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NetNotes


Edited by Thomas E. Phillips University of Missouri


phillipst@missouri.edu


Selected postings from the Microscopy Listserver from January 1, 2016 to February 29, 2016. Complete listings and subscription information can be obtained at http://www.microscopy.com . Postings may have been edited to conserve space or for clarity.


Specimen Preparation: agar embedding cells


Some advice please on colorizing agar for embedding cells. In the past, my osmicated pellets in agar blocks were easily trackable, but when the cells are in a thin layer the cells don’t give much contrast, especially if not osmicated. I am collecting naked eukaryotic cells in suspension culture, but have problems bring down the smaller cells by centrifugation. Cultures containing cells are fi xed in aldehyde or aldehyde/osmium cocktail, collected onto Millipore or Nucleopore fi lters, then held in place by adding warm agar over them on the fi lter. When cooled, I peel off the fi lter, leaving me with a thin layer of cells embedded in the agar. I then cut the agar layer into small blocks to enhance fl uid exchange during subsequent treatments. My problem is, the cells are light colored to faintly black, and the thin layer of cells does not stand out well, particularly when in the various stages of Epon infi ltration/polymerization. I tried adding some azure II blue dye I had on hand to the agar so I could see the agar blocks more easily, but by the time I got up to epon/acetone, the color was gone. Any sugges- tions for a dye that will withstand both the water based and acetone solutions, or an alternative way to mark the little agar blocks so I can more easily monitor them throughout the process? Steve Barlow sbarlow@mail.sdsu.edu Fri Jan 29


I have had luck with fast green added at the 100% ethanol step.


I make a saturated Fast Green solution in ethanol and add a drop or two at the 100% ethanol step. It kept the agar blocks rather colored through subsequent infi ltrations and polymerization. Sorry I don’t have handy what “saturated” means in terms of mg/mL but if you are interested I could probably dig that up somewhere. Tobias Baskin baskin@bio.umass.edu Mon Feb 1


Not directly answering your question but making a remark: You are working with a cell monolayer, meaning approx. 50μm thin layer. T is is so thin that you don’t have to care about fl uid exchange; I don’t think that cutting your agar in small blocks will add anything. I would keep the whole agar block standing at the bottom of a jar, so you always know that the cells are on the upper side. Stephane Nizets nizets2@ yahoo.com Tue Feb 2


Specimen Preparation: speckled TEM samples


I am having a terrible time getting clean TEM sections. I fi lter the


primary fi x mixture with a 0.1µm vacuum fi ltration unit; I’ve tried changing the water and am currently using RICCA ultra-pure H 2 O; I have stopped en bloc staining altogether; I use Spurr’s Low Viscosity embedding resin. Most recently I have been ethanol-washing all razor blades used during specimen collection; I use a 0.04% FSG quench aſt er fi xation (didn’t seem to make a diff erence). T e fi ne speckling is most noticeable in muscle tissue, although I see it in other tissues as well. I just looked at some freshly cut, unstained mouse foot pad muscle yesterday and there were the black speckles all over the place! T e specks are tiny,


62


much smaller than 10 nm. It almost looks like immuno-gold, except that I haven’t done immuno-gold staining in years, and the specks are not rounded as you might expect with gold particles. T ey don’t seem to be clustered at any specifi c organelle, and they are defi nitely in the tissue, not on the surface. Someone please help, this is driving me nuts! Debra M. Townley debrat@bcm.edu T u Feb 4 T is could be a couple of things. First, did you use phosphate


buff er? If your concentration was a 100 mM or over, the glutaraldehyde, osmium and phosphate can precipitate in the tissue. Check unstained sections to confi rm. Also make sure all the glutaraldehyde is rinsed out of tissue before you get to osmium. Are the tissue pieces small? You can also microwave fi xative to get better penetration. Be careful with using reduced osmium and then staining muscle. Lipid granules are aff ected with the post-section staining with uranyl acetate and lead. Apparently the lipid material gets lost aſt er staining and rinsing, I have seen this phenomena and sometimes it will contaminate parts of the section. If you are sure it isn’t your staining protocol, then it must be the fi xation. Michael Delannoy mdelann1@jhmi.edu T u Feb 4 It would be interesting to see an image of your speckles. Wolfgang


Muss wij.muss@aon.at Fri Feb 5 Image of speckled mouse muscle: http://s450.photobucket.com/ user/wassupdoc1/library/Science Debra M. Townley debrat@bcm.edu Fri Feb 5 T ank you for showing us the image with “speckles”. I think that the problem might be in lead staining procedure. Please, look at Arvid B. Maunsbach & Björn A. Afzelius “Biomedical Electron Microscopy (Illustrated Methods and Interpretations)” 1999, Pages 207–228 <doi:10.1016/B978-012480610-8/50011-1> for explanation. We had also similar problems with lead carbonate precipitations in the past. Since our technician started to wear a face shield during sections staining with lead citrate the precipitates (contamination) in them almost vanished. T ere is an old paper on “How to remove such precipitates from ultrathin sections”: J Kuo, “A simple method for removing stain precipitates from biological sections for transmission electron microscopy,” Journal of Microscopy 120 (1980) 221–24. <doi:10.1111/j.1365-2818.1980.tb04140.x>. We tried this approach several times, but the results were not ideal in our hands. Oldrich Benada benada@biomed.cas.cz Fri Feb 5 Diffi cult to say when we don’t know your primary buff er / fi xation protocol, but I would agree that it looks like it may be due to either precipitate from a phosphate buff er, possibly due to insuffi cient washing between glutaraldehyde/OsO 4 steps, or osmium ‘peppering’, usually produced with buff er reactions in the osmium steps, or prolonged fi xation in osmium. I used to have similar problems, and resolved them by changing my buff er to HEPES or cacodylate, extra wash steps between the glutaraldehyde and osmium fi xes, and secondary fi xing in aqueous osmium as opposed to buff ered osmium. Good luck resolving it - let us know how you get on! Natalie Allcock nsa2@leicester.ac.uk Fri Feb 5


doi: 10.1017/S1551929516000286 www.microscopy-today.com • 2016 May


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