We don't seem } to have any problems, and the photos are crisp. What are others doing? } Dear Sharron, We go through an extensive procedure daily. In addition for dif- fraction we also check the condenser aperture position and do further align- ment in selected area and diffraction modes. About once a month we check the mechanical alignment of the objective upper pole piece, and once a year we disassemble and clean the entire lens column and do a complete mechanical alignment. I don't think the normal user--doing imaging of thick biological specimens at ~10k mag--would notice the difference in pictures if we didn't do the alignment as often, but there are uses for which it is critical, and it tells us a lot about the machine. Since we do all the maintenance--no service contract for the HVEM--the information we get from doing frequent alignments is quite useful. Yours, Bill Tivol
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Greg wrote: } } Thanks to all who responded to my Hummer Sputter target question. Many } of you have an adress for Anatech that is old. } The new address is: } Anatech LTD. } 6621-F Electronic Dr. } Springfield, Va 22151-4303 } 1-800-752-7629, Fax 703-941-8077 } } Greg Rudomen } S.U.N.Y. at Stony Brook } University Microscopy Imaging Center } greg-at-umic.sunysb.edu
I generally do at least some cursory alignment procedures each time I sit at the microscope. Most of the time all is well, but sometimes the last person had it in a different mode and/or did not bring it back to a "standard state". However, like stretching before excercising, I also find that running through the alignment procedures gets me in the proper state of mind for doing microscopy (as well as allowing my eyes to get dark-adjusted). Therefore, I would run through it just for that purpose. Is that kind of a Zen thing? :-)
Cheers,
John Vetrano _______________________________________________________________________________
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Greetings All, I'm curious about how often everyone aligns the beam on their 'scopes. I use a JEOL - JEM 100CX II. In the past, the alignment procedure was done every morning. Now, we only do it once a week. We don't seem to have any problems, and the photos are crisp. What are others doing?
Sharron G. Chism HT (ASCP) Electron Microscopy Lab Harris Methodist Hospital Fort Worth, Texas
I started life as an EM service engineer in 1966, since then I think I ha= ve handled almost every type of commercial TEM and SEM, therefore I have bee= n able to take a good look at the subject of alignment.
How often you need to align a TEM depends very much on its stability, =
thermal and mechanical. If you leave the lenses switched on at a controlled temperature, about 2 deg centigrade below room temperature is ideal, there is no reason for constantly aligning the lenses. =
If the machine has mechanical lens alignment it is my experience that the=
more you move them the more they move, compromise! However you must do a=
quick check prior to use of 1. gun alignment - viewing the spot and halo= : 2. illumination - in relation to the screen centre: 3. condenser apertur= e - in relation to the screen when C2 is spread overfocus: 4. objective aperture - in relation to the diffraction spot.
Of course the alignments that you need to perform should relate to the levels that you expect to reach when using the instrument. Less than 10,000X then you can get away with a very quick check of saturation. =
Should your target be 500,000X then the most important alignment will be voltage alignment, as clearly resolution will be your goal. In this case=
forget using the instrument within 2 hours of switching on the high voltage; that is unless you have a gas filled HT tank. It takes this tim= e for the high voltage to stabilise due to heat gained in the tank being required to reach the same level as heat lost. Gas filled tanks seem to take about 45 minutes at 120kV!
Most people over align their instruments as if the feat of completing the=
alignment is part of their religion! =
The time when an alignment becomes critical is when the lens in question = or the high voltage become unstable. To mis align the lenses is a standard engineer trick to isolate a fault to a particular part of the instrument.= =
See my book "Maintaining & Monitoring the Transmission electron Microscop= e" published by the Royal Microscopical Society ISBN 0-19-856407-4. This bo= ok also outlines all the TEM alignment procedures, including deflection coil= s and stigmators as well as covering typical image defects due to alignment=
and instability problems.
In spite of what some manufacturers may claim the alignment of a TEM follows basic procedures which have not changed since the 1960s, how can they, they are just electron optics!
Greg wrote: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } Thanks to all who responded to my Hummer Sputter target question. Many } of you have an adress for Anatech that is old. } The new address is: } Anatech LTD. } 6621-F Electronic Dr. } Springfield, Va 22151-4303 } 1-800-752-7629, Fax 703-941-8077 } } Greg Rudomen } S.U.N.Y. at Stony Brook } University Microscopy Imaging Center } greg-at-umic.sunysb.edu
Web address of Anatech LTD is:
http://www.anatechltd.com
-- Henrik Kaker SEM-EDS Laboratory, Metal Ravne d.o.o. Koroska c.14, 2390 Ravne, Slovenia Tel: +386-602-21-131, Fax: +386-602-20-436 SEM-EDS Laboratory Web Site http://www2.arnes.si/guest/sgszmera1/index.html Microscopy Vendors Database http://www.kaker.com/mvd/vendors.html Kaker.Com http://www.kaker.com
In reference to Cynthia Bennett's and Robert H. Olley's information request, there are in fact commercially available multi-process radio frequency etching systems at reasonable costs. For a full range of etch capability, it is necessary to have multi-gas inputs for user determined ratios of more than one species and the versatility to select more than one etching gas. Various gases can be used for selective etch of specific materials as has been done in the semiconductor industry since the 1960's. There are a couple reference texts available with specific designs of etching systems including parallel plate, ECR, inductivly coupled plasma and microwave technology. "Thin Film Processes" by Vossen, version I (1978) and II is an excellent reference to these processes. Also "Glow Discharge Processes" by Chapman (1980) covers these techniques in detail. As indicated in other responses, oxygen is most common for etching or removing hydrocarbons, organics and polymers such as photoresist. The O2 gas provides a chemical etch and in some cases it is desired to speed up the process by adding a heavier, non-reactive atom such as argon. This will cause a physical etch as well as a reactive chemical etch process.
To avoid "advertising" on this network, please refer to the South Bay Technology home page for additional information on one commercial versatile RF plasma etching system or contact me direct for details on the unit.
Regards, Steve
Steve Collins scollins-at-southbaytech.com Ph: 703-486-7999 (east coast USA) 714-492-2600 (west coast USA) 800-728-2233 (Toll Free) Fax: 714-492-1499 Web Page: http://www.southbaytech.com
At 10:10 AM 7/28/97 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Column alignment of TEMs is mostly for convenience and does not much affect resolution.
Filament traverse and tilt are so a maximal amount of beam enters the condenser system
Condenser alignment is so the illumination stays on the screen when condenser lens focal length is changed. The condenser/gun system is aligned with the axis of the objective so the illumination stays on the screen as objective focal length is changed.
Imaging lenses have their axes put on the line joining the centre of the objective with the centre of the screen so the image stays central and in view when magnification is altered.
The objective aperture MUST be centred on the axis of the objective. We use the zero order spot in the diffration pattern to define the axis.
For best resolution the entire illumination system should be tilted so its axis coincides with either the current or voltage centre of the objective. Voltage centre has an advantage as it aligns with the mean axes of all the imaging lenses, not just the objective. But there is not much in it. So when you are shooting for really good resolution, centre the objective aperture, correct astigmatism using background phase speckle on your specimen, use high voltage modulation to test illumination tilt and adjust for minimal image wobble, and shoot.
Maybe once a week I check condenser aperture alignment, every operator must check objective aperture alignment, gun alignment is checked after filament change of voltage change. The rest doesn't matter. We have not done a full alignment on our Hitachi H-7000 in 7 years. Mel Dickson President, Australian Society for Electron Microscopy Director, Electron Microscope Unit, University of New South Wales. Sydney NSW 2052 Australia
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Hello All:
In reference to Cynthia Bennett's and Robert H. Olley's information request, there are in fact commercially available multi-process radio frequency etching systems at reasonable costs. For a full range of etch capability, it is necessary to have multi-gas inputs for user determined ratios of more than one species and the versatility to select more than one etching gas. Various gases can be used for selective etch of specific materials as has been done in the semiconductor industry since the 1960's. There are a couple reference texts available with specific designs of etching systems including parallel plate, ECR, inductivly coupled plasma and microwave technology. "Thin Film Processes" by Vossen, version I (1978) and II is an excellent reference to these processes. Also "Glow Discharge Processes" by Chapman (1980) covers these techniques in detail. As indicated in other responses, oxygen is most common for etching or removing hydrocarbons, organics and polymers such as photoresist. The O2 gas provides a chemical etch and in some cases it is desired to speed up the process by adding a heavier, non-reactive atom such as argon. This will cause a physical etch as well as a reactive chemical etch process.
To avoid "advertising" on this network, please refer to the South Bay Technology home page for additional information on one commercial versatile RF plasma etching system or contact me direct for details on the unit.
Regards, Steve
Steve Collins scollins-at-southbaytech.com Ph: 703-486-7999 (east coast USA) 714-492-2600 (west coast USA) 800-728-2233 (Toll Free) Fax: 714-492-1499 Web Page: http://www.southbaytech.com
I also have a 100CX. I align the microscope everytime I sit down to it. If it is well aligned already the whole time takes just a few minutes out of my day. It's just a good habit to get into particularly if you have a multiuser environment.
And please pardon the bandwidth if anybody feels it's inappropriate. = This is the easiest way to widely disperse the message.
I have a substantial supply of Epon 812 (the real thing) that I find I = must part with. If anyone is interested in retro-embedding their = specimens, please contact me privately.
If you have access to a microscope with DIC (differential interference contrast) You can visualize the muscle very nicely and see the peroxidase product. You can also counterstain with hematoxylin in get more reference points.
Bob Morphology Core
On Mon, 28 Jul 1997, Lisa Brown wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } I have recently used 10 micron frozen sections of muscle fibers for } immunolabelling using an antibody produced in our lab that is specific for } a protein on the myosin filaments. Visualization was achieved by a } flourescent secondary antibody. The problem is that we are not able to } obtain the resolution necessary with the flourescent antibody in order to } access fiber to fiber variation in labelling within the sarcomere or } between sarcomeres. } } I am thinking of using a biotinylated secondary antibody such that a } biotin/avidin horseradish peroxidase visualization system can be used. This } would allow us to continue this study at the light microscope level. } Eventually EM wil be utilized, but light microscopy will allow for broader } asessment of this large muscle. Does anyone know of a dehydration and } counterstaining protocol that can be used for final viewing of the muscle } tissue? } } }
Hi folks We are trying to find the most economic source for FUji Pictrography supplies, I would welcome info on where current users buy their materials and how much they pay for the various components. I will compile the results and send back to anyone interested. If vendors wish to add to this cost survey please contact me directly Tx
-- Simon C. Watkins Ph.D. Associate Professor Director CBI University of Pittsburgh Pittsburgh PA 15261 tel:412-648-3051 Fax:412-648-2004 URL:http://sbic6.sbic.pitt.edu
We are in a tight spot. We have collected data with a monoclonal antibody (Snap-25, reactive with fusion proteins from COS cells - the immunogen was crude synaptic immunoprecipitate from humans) which was raised in mouse. We used rats for data collection. Our secondary was FITC-conjugated AFfiniPure Goat Anti-mouse IgG. Now we find that we cannot obtain a peptide for control purposes. And, if we omit the primary, and use only the above secondary, we get a definite immuno response: It looks exactly as though the primary had been applied. The manufacturer of the FITC states that the anti-mouse antibody may cross-react with other species. Now What? We cannot purchase polyclonal Snap-25. Does anyone have any ideas? Did we make a huge mistake in not testing for control situations at the outset? We are relatively new at this game - do others get down these blind alleys, singing and dancing all the way until the lights go out? Bye, Hildy
Thanks to all who responded to my question on the frequency of beam alignment. It was interesting to hear from all of you. My favorite response included the advice of "If it ain't broke, don't fix it!". The majority of those that answered do a quick alignment every day ... mostly because of so many pairs of hands that fiddle with the 'scope during the day. Since I am the only tech that works with this 'scope, and always return it to "square one", and only have two pathologists that operate it ... once a week alignment seems to be all it needs. Our normal operation takes us up to 14k or 20k mag and seldom higher than 60k - 80k. (The specimens are about 70nm in thickness.) Still others have said that they, too, have a CX100 and only align the beam once a week. The information has been very helpful. Thanks, again for your info!
Sharron G. Chism HT (ASCP) Electron Microscopy Lab Harris Methodist Hospital Fort Worth, Texas
Our Local Arrangements Committee is chewing over a question that on which we have differing opinions. The question is " What is the true function of the Sunday social and how does this function relate to the venue?" We have come up with a couple of options.
Option 1: Is this function serving to rekindle acquaintances between colleagues.
Option 2: Is this function serving to not only rekindle acquaintances between colleagues but also to set the tone for the coming meeting, and show case the hosting city.
For those list members that attend the meetings, and any others that would like to comment all your input will be welcome. If you want to reply directly to me, I will be delighted to post a summary after comments have been received.
Bob Kayton, Ph.D. Histology/E.M. Core Director C.R.O.E.T. Oregon Health Sciences University Portland, OR 97201 W-503-494-2504 Fax-503-494-6831 H-503-590-7801
Jackson Immunoresearch carries secondary antibodies that are pre-absorbed against several different animal seras including rat. We routinely do double label ICC with mouse and rat monoclonals and purchase our secodary antibodies from them. See Braisted et al. Development 120:2409-2419 (1994). There is no cross reactivity seen with the mouse secondary to the rat immunoglobins. These preabsorbed antibodies should work well on your mouse tissue. Other companies carry pre-absorbed secondaries also. We have been using Jackson for years and have been very satisfied with them. Linda Barthel Research Associate II Department of Anatomy and Cell Biology University of Michigan lab (313) 764-7476 fax (313) 763-1166 barthel-at-umich.edu
On Tue, 29 Jul 1997, HILDEGARD CROWLEY wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } Dear Immuno LM Friends, } } We are in a tight spot. We have collected data with a monoclonal } antibody (Snap-25, reactive with fusion proteins from COS cells - the } immunogen was crude synaptic immunoprecipitate from humans) which was } raised in mouse. We used rats for data } collection. Our secondary was FITC-conjugated AFfiniPure Goat Anti-mouse } IgG. Now we find that we cannot obtain a peptide for control purposes. } And, if we omit the primary, and use only the above secondary, we get a } definite immuno response: It looks exactly as though the primary had been } applied. The manufacturer of the FITC } states that the anti-mouse antibody may cross-react with other species. } Now What? } We cannot purchase polyclonal Snap-25. Does anyone have any ideas? Did } we make a huge mistake in not testing for control situations at the outset? } We are relatively new at this game - do others get down these blind } alleys, singing and dancing all the way until the lights go out? } Bye, } Hildy }
In my opinion concerning 100C(X) alignment, don't make knots to your neurons! 100 cx is a very complete but simple microscope to use.Only 6 lenses and 4 alignments coils including condenser and objective stigmators. All I'm describing is for routine work, for High resolution job it's an other problem. The main mechanical alignment is intermediate and projector lens and must be perform once or two times a year. alignment of condenser aperture is OK until people change the aperture. Gun tilt and shift are generally OK until you change the filament. The only thing to check is Spot size 1 and center with Gun shift then spot size 3 and center with Trans and repeat (once or two time) until the position of beam is the same between spot 1 and 3. If you have turn a lot the gun shift, you can check the maximunm of brightness or the image filament with gun tilt. If the precedent user has made dark field you can check the voltage center with HV wobbler and tilt knob. That's all, no more than a couple of minutes. Of course centering of objective aperture and setting of objective stig must be perform by user but it's not alignment it's just like to focus the image to obtain the best you can have. Hope that helps. ========================================================== Jacky Larnould mailto:larnould-at-mnet.fr voice:33 (0)4 67 72 28 26 fax :33 (0)4 67 79 54 90
I am looking for information about the ISO 9001 standard with regard to the operation of FT-IR microscopy and SEM systems for materials analysis, and would very much appreciate communicating with list members who have practical experience in this area.
Hi all: EMSL has a Philips 400 TEM/STEM to give away to any qualified non-profit. The instrument is in good condition and is located in Greensboro, NC. Recipient is responsible for packing and moving the instrument. Interested parties should contact Ron Mahoney at EMSL 910-297-1487 for more information. Cheers, Julian
Julian P.S. Smith III Biology Winthrop University Rock Hill, SC 29733 803-323-2111 x227 (vox) 803-323-2246 (fax)
} Date: Tue, 29 Jul 1997 11:02:50 -0600 (MDT) } From: HILDEGARD CROWLEY {hcrowley-at-du.edu} } To: postmessage {Microscopy-at-sparc5.microscopy.com} } Subject: LM:Mess-Polyclonal-mono-control } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } Dear Immuno LM Friends, } } We are in a tight spot. We have collected data with a monoclonal } antibody (Snap-25, reactive with fusion proteins from COS cells - the } immunogen was crude synaptic immunoprecipitate from humans) which was } raised in mouse. We used rats for data } collection. Our secondary was FITC-conjugated AFfiniPure Goat Anti-mouse } IgG. Now we find that we cannot obtain a peptide for control purposes. } And, if we omit the primary, and use only the above secondary, we get a } definite immuno response: It looks exactly as though the primary had been } applied. The manufacturer of the FITC } states that the anti-mouse antibody may cross-react with other species. } Now What? } We cannot purchase polyclonal Snap-25. Does anyone have any ideas? Did } we make a huge mistake in not testing for control situations at the outset? } YES
We are relatively new at this game - do others get down these blind } alleys, singing and dancing all the way until the lights go out?
SUGGEST YOU TALK TO AN IMMUNOLOGIST TO HELP DESIGN YOUR IMMUNOEXPERIMENTS. THERE ARE LOTS OF REACTIVITIES THAT EVEN EXEPRIENCED MICROSCOPISTS DON'T KNOW ABOUT WHICH ARE COMMON KNOWLEDGE AMONG IMMUNOLOGY FOLKS. FORTUNATELY, I MARRIED AN IMMUNOPATHOLOGIST WITH A PH D IN IMMUNOLOGY--HE SOLVES MY REACTIVITY PROBLEMS.
} Bye, } Hildy } Rats and mice are very similar. I'm not surprised you got cross reactivity. You should always do negative controls-- in every experiment--one of which is your secondary without the primary. But this is not enough; you should have a primary control (one that is the same type as your experimental), either a preimmune serum if polyclonal or a non-reactive mono (proven non-reactive) that is the same species and same isotype.
You can buy a rat antimouse-FITC which probably won't react with rat tissue (ours didn't). Also, you can try absorbing your secondary with normal mouse serum, but you will probably lose a lot of your specific reactivity too.
Good luck, Sara
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
Anyone have any suggestions for the question below. This is out of my area...
Nestor
} Date: Tue, 29 Jul 1997 10:48:01 -0500 } To: Zaluzec-at-sparc5.microscopy.com } From: lmtuhela-at-cc.owu.edu () } Subject: Ask-A-Microscopist } } Below is the result of your feedback form. It was submitted by } (lmtuhela-at-cc.owu.edu) on Tuesday, July 29, 1997 at 10:48:00 } --------------------------------------------------------------------------- } } Email: lmtuhela-at-cc.owu.edu } Name: Laura Tuhela-Reuning } } School: Ohio Wesleyan University } } State: OH } } Zip: 43015 } } Question: A faculty member is interested in using our SEM to observe the } giant chromosome in Drosophila for an upcoming genetics class. Are there } any suggestions for preparing the samples? We have a cryo system } available as well as variable pressure but do not have a critical point } dryer. Thank you! } } --------------------------------------------------------------------------- }
We do a lot of immunocytochem on Rat tissue and Rat -derived cultured cells and have (almost) always had good reults using anti-mouse secondaries from Jackson Immunoresearch (I've no stake in this company). They have antibodies raised in Donkey that are already preadsorbed against a number of other species, including Rat. Generally they give very low non-specific binding on our specimens. Negative controls are very important. We use the "no primary" and "pre-immune (non-immune)" controls often, but prefer the "irrelevent primary" control where possible.
Greg Martin Dept. of Cell Biology and Anatomy Johns Hopkins School of Medicine
On Tue, 29 Jul 1997, HILDEGARD CROWLEY wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------. } } } Dear Immuno LM Friends, } } We are in a tight spot. We have collected data with a monoclonal } antibody (Snap-25, reactive with fusion proteins from COS cells - the } immunogen was crude synaptic immunoprecipitate from humans) which was } raised in mouse. We used rats for data } collection. Our secondary was FITC-conjugated AFfiniPure Goat Anti-mouse } IgG. Now we find that we cannot obtain a peptide for control purposes. } And, if we omit the primary, and use only the above secondary, we get a } definite immuno response: It looks exactly as though the primary had been } applied. The manufacturer of the FITC } states that the anti-mouse antibody may cross-react with other species. } Now What? } We cannot purchase polyclonal Snap-25. Does anyone have any ideas? Did } we make a huge mistake in not testing for control situations at the outset? } We are relatively new at this game - do others get down these blind } alleys, singing and dancing all the way until the lights go out? } Bye, } Hildy }
Michigan State University has an electron microscope available it is a Philips model EM201, purchased in 1972. Anyone interested contact Roger Cargill (517)355-0364 or cargill-at-pilot.msu.edu. thank you
A collegue has been offered a free Zeiss 25. It would be shared between the biology group and the materials group and the cost to them would only be shared maintenance. He is interested in its use as a materials science TEM.
I'm looking for any information about this microscope and comments from those familiar with it. There was no information on any TEMs at Zeiss' web sites. Are they still producing them? What about the Omega filter?
Michael Cinibulk UES Inc. at Wright Laboratory Wright-Patterson Air Force Base, Ohio cinibumk-at-ml.wpafb.af.mil
I need to purchase 300 mesh copper and 1000micron-slotted or hole copper grids in the 2.3 mm size. Does any vendor in the US supply these? I know that Agar does in the UK but I thought it might be faster to purchase them on this side of the pond. Can anybody out there help me? Thanks so much.
Cheers, Peggy Bisher.
NEC Research Institute 4 Independence Way Princeton, NJ 08540. Tel.: (609) 951-2629 Fax: (609) 951-2496 e-mail: peggy-at-research.nj.nec.com
(1) there is no current manufacturer of 35mm unperforated orthochromatic film;
(2) we have been offered a supply of some old stock at reasonable price, but this will not last for all that long;
(3) it would be nice if we could pressurize some manufacturer into re-doing the stuff. The Agfa Scientia appears to be the best;
(4) a gentleman from Kodak in New York sent me a roll of TX100 to try in our optical microscope / SEM. This material allows a large range of greyscales, and it works! It did particularly well for micrographs between crossed polars, where objects type A are only just off extinction and in the same field, objects type B were displaying full birefringence. And the quality was good - "brilliant grey", if such a colour exists.
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
} A collegue has been offered a free Zeiss 25. It would be shared between the } biology group and the materials group and the cost to them would only be } shared maintenance. He is interested in its use as a materials science TEM. } } I'm looking for any information about this microscope and comments from those } familiar with it. There was no information on any TEMs at Zeiss' web sites. } Are they still producing them? What about the Omega filter?
For Zeiss, see now: http://www.leo-em.co.uk/ or http://www.mwrn.com/leo/leocont.htm
Yves MANIETTE Universitat de Barcelona Serveis Cientifico Tecnics Unitat ESCA TEM Carrer Lluis Sole i Sabaris, 1-3 E-08028 BARCELONA ESPANYA
Our Lab is ISO certified and I run the (materials) SEM Lab. Since ISO seems to be oriented toward a manufacturing environment where repetitive steps are involved, I found it a "pain in the butt" for a research/failure analysis oriented SEM facility. Procedures is the "magic word" (translate that to lots of paper work). I have procedures for operating the SEM, EDS, and WDS and also for periodic calibration checks. These procedures should reflect what is necessary to operate the equipment and generate "good" data, but at the same time, be as general as possible. This is so that under unusual circumstances your analysis strategies and conditions are not too limited. Good records keeping and adherence to the procedures are closely scrutinized. Use of ASTM methods and NIST certified standards will "lubricate" the ISO approval procedure.
Good Luck! Woody White Mcdermott Technologies, Inc
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I am looking for information about the ISO 9001 standard with regard to the operation of FT-IR microscopy and SEM systems for materials analysis, and would very much appreciate communicating with list members who have practical experience in this area.
Is anyone familiar with stains and protocols for tissues embedded in Spurr's low viscosity resin? I have been using toluidine blue on one microns and need to achieve a greater differentiation among collagen fibers and keratocytes within the stroma of the human cornea.
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Clearly, generic questions about Word Processing do not } belong in } this discussion forum, so just use common sense. } } If there is any question in your mind about something you wish to } post, send } it to me first at Zaluzec-at-MSA.Microscopy.Com and I will give you my } opinion/comments. } } ----------------- } ---------------------------------------------------------- } Can I post an Announcement of a Job Opening or a Meeting? } --------------------------------------------------------- } ------------------ } } Yes that falls within the bounds of the subject of this list as long } as it } is related to Microscopy/Microanalysis. } } --------------------------------------- } ------------------------------------ } Can I post my Resume'? } ---------------------- } ----------------------------------------------------- } } No. This forum was not created for that purpose. } } ------------------------------------------------ } --------------------------- } Can I post an Advertisement? } ---------------------------- } ----------------------------------------------- } } No, that does not fit within the bounds of this discussion forum. } } This listserver is not intended to be a Sales mechanism for commerical } } organizations, but rather it is an open discussion area about } microscopy and } microanalysis problems and solutions. 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These alternative } Internet } services, are provided independently of the Listserver Operation, } which MSA } provides as a FREE service to the WorldWide Microscopy and } Microanalysis } Community. Any funds derived from the above are used to defray the } costs of } running MSA's Internet site. } } ---------------------------- } ----------------------------------------------- } I'm only interested in XYZ Microscopy is there sublist? } ------------------------------------------------------- } -------------------- } } Nope, not here. There are other listservers which deal with specific } topics, } but this one is general. At sometime in the future I may consider } creating } subtopics. But really that is what the SUBJECT line is for. Please use } a } good description for your message! I've learned alot by just } "listening" in } on discussions about topics I know nothing about. You may also find } the same } is true for you. } } ---------------- } ----------------------------------------------------------- } Can I access this list via NewsGroups on the Internet? } ------------------------------------------------------ } --------------------- } } Not any longer. For a time this Listserver automatically forwarded all } } postings to the SciTechnqiue.Microscopy Newsgroup. However, the link } to that } newsgroup has been lost at the receiving end. A link may once again be } } possible in the future, when hardware and software is upgraded. } } --------------------------------------------------------------- } ------------ } I think I need to test the Email, what do I do? } ----------------------------------------------- } ---------------------------- } } First, if you received the instructions, after submitting } instructions, then } your Email program must be working. So no further testing is really } necessary. } } DO NOT send TEST/HELLO messages to Microscopy-at-MSA.Microscopy.Com it } needlessly clutters up the subscribers mailboxes. } } Remember every time you post a message to the Listserver over 2000 } copies } are sent. This not only wastes processing time on my computer, but } gets lots } of people angry at wasted bandwidth. You should also remember some } people } have to pay $$ just to read their Email (think of those subscribers on } } CompuServe, AOL, GENIE, etc....) We have a lot of them, and this } wastes time } &/or money! } } If you really feel you must test your Email Package or think you need } help } then send a message to: } } Zaluzec-at-MSA.Microscopy.Com. } } He's sometimes a patient soul, who at times has been known to read his } Email } and occasionally answers questions. Especially, late at night. } } -------------------------------------------------------------- } ------------- } Is there a Digest Mode? } ----------------------- } ---------------------------------------------------- } } Digest Mode is not available on the present server. It is an option } which } will be added in the future. } } Digest Mode compresses all message from a single day into one long } message } which is then sent to all subscribers. It is a nice option, and I } intend to } get it running eventually. } } -------------------------- } ------------------------------------------------- } When I send a message I get (sometime many) Bad/Bounced Messages back } why? } ---- } ----------------------------------------------------------------------- } } Because this list acts as a reflector, when message delivery fails and } is } returned to the system it goes to the originator of the message. 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He's usually easy to recognize, and his } arm } twists easily. } } -------------- } ------------------------------------------------------------- } Who do I contact with problems? } ------------------------------- } -------------------------------------------- } } God-at-Garden.Eden.Earth.Com } } ------------------------- } -------------------------------------------------- } Who Runs this? } -------------- } ------------------------------------------------------------- } } This listserver is run by Nestor J. Zaluzec } (Zaluzec-at-MSA.Microscopy.Com) It } has been & continues to be run/administered/babied mostly in his spare } time } and usually late in the evening or earlyhours of the morning. } } ------------------------------------------------------------- } -------------- } Special Announcement/News } ------------------------- } -------------------------------------------------- } } At the 1995 Winter Council Meeting, the Microscopy Society of America } (MSA) } approved a proposal to support this server as well as other } telecommunications options as a FREE Service to ALL } persons/organizations } worldwide who are involved or interested in Microscopy and/or } Microanalysis. } } Thanks to this support, the listserver will be upgrading the hardware } software and services for use by Microscopist's/Microanalysts } Worldwide. } } ---------- } ----------------------------------------------------------------- } Is there an FTP and/or WWW site? } -------------------------------- } ------------------------------------------- } } Yes, both an FTP and WWW sites exist here are the addresses: } } WWW http://www.msa.microscopy.com } } FTP ftp.msa.microscopy.com (Anonymous Login Enabled) } } ********************************* } Last Updated April 15, 1997 } Nestor J. Zaluzec } Your Friendly Neighborhood SysOp. } ********************************* } End of File } *********************************
} } The Company: } } Samsung Austin in Austin, Texas wants to offer you more than a job. We want } to offer you the chance to develop a career. Becoming involved in a } fast-paced start-up operation is both exciting and challenging and we look } for employees who are energetic, flexible, and team-oriented. } Are you willing to go the extra mile? Are you comfortable with change? Are } you interested in great benefits? Are you excited about working in a start-up } environment? Are you interested in a career where you will be an integral } part of a new company? Are you comfortable making decisions that will } influence the development of our corporate culture? } If the answer to these questions is "yes," then Samsung Austin is the company } for you. } For more information visit our web site at www.sas.samsung.com } } } The Position } } We are currently looking to hire an Entry Level TEM Technician. The } requirements for this position is simply an Associate Degree in a technical } major. Experience is preferred but not necessary. The position will involve } shift work. } } If someone know of a person who would consider this position or if you know } of a 2 year college that offers courses on TEM Sample Prep please feel free to contact me. We are going to start interviewing next week so please } get your resume to me quickly. } } Resumes can be faxed or emailed to: } David A. Griffiths } Fax (512)491-1165 } Pager (512)209-4132 } e-mail DGriffiths-at-sas.samsung.com }
Thanks to all for your help! We were able to get a pattern after tweaking the specimen quality.
************************************************************************** Lucille A. Giannuzzi, Ph.D. phone: 407 823-5770 University of Central Florida fax: 407 823-0208 Dept. of Mechanical, Materials, and Aerospace Engineering PO Box 162450 4000 Central Florida Blvd. Orlando, FL 32816-2450 email: lag-at-pegasus.cc.ucf.edu **************************************************************************
Peggy Bisher wrote: ================================================= I need to purchase 300 mesh copper and 1000micron-slotted or hole copper grids in the 2.3 mm size. Does any vendor in the US supply these? I know that Agar does in the UK but I thought it might be faster to purchase them on this side of the pond. Can anybody out there help me? ================== ================================ Because of such little demand for the 2.3 mm size, in North America, they not found as readily here as they are in Europe. However, you can find them in a few mesh sizes on our website under "regular" grids, "micron" style. Other mesh sizes and types are available also but are not yet up on the website. I think that some of the other EM suppliers of consumables in the USA offer 2.3 mm grids as well, or at least they did until recently.
Chuck
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I'd like to examine a sample in my SEM. My concern is that if the beam heats the sample, it may volatile and contaminate my column. I don't have a cold finger. Does the beam heat the sample? What is the temperature of the beam? What sort of elevation in temperature does a sample go through during an examination? What about different accelerating voltages, do they produce beams with different temperatures? Thanks Mark E. Darus
Dear colleagues and calibration standard manufacturers.
A committee within the Dutch Microscopy society is working on the general issue of calibration. In June we made a request on the microscopy listserver. The only helpful item that came up was a reference for the NIST calibration standard SRM484 G (which I already own).
On the Internet there is however a large variety in calibration standard available. It's a pity that so little people have responded therefore one can only assume that they are: 1. not using the standards 2. not interested in obtaining accurate results to satisfy the ever demanding customer 3. not on the Internet 4. don't feel the need to talk to us (were told not to by there boss or were shy) 5. under the impression we're not nice persons 8(
Therefore I see no alternative then to ask you again a few questions. The questions are intended for users of a SEM and the manufacturers of calibration standards but anyone who has some interesting things to say is welcome.
Are you all aware of the writing of Herr Clive Walker on "The Report on the 4th Plenary Meeting of ISO/TC202 Londen 6-9 th May 1997" ??. In this writing there are still some issues that must be dealt with. One issue is calibration but unfortunately a consensus was not reached and the issue remains open. So, let's say I want to apply for a STER-lab certificate; what issues are to be dealt with other than calibration (other than the references given in the NIST calibration routine).
Some questions that come to my mind are: * Standard defining terms used in Micro Beam analysis. * General machine settings ? * Info about the mounting size. * Is the sample 1D or 2D (the NIST standard is a 1Dimension standard, IBSEN has a 2D standard) are they the same in the sense that they can both be used for the same calibration routine (for instance the ASTM E 766-93). * Is it important to know about the difference in Z (atom number). The NIST standard has thin gold lines in a nickel base. Others use a silicon grid. Does the difference in Z also mean that there might be a difference in signal to noise ratio when the same machine offsets are used. * Can you give us some photograph's of the calibration sample at let's say 3 different magnifications, 3 different accelerating voltages and corresponding spotsizes. * Can you give us detailed information about the accuracy of the calibration standard and a normal achievable level of accuracy with an average SEM (knowing it's hard to define a normal SEM) including the methods used for statistical proceedings eg. . * Do you have references from laboratory that have a STERlab certificate(not ISO 9000) or equal and I refer to the new ISO/IEC guide 25 using your calibration standards.
I know of the following calibration standard manufactures: Ernest F. Fullam, Inc. SPI. Energy Beam sciences Inc. NIST. MOXTEK, order at Ted Pella, Inc. TCL (I am not sure if they make standards but they do SEM calibration) IBSEN MAG*I*CAL (National Research Council of Canada)
All info is derived from the Internet using common search engines. The list is probably not complete and I would appreciate it if the list can be made up to date with more info.
It is not my intention to give a rating for the calibration standards that are provided with a large variety in spec's and cost. What I (we) want is to give people who are on the road for ISO/IEC certification some info about the calibration standards and the issues that come along in the process. Therefore I give everybody a equal opportunity to give info about his or her calibration sample making it more easy for us people to make a choice in what calibration standard the most suited is for his or her specific use.
As stated above this request goes to all the known manufacturers on the list and this request will also be posted on the Microscopy list server.
Many thanks in advance,
Gert ten Brink Philips Semiconductor BV. Fysisch analyst postbus 10 9500AA Stadskanaal tel 0599632380 fax 0599632505 e-mail brink-at-skn.sc.philips.com privat asslab-at-xs4all.nl
Ps. all cost made and found reasonable (photo material, stamps etc.) will be refunded. On the other hand: if this leads to a large rise in sales in calibration standards can I give you the number of my bankaccount ?
NIST t.a.v. Thomas E. Gills Gaithersburg, Maryland, MD 20899
Allen R. Sampson Advanced Research Systems St. Charles, Illinois ISO/IEC Guide 25-draft four Recommendations for implementation of ISO Quality Standards in an SEM laboratory Both can be found on the Internet
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Dear all, I am interested in this question, I sometimes see evidence of local heating of samples during TEM observation but I have never had much idea about quite how much heat is generated by the electron beam. Does anyone know or know how to find out?
Also, does anyone have any idea why oily blobs (obviously from oil in the vacuum system) sometimes appear on the sample in the very place that you are observing (as opposed to any other place on the sample)?
Thanks
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ Ian MacLaren, Tel: (44) (0) 121 414 3447 IRC in Materials for FAX: (44) (0) 121 414 3441 High Performance Applications, email: I.MacLaren-at-bham.ac.uk The University of Birmingham, http://web.bham.ac.uk/I.MacLaren/ Birmingham B15 2TT, England. ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
} I'd like to examine a sample in my SEM. My concern is that if the beam } heats the sample, it may volatile and contaminate my column. I don't have } a cold finger. Does the beam heat the sample? What is the temperature of } the beam? What sort of elevation in temperature does a sample go through } during an examination? What about different accelerating voltages, do they } produce beams with different temperatures? } Thanks } Mark E. Darus
The beam certainly can heat some specimens to a sufficiently high temperature to cause evaporation. The temperature reached is going to depend on all sorts of things - specimen thermal conductivity, how well specimen is connected to mount, how well mount is connected to stage, beam energy and current, scan speed (versus leaving probe stationary), probe size, etc.
Personally, unless you have got something really nasty - elemental arsenic maybe, or mercury - under normal circumstances, I would say that the volume evaporated is going to be so small that there is no need to worry. Although if you are going to be looking at a volatile specimen for 12 hrs a days, 7 days a week you might want to start thinking about a cold trap.
One thing to try to check is vacuum level - it this doesn't change when examining the potentially volatile specimen, I wouldn't get too concerned.
I work in a hospital lab and our pathologist would like some EM pictures of plts. Does anyone have a procedure for isolating plts. from whole blood and embedding them in Spurrs?
I know prices are high. Must have to do with low volume, lots of labor costs, supply and demand, profit??? Anyway... My "primary" standard is the NIST SRM-484 (last time I looked was in the $700-800 range). I also use NIST tracable sphere suspensions from Duke Scientific. Another "standard" I use for very low magnification is a section from an etched steel rule (from Starret, or equal). The rule does not come with a "pedigree", but is NIST tracable through the manufacturer and is an "industry accepted" measuring device.
Woody White Mcdermott Technology, Inc. http://www.mtiresearch.com/ http://www.geocities.com/capecanaveral/3722
Dear Woody, Do you use NIST-certified magnification standards? I found one from Geller Microanalytical which is $1500. Do you know of other suppliers, perhaps cheaper? Thanks, Melanie Behrens (going ISO as soon as I get all this paperwork done....)
The easist way to isolate platlets is to obtain a 7cc tube of blood in an ACD tube (EDTA will also work), centrifuge for 15 minutes at 100 x g. Remove the supernatant (this is the platlet rich plasma), high speed spin this to form a pellet and process as normal.
Best of Luck, Ed Calomeni Dept. Pathology Medical College of Ohio Toledo, OH 43614-2598 emlab-at-opus.mco.edu
The easist way to isolate platlets is to obtain a 7cc tube of blood in an ACD tube (EDTA will also work), centrifuge for 15 minutes at 100 x g. Remove the supernatant (this is the platlet rich plasma), high speed spin this to form a pellet and process as normal.
Best of Luck, Ed Calomeni Dept. Pathology Medical College of Ohio Toledo, OH 43614-2598 emlab-at-opus.mco.edu
Ian MacLaren wrote: } I am interested in this question, I sometimes see evidence of local } heating of samples during TEM observation but I have never had much } idea about quite how much heat is generated by the electron beam. } Does anyone know or know how to find out?
Dear Ian,
L. W. Hobbs has a contribution entitled "Radiation Effects in Analysis of Inorganic Specimens by TEM" in "Introduction to Analytical Electron Microscopy" edited by J. J. Hren, J. I. Goldstein and D. C. Joy, Plenum Press 1979. In this paper there is on page 441 a section on "Electron-Beam Heating" where you will find the relevant equations.
The sample temperature will depend on beam current, thermal conductivity and specimen geometry. Hobbs mentions that under the worst circumstances it is possible to melt refractory ceramics.
Best wishes, Joergen.
J. B. Bilde-Soerensen Senior Research Scientist, ph. d. Materials Research Department Risoe National Laboratory DK-4000 Roskilde Denmark
Dear all, A few months ago I asked my Professor about heating of samples during TEM observation and he suggested me to read the monograph of L.Reimer: Transmission Electron Microscopy (Springer-Verlag, Berlin, 1984). I have read and I can recommend you as the good source regarding "Beam Temperature".
In response to Ian MacLaren's request, hydrocarbon contamination is resident on EM specimens as a result of preparation and handling techniques, , ambient conditions, and microscope vacuum contamination, although in most cases it has been found that the microscope vacuum is actually quite clean.
Under vacuum conditions, the hydrocarbons are mobile on the specimen surface. As they pass the impingiment point of the electron beam on the specimen, they are essentially polymerized. With the beam focussed at one area on the specimen, as is the case when conducting fine probe microanalysis in a TEM, a carbon cone is generated. On a TEM specimen, the carbon formation is created simultaneously from both specimen surfaces. These carbon formations or "oily blobs" as Ian MacLaren called them preclude both imaging and the acquistion of analytical data.
It has been found that low-energy plasma cleaning of the specimen prior to EM observation, utilizing an oxygen-based process gas, chemically converts the hydrocarbon contamination to CO, CO2 and H2O. This process virtually eliminates the contamination issue without altering the specimen's properties. The resultant is enhanced imaging and analytical data.
Best regards,
Paul E. Fischione E.A. Fischione Instruments, Inc. 9003 Corporate Circle Export, PA 15632 USA Phone 412-325-5444 FAX 412-325-5443 Web site: www.fischione.com
Hi everybody According to Castaing in thesis the temperature rise at the point of irradiation theta in Celsius is=20 approximately as follow:
theta =3D 1.14 IaV/Cd
Where Ia specimen absorbed current in microamp V accelerating voltage in kV C Thermal conductivity (cal/cm sec deg) d electron probe diameter in micronmeter
for example with glass at 10kv and d=3D0.1=B5 and Ia=3D0.1nA rise of temp is about 7 degrees celsius
If somebody interresting I also have a diagram showing temperature rise vs thermal conductivity (about 50k in TIF) Hope that helps.
Eugene, I use Spurr's exclusively for E.M. The stain I use on 0.5 micron sections is Paragon. It is a combination of Toluidine Blue and Basic Fuchsin and is a really beautiful stain. The reference for this is Dr. Frieda Carson's book "Histologic Techniques In Electron Microscopy", published by the American Society of Medical Technology, 1979. Hope this helps. Sharron G. Chism HT (ASCP) Electron Microscopy Lab Harris Methodist Hospital Fort Worth, Texas
Please help me with some recommendations; We are a multi-user, University, primarily biological, microscopy facility used extensively by staff and postgraduates from the Faculties of Science and Agriculture.
For the past ten years we have made extensive use of a steam(286)-driven Kontron Vidas full colour Image Analysis system which, though competent and versatile, was soon dubbed 'user-hostile' by our students.
We now have a very limited budget with which to replace this apparatus. Any new system would need to be Windows-based (to satisfy those students !) and have a high specification of PC hardware which is easily obtained from local suppliers.
I am looking for proven recommendations for software used in similar applications to our own. Further detail can be supplied on request.
I would particularly appreciate recommendations on reasonably priced products via fellow microscopists. Our applications range over particle counting through a great variety of macro and micro area measurement to measuring the black vs white area on dairy cows !!
Commercial responses should be mailed directly to me in the spirit of correct net protocol.
Tony Bruton Head, Centre for Electron Microscopy University of Natal, Pietermaritzburg. KwaZulu-Natal, South Africa Tel: +27 331 2605155 Fax: +27 331 2605776 E-mail: bruton-at-emu.unp.ac.za
FALL 1997 COURSE ANNOUNCEMENT - Transmission Electron Microscopy (BIO. 221-V2)
NASSAU COMMUNITY COLLEGE
A fifteen week, fall 1997 semester, course in Biological Transmission Electron Microscopy is being offered by the Biology Department of Nassau Community College. This is a 4 credit course offered ONE EVENING PER WEEK, Thursdays, starting at 5:30pm. Classes will begin on Sept. 4 and end on Dec. 18, 1997.
This is a "hands-on" course emphasizing biological specimen preparation, ultra-thin sectioning involving block trimming, glass knifemaking and operation of the ultramicrotomes (Sorvall MT-2B and LKB Ultrotome III), thick and ultra-thin section mounting and contrast staining (UA and Pb citrate), grid support films (formvar, carbon), student operation of the TEM (Hitachi HS-8, Philips EM 300) and production of electron micrographs through the process of black & white photography, and electron micrograph analysis. Students will work on a chosen sample(s) with the goal of producing a portfolio of high quality TEM photomicrographs of that sample(s).
The course is widely transferrable and the cost per credit is reasonable at $84 per credit.
More information about the Bio-Imaging Center at NCC, course descriptions and syllabi, and the beginnings of a student gallery of EM photomicrographs is available at our web site. The URL is {http://www.sunynassau.edu/webpages/biology/becks.htm} .
For those without www access, the catalog description is specified below. If you have further questions, you should e-mail me directly at the address below.
Interested individuals should register early (prior to Aug. 15) since the course is limited to a total enrollment of ten (10) students.
Questions regarding the actual registration process can be directed to our registrar at (516) 572-7355. ________________________________________________________________________________
CATALOG DESCRIPTION BIO 221: Transmission Electron Microscopy -- 4 credits Prerequisites: BIO 109-110 or equivalent, CHE 151-152 or equivalent. An introduction to the basic principles of transmission electron microscopy including tissue preparation, microscope (TEM) operation, black & white photography, and micrograph interpretation. The entire laboratory is devoted to the development of skills and preparative techniques involved with the operation of an actual transmission electron microscope. (3 lecture, 3 laboratory hours). Laboratory fee applies. ________________________________________________________________________________
Stephen J. Beck Associate Professor Bio-Imaging Center/Electron Microscopy Department of Biology Nassau Community College Garden City, NY 11530 Voice Mail: (516) 572-7829 Email: {becks-at-sunynassau.edu} URL: {http://www.sunynassau.edu/webpages/biology/becks.htm}
Dear All, A student, as part of an electron microscopy project, has ferrosilicon samples from powder up to 2mm diameter. After heat treating, he wants to chemically polish the surface to remove contamination. Does anyone know of a chemical polish that would do this? Preferably it should polish not etch the surface. Thanks for any suggestions. Mike
Michael J Witcomb PhD Electron Microscope Unit University of the Witwatersrand Private Bag 3 WITS 2050 South Africa
I am doing in situ hybridization on parafin embedded and sectined plant material using a DIG labelled probe. However, I find that the binds non-specifically to walls and cytoplasm. I have tested the anti-dig AB and the do not show any non-specific binding.
Does someone have an idear on how to block non-specific binding of the probe prior to hybridization without damaging the DNA (or RNA) in the sections?
Peggy Bisher wrote: } } ------------------------------------------------------------------------} The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } -----------------------------------------------------------------------.} } Dear Newsgroup: } } I need to purchase 300 mesh copper and 1000micron-slotted or hole copper } grids in the 2.3 mm size. } Does any vendor in the US supply these? I know that Agar does in the UK but } I thought it might be faster to purchase them on this side of the pond. Can } anybody out there help me? Thanks so much. } } Cheers, Peggy Bisher. } } NEC Research Institute } 4 Independence Way } Princeton, NJ 08540. } Tel.: (609) 951-2629 } Fax: (609) 951-2496 } e-mail: peggy-at-research.nj.nec.com
Peggy, We have the following 2.3 mm copper grids in stock. If any are of interest, please e-mail me and I will get you pricing:
Dear Mark, Others have answered the other questions you asked, so I'll just tackle these two.
} What is the temperature of } the beam?
The beam is far from equilibrium, so such concepts as temperature don't really apply, but for a monatomic gas, E = 3/2kT, so for a "gas" of electrons, you can use the same equation. If you had a gas of hot electrons in a chamber with a small hole in one wall, there would be a stream of electrons emerging from the hole whose average energy would be 3/2kT. This is not exactly a mono-energetic electron electron beam, but the same concept can be applied. 1 eV is about 10^4 K, so a 10 keV beam has a "temperature" of about 10^8 K.
} What about different accelerating voltages, do they } produce beams with different temperatures?
From the considerations above, yes.
The connection between the very high temperature of the beam (about that of a stellar interior) and the heating of the specimen is through the energy deposited in the specimen as the electrons are slowed to a stop. In the SEM each electron is stopped in the specimen, so all the energy is converted to heat (except that used in the produc- tion of secondary electrons, taken away by backscattered electrons, etc.). A 10 keV electron has a range of 0.28 mg/cm^2, or--since carbon has a density of ~2 g/cm^3--~1.4 micro meter. Thus, all the energy is deposi- ted in a thin layer near the surface. As others have said, the final specimen temperature depends on how this heat is dissipated. Yours, Bill Tivol
Yes, I have a procedure for what you want. We are experimenting with the following process ...so far so good! (About 5 - 7 mls of blood is collected in an EDTA tube.) 1. Centrifuge blood sample at 1,300 rpm for 10 minutes. 2. Draw off half of the plasma and discard. 3. Centrifuge again at 1,300 rpm for 10 minutes 4. Draw off plasma leaving a 2mm layer over the cells. Be VERY careful not to disturb the cells. 5. Replace the plasma with an equal amount of 2.5% buffered glutaraldehyde. 6. Place this in the 'fridge for at least 4 hours ... overnight is ok. 7. Carefully draw off the fixative and discard. 8. Remove the button of cells with a sharp applicator stick, and place in a dissecting dish. (Try not to break the button too much ... you can probably get it out in two pieces.) 9. Rinse the cells with fixative and remove as much of the red cells as possible with a razor blade. 10. Dice the button into 1mm squares and process as you would tissue. We go through Osmium, graduated alcohols and eventually embedding in Spurr's.
The only difference in this process and our routine buffy coat is the first three steps. We usually centrifuge at 3,000 rpm for 10 minutes and skip steps #2 and #3 for leukocyte study. We have found that the faster rpm can damage platelets so we've come up with this ... see what you think.
Sharron G. Chism HT (ASCP) Electron Microscopy Lab Harris Methodist Hospital Fort Worth, Texas
I work in a hospital lab and our pathologist would like some EM pictures of plts. Does anyone have a procedure for isolating plts. from whole blood and embedding them in Spurrs?
Just a follow up to Paul F.s comment. I would disagree with Paul's statement saying..
"although in most cases it has been found that the microscope vacuum is actually quite clean".
I've done some pretty extensive work on this topic for more years than I'd like to admit to and can show that contamination is also derived from the microscope "vacuum". I've worked with microscopes operating from 10**-5 to 10**-10 torr. Cleaning a specimen with reactive gas plasma minimizes the initial specimen borne components. But if a specimen is left in even a relative modern microscope over night (~ 10**-7 to 10**-8) the contamination effects can return albeit at a reduced level. Subsequent retreatment of the specimen with a plasma will remove this but if you leave it sitting in the microscope it will eventually return.
Stop by the poster session at the Microscopy & Microanalysis 97 meeting in Cleveland and I'll be glad to fill you in.
I recently went through an extensive evaluation of several systems to replace our Kontron system. We examined systems for Mac, Unix and PC systems and determined that a software package called Optimas is the best value for the money. It is a PC based system that works well with Win95 or Win NT (or Win 3.11 for that matter!) The software has many pre defined macros which may help in your application, but also has a very powerful, vector based, C-like language which is fairly easy to use. The support is top notch with and excellent web page: http://www.optimas.com which gives really good support and their phone support is also very good. The company is located in Washington state and our local vendor sold a single license of the software for $3995. We bought our own frame grabber and computer. If you would like to contact me off line to ask further questions, my email is David_Bell-at-millipore.com and my number is (617) 533-2108.
I am in no way connected to Optimas or any Optimas reseller, I am just a very satisfied user.
We are preparing to examine dairy products for protein, fat and starch. If anyone has had experience with this application using a Biorad krypton-argon laser and could give any suggestion on technique and suggest the appropriate flourochromes it would be greatly appreticated.
William R. McManus Electron Microscopy Facility Department of Biology Utah State University Logan UT 84322-5305 1-801-797-1920
Some principles I think are important. Stay away from systems that require proprietary hardware boards for the software to run. These systems tend to become obsolete quickly or are expensive to upgrade. The less expensive image analysis software programs tend to be easy to use but lack the flexibility and power when confronted with a difficult problem. For overall cost and performance, I think PC based systems are the best. My lab has chosen Optimas software (runs under Win95 or NT) as the main image analysis platform, and sofar it has been able to do everthing we require.
Regards,
John J. Turek, Ph.D. Associate Professor Director, Electron Microscopy Laboratory and Core Laboratory for Image Analysis and Multidimensional Applications (CRISTAL) Department of Basic Medical Sciences 1246 Lynn Hall, G193C Purdue University W. Lafayette, IN 47907-1246 Phone: 765-494-5854 Fax: 765-494-0781 Email: jjt-at-vet.purdue.edu
On Thu, 31 Jul 1997 David_Bell-at-Millipore.com wrote:
} I recently went through an extensive evaluation of several systems to } replace our Kontron system. } We examined systems for Mac, Unix and PC systems and determined that a } software package called Optimas is the best value for the money. It is a } PC based system that works well with Win95 or Win NT (or Win 3.11 for that } matter!) The software has many pre defined macros which may help in your } application, but also has a very powerful, vector based, C-like language } which is fairly easy to use.
We did the same and decided on Visilog in part because it uses real C-code and includes a C-interpreter which aids programming. All the other stuff too but the price is a bit more.
Talking about long term storage of tissue reminds of one of my favorite sayings - I always reserve the right to be wrong! - (But I will do my best).
Unfortunately I have not real hard data on tissue storage. This would take an immense amount of time to accumulate. But over many, many years, the following have proven to be good. After glutaraldehyde the tissue remanins quite permeable. Fluids and material may flow in both directions*. Some lipid has been lost*. This is evident when mycobacterium a prefixed with glut and malachite green, and lipid containing filaments which perhaps carry antigen, remain visible. The malachite prevents lipid loss at the outset*. Immediate, superfast, fixation and processing of tissue results in the most "brilliant" of sections and intact cytoplasm. This comes into evidence when one does pathological tissue TEM investigation - wet tissue to paper micrographs in 5.5 hours. I have done this, and the results are astonishing. But we cannot indulge in this. Frequently we must store for weeks, months. This was the case when I worked at a research institution which had to store lung tissue. We stored it in 0.1M buffer after fixation for 4 hours in glut. And we stored in in 0.2M buffer on the assumption (Note: assumption is the mother of all screw-ups) that the increased hypertonicity of the buffer would prevent exodus through membranes. We felt that this improved storage conditions, but we did not do a systematic study, or spend much time at very high TEM magnifications. But we did end up adopting that strategy. Osmium fixes lipids and some proteins*. At a later date I started walking tissue through the osmium step and then storing for long term in buffer. I felt that this was quite an improvement. I have no comparative data. If asked how I would store tissue today, my answer would be to carefully fix it with phosphate buffer and fresh glut, quick rinse it, and refix it in osmium, constantly keeping the tissue in motion, and store it in phosphate buffer in the refrigerator, tightly capped. Storage in alcohol is not advisable, as the alcohol can dislodge the osmium. We see this when the alcohol turns brownish. Long term storage in glut brings up the question of the polymerization and degredation of glut over time*. What effect would this have on the tissue? To really aquire an answer one would have to do a "blind study". That is, the microscopist would have to be required to take the micrographs and then sort them according to their storage conditions without the benefit of any ID data sheet. *Denotes references available for these statements. Bye, Hope to see you all in Cleveland, Hildy
If you worry about cooking a specimen the following guide lines may be of=
assistance.
1. Use a low kV, the lowest you can use with comfort 2 to 5 is possible with most conventional instruments although I have looked at uncoated photoresist at 200v with Lab6. FEG makes the job too easy. 2. Use a low emission current about 20- 50uA with W 3. Use a small spot size, not for resolution but as a safety device.=
4. Set the stage so that when you switch on the beam you are NOT on the specimen 5. Set up off the specimen material and when on the material make yo= ur adjustments about a screen width away from the area required. 6. Only move the beam at the last moment, do not move the stage as this may change the Z .You need a fairly flat specimen or a good depth of=
field to be successful here. 7. Practice focussing and stigmating then press photo and then move the specimen the known amount with a X or Y beam shift. Most instruments=
pause between the photo button press and actually starting the scan. 8. In my experience some specimens will be damaged or contaminated s= o they look different after even one additional scan. This one scan photo method is about a pure an image as you can get.
1 to 3 above are the safety features, cut down the kV and cut down the be= am current and you save your specimen. Keep the kV off and they last even longer :-)
I visit on average one SEM laboratory per week throughout the year and it=
was initially quite a shock to see that very very few really know how good/bad their microscope calibration is!
Coming from the direction where as a TEM engineer I calibrated all of the=
microscopes that I attended once each year, in my teaching I carry this practise over to the SEM. I routinely carry out SEM resolution, magnification calibration and contamination rate tests on the instruments=
that I use. At first I tried drift rate tests too but the results came a= s a shock!
Resolution - most instruments are set up incorrectly. The electron gun i= s always in economy mode i.e. the filament is too far from the cathode to enable spec resolution to be attained. Correct this problem or tune the gun further and it is good to see how many old instruments are capable of=
beating their spec resolution. I use my well know sputtered gold on late= x spheres for this test.
Magnification Calibration - Most instruments are within the standard I fe= el is respectable which is plus or minus 10% of the readout with no more tha= n a 5% error between X and Y directions. What people fail to recognise is that different spot sizes on the same area at the same magnification provide different calibration values. Typical is a ten turn potentiomete= r on old Hitachi instruments 2 turns give a 5% change in calibration. Peopl= e do not seem to recognise that if you change the focus after some other adjustment you have just changed the effective working distance and therefore the magnification. I use an Agar TEM carbon grating replica a cross grating of 2160 line per millimetre. I prefer this specimen as it also makes a very good demonstration specimen on the effect of kV on imag= e form. See "Working With A SEM" S.K. Chapman ISBN 0 850770 93 9. It is m= y experience that on some SEM the magnification calibration is very good at=
certain kV, but bad on others. Machines also seem good at certain WD but=
not at others. In courses each student measures each picture and we have= a spread of 4 to 7% amongst them! It is not that easy to calibrate a SEM!
Contamination Rate - all this comment about oil and specimen damage, is n= ot contamination a cracking of vapours within the vacuum by the heat of the beam on a surface. Is it not hydrocarbons and silicons being deposited hence the low signal level dark lines or rectangles? SEM contamination rate is very much specimen dependant but by taking a constant approach th= is may be a useful test. I use sputter coated latex spheres the specimen being in the microscope one hour prior to the test. A typical rough toug= h microscope used without any care gives 10nm/min over my 20 minute test period. Under similar (emphasise similar) conditions a well kept air locked instrument will come down to 2.5nm/min. Add a cold finger around the final lens similar to that used in a cryo system and you are down to {1.5nm/min.
Drift Rate - I thought SEM stages were lousy however testing a good numbe= r of instruments over a wide price range I found that over a twenty minute period the amount of drift was less than the instruments resolution, in other words the sample did not move. If it did I always found an earth problem not a stage drift problem. I no longer bother with this test unless I have a worry about a particular stage stability. =
Most of my work has been on run of the mill instruments with the best results from the modern twin detector FEG systems. In these instruments = a good cold finger sitting around the final lens is the difference between good and amazing results - contamination IS the killer of high resolution=
If you worry about cooking a specimen the following guide lines may be of=
assistance.
1. Use a low kV, the lowest you can use with comfort 2 to 5 is possible with most conventional instruments although I have looked at uncoated photoresist at 200v with Lab6. FEG makes the job too easy. 2. Use a low emission current about 20- 50uA with W 3. Use a small spot size, not for resolution but as a safety device.=
4. Set the stage so that when you switch on the beam you are NOT on the specimen 5. Set up off the specimen material and when on the material make yo= ur adjustments about a screen width away from the area required. 6. Only move the beam at the last moment, do not move the stage as this may change the Z .You need a fairly flat specimen or a good depth of=
field to be successful here. 7. Practice focussing and stigmating then press photo and then move the specimen the known amount with a X or Y beam shift. Most instruments=
pause between the photo button press and actually starting the scan. 8. In my experience some specimens will be damaged or contaminated s= o they look different after even one additional scan. This one scan photo method is about a pure an image as you can get.
1 to 3 above are the safety features, cut down the kV and cut down the be= am current and you save your specimen. Keep the kV off and they last even longer :-)
I visit on average one SEM laboratory per week throughout the year and it=
was initially quite a shock to see that very very few really know how good/bad their microscope calibration is!
Coming from the direction where as a TEM engineer I calibrated all of the=
microscopes that I attended once each year, in my teaching I carry this practise over to the SEM. I routinely carry out SEM resolution, magnification calibration and contamination rate tests on the instruments=
that I use. At first I tried drift rate tests too but the results came a= s a shock!
Resolution - most instruments are set up incorrectly. The electron gun i= s always in economy mode i.e. the filament is too far from the cathode to enable spec resolution to be attained. Correct this problem or tune the gun further and it is good to see how many old instruments are capable of=
beating their spec resolution. I use my well know sputtered gold on late= x spheres for this test.
Magnification Calibration - Most instruments are within the standard I fe= el is respectable which is plus or minus 10% of the readout with no more tha= n a 5% error between X and Y directions. What people fail to recognise is that different spot sizes on the same area at the same magnification provide different calibration values. Typical is a ten turn potentiomete= r on old Hitachi instruments 2 turns give a 5% change in calibration. Peopl= e do not seem to recognise that if you change the focus after some other adjustment you have just changed the effective working distance and therefore the magnification. I use an Agar TEM carbon grating replica a cross grating of 2160 line per millimetre. I prefer this specimen as it also makes a very good demonstration specimen on the effect of kV on imag= e form. See "Working With A SEM" S.K. Chapman ISBN 0 850770 93 9. It is m= y experience that on some SEM the magnification calibration is very good at=
certain kV, but bad on others. Machines also seem good at certain WD but=
not at others. In courses each student measures each picture and we have= a spread of 4 to 7% amongst them! It is not that easy to calibrate a SEM!
Contamination Rate - all this comment about oil and specimen damage, is n= ot contamination a cracking of vapours within the vacuum by the heat of the beam on a surface. Is it not hydrocarbons and silicons being deposited hence the low signal level dark lines or rectangles? SEM contamination rate is very much specimen dependant but by taking a constant approach th= is may be a useful test. I use sputter coated latex spheres the specimen being in the microscope one hour prior to the test. A typical rough toug= h microscope used without any care gives 10nm/min over my 20 minute test period. Under similar (emphasise similar) conditions a well kept air locked instrument will come down to 2.5nm/min. Add a cold finger around the final lens similar to that used in a cryo system and you are down to {1.5nm/min.
Drift Rate - I thought SEM stages were lousy however testing a good numbe= r of instruments over a wide price range I found that over a twenty minute period the amount of drift was less than the instruments resolution, in other words the sample did not move. If it did I always found an earth problem not a stage drift problem. I no longer bother with this test unless I have a worry about a particular stage stability. =
Most of my work has been on run of the mill instruments with the best results from the modern twin detector FEG systems. In these instruments = a good cold finger sitting around the final lens is the difference between good and amazing results - contamination IS the killer of high resolution=
Greetings, I inherited a Lietz Orthomat Camera system. It has the controler and the camera part. Alas, it says on the box that it is broken and the estimate for repair made in 1986 was $1000 US dollars. This item is not quite old enough to be a "collectors item" but when these function, they are very very good. I would hate to throw this away. I was just wondering if there were perhaps someone who could use the parts? Or who might be able to fix the camera and use it? Thanks, Tobias
We saw the same thing in our XPS system operating in the low 10E-9 to high 10E-10 Torr range when we did our Contamination study that we presented at the Spring MRS97 meeting. We wanted to measure very small quantities of HC's on the surface and wanted to know if the vacuum was contributing. (We were trying to find our minimum detectability limit.) We left the sample overnight after sputter cleaning to a fresh, i.e. no C peaks, and ran the XPS first thing in the morning. A significant surface C peak was present. Of course, any good RGA will tell you how much HC's you have in a vacuum system. -Scott Walck
} } I've done some pretty extensive work on this topic for more years } than I'd like to admit to and can show that } contamination is also derived from the microscope "vacuum". I've worked } with microscopes operating from 10**-5 to 10**-10 torr. Cleaning a } specimen with reactive gas plasma minimizes the initial specimen borne } components. But } if a specimen is left in even a relative modern microscope } over night (~ 10**-7 to 10**-8) the contamination effects can return albeit at } a reduced level. Subsequent retreatment of the specimen with } a plasma will remove this but if you leave it sitting in } the microscope it will eventually return. } } } Stop by the poster session at the Microscopy & Microanalysis 97 } meeting in Cleveland and I'll be glad to fill you in. } } } Nestor Zaluzec } } Your Friendly Neighborhood SysOp. } }
The question of thermal effect caused by electron beam in SEM and X-Ray microanalysis was a subject of studies by Dr. M.N. Filippov in his Doctor of Science Dissertation devoted to the microprobe analysis of unstable samples.
It was found that the theoretical estimation of the overheating of the sample may be expressed as DT = 7.8* Io*Eo*ro/lambda/(ro*do+0.13*Eo^1.7), where Io is the probe current (MickroAm), Eo - electron energy (keV), ro - sample density (g/.sm^3), lambda is the heat conductivity of the sample(Wt/cm/K). DT -s the sample overheating (in K). In his works Dr. filippov also derived the equation to estimate the time, required for the achieving the overheating temperature
t = 2.5E-7*(c*ro/lambda)*(0.5d*do+6.4E-2*(Eo^1.7)/ro)^2. Here t is time in sec, c is the specific heat capasity (J/g/K).
As a consequense of this equations, it was found (and cofirmed experimentally) that the for a lot of samples which suffer from the electron beam induced overheating, the maximum overheating is at about 25-30 kV. If you increase the beam energy, despite the fact that you are starting to pump more energy to the sample, you are also increasing the dissipation surface, and thus, the overheating of the sample often at 50 kV is smaller than the one at 30 kV.
More detailed information might be obtained from Dr. Filippov. (As far as I know his E-Mail is fil-at-pel157a.phys.msu.su Hopefully, this may be usefull.
E-Mail : _ . Nick Kinaev ,~' (_|\Centre for Electron Microscopy(CMM) nick-at-mama.minmet.uq.oz.au ,-' \ The University of Queensland Ph. home : +61 7 3279 4771 ( * {----Brisbane Ph. Dept:+61 7 3365 3743 \ __ / Qld 4072 Fax: +61 7 365 3888 \,~' "\__/ Australia
Bo Johansen laments: } I am doing in situ hybridization on parafin embedded and sectined plant } material using a DIG labelled probe. However, I find that the binds } non-specifically to walls and cytoplasm. I have tested the anti-dig AB } and the do not show any non-specific binding.
Bo binding to cell walls is just one of the joys of working with plant material :-). If you are working with RNA probes try adding tRNA to your hyb buffer. Might also be worth trying things like a "blotto" pre-hyb step similar to membrane hybs.
contact me if you want to talk about this some more.
--
Daryl Webb (dwebb-at-waite.adelaide.edu.au) Dept. of Plant Science, Waite Institute University of Adelaide, Glen Osmond S.A. 5064 Australia. Voice:61_8 8303 7426 Fax:61_8 8303 7102
James Martin wrote: ==================================================== I am looking for information about the ISO 9001 standard with regard to the operation of FT-IR microscopy and SEM systems for materials analysis, and would very much appreciate communicating with list members who have practical experience in this area. ===================================================== Our analytical services laboratory is accredited by the American Association for Laboratory Accreditation (A2LA) to the standard of ISO Guide 25. While on the one hand, it seems like there is a paperwork requirement the describes virtually everything, and while that is at times frustrating, I am quite confident that we have a laboratory running on a far higher level since everyone is much more accountable. While in a sense it does add to our costs, the "cost of rework", that is, the cost of doing samples over again because they were not done right the first time, has gone down more than enough to compensate.
You can contact A2LA directly at the following:
American Association for Laboratory Accreditation 656 Quince Orchard Rd. #620 Gaithersburg, MD 20878-1409 301-670-1377, FAX 301-869-1495 http://www.a2la.org/
A2LA has been accrediting EM and LM laboratories under the discipline "Chemical Analysis" and subgroup "Microscopy". I would imagine that the extension from EM/LM labs to FT/IR microscopy would not be a very great leap.
We have no connection to A2LA except as being one of their accredited laboratories. A satisfied customer, in other words.
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 ############################ ==================================================
James Martin wrote: ==================================================== I am looking for information about the ISO 9001 standard with regard to the operation of FT-IR microscopy and SEM systems for materials analysis, and would very much appreciate communicating with list members who have practical experience in this area. ===================================================== Our analytical services laboratory is accredited by the American Association for Laboratory Accreditation (A2LA) to the standard of ISO Guide 25. While on the one hand, it seems like there is a paperwork requirement the describes virtually everything, and while that is at times frustrating, I am quite confident that we have a laboratory running on a far higher level since everyone is much more accountable. While in a sense it does add to our costs, the "cost of rework", that is, the cost of doing samples over again because they were not done right the first time, has gone down more than enough to compensate.
You can contact A2LA directly at the following:
American Association for Laboratory Accreditation 656 Quince Orchard Rd. #620 Gaithersburg, MD 20878-1409 301-670-1377, FAX 301-869-1495 http://www.a2la.org/
A2LA has been accrediting EM and LM laboratories under the discipline "Chemical Analysis" and subgroup "Microscopy". I would imagine that the extension from EM/LM labs to FT/IR microscopy would not be a very great leap.
We have no connection to A2LA except as being one of their accredited laboratories. A satisfied customer, in other words.
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 ############################ ==================================================
I would appreciate any comments regarding charge back fees for SEM and TEM services. Specifically, I would like to know how the basis for the charges are derived. The rates listed in the Tech Forum varied so much I was wondering if anyone has worked out a formula on whether to charge per sample or per hour. We are trying to compare our rates with those from other facilities and would really appreciate comments, suggestions etc.
Thank you,
Cora Bucana UTMDACC Houston, Texas
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