NetNotes
You raised an interesting topic. Last month, I posted an entry
reporting some infiltration problems using EmBed 812. I use a similar protocol for Xenopus tadpole eyes (which are approximately 1 mm3), involving osmicating the samples with potassium ferricyanide. As a result, the samples are (again) impossible to cut, although the ultrastructure seemed well preserved, indicating that there is a dehydration/ infiltration problem. I am currently modifying our standard protocol (basically increasing dehydration and infiltration times) and will consider removing the potassium ferricyanide, as I was already advised. Tami Bogea
tbogea@interchange.ubc.ca Fri May 14 RE the query whether the FeCN might react with the resin, I
can’t help remembering discovery of the necessity for excluding NMA (MNA) from original Epon mix, using Epon DDSA (or Araldite-DDSA formula) instead, if one wishes to stain sections with permanganate rather than oxidize the embedding resin. (J Cell Biol 1965 26:309). Mike Reedy
mike.reedy@
cellbio.duke.edu Fri May 14 I have used permanganate as a fixative and stain and it is a whole
different beast. Te problem with permanganate is that it is such a strong oxidizing agent. Adding ferrocyanide to osmium reduces the oxidizing power of the osmium so should lower the risk that osmium would react with any resin component. Furthermore, the use of FeCN-Os is as a fixative so the tissue has been typically washed and dehydrated prior to exposure to the resin unlike the case where permanganate is used on sections. Tom Phillips phillipst@missouri. edu Tu May 13 I don’t know if this works for animal tissues, or works as well
as the OTO method, but a less toxic (yet equally tissue-blackening) protocol involves incorporating tannic acid into glutaraldehyde and/ or formaldehyde fixative, then “developing” with ferric chloride. Alternatively, just adding tannic acid to the fixative before osmium staining can also enhance membranes. For an older review: Chaplin, AJ (1985) Tannic acid in histology: an historical perspective. Stain Technology 60: 219–231. Cheers, Dr Rosemary White rosemary.
white@csiro.au Mon May 24 I agree tannic acid is a good approach. It is important to use
the “right” type of tannic acid. I think it is the low molecular weight species but I would check this before proceeding. Tom Phillips
phillipst@missouri.edu Mon May 24 Tom is right: it’s important to use the monomeric kind, like
Mallinckrodt 1764. Phil Oshel
oshel1pe@cmich.edu Tue May 25 You can use tannic acid as a mordant to lead to enhance
membranes stain. Simonesque and Simonesque, Journal of Cell Biology, Jan. 1976 is the reference. I don’t know if you can still get EM grade tannic acid from Mallinquot or not. Tat’s what you need. Tat’s what the Simonesque’s used. It will enhance membranes beautifully. Your fixation should be a standard glutarladehyde fix. Barbara L. Plowman
Bplowman@pacific.edu Tue May 25 So I took some more of the same brain tissue and used fresh
osmium w/o ferrocyanide and I embedded 100 micron Vibratome sections in fresh Spurr (according to Anne Ellis). Same result! Badly fixed brains (by another lab) seems to be the problem, not a reaction between osmium+ferrocyanide and EmBed 812. Geoff McAuliffe
mcauliff@umdnj.edu Tu Jun 3
Specimen Preparation: enhancing membranes I have a postdoc who would like to see the membranes of the
vesicles she is interested in “stand out” better from the cytoplasm in the cells. She brought me an old paper where the fixation protocol pretty much extracted everything out the cells, except membranes—similar to the days of using potassium permaganate, etc. I would appreciate any and all advice on fixation protocols that may help to enhance the
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visibility of membranes, while maintaining a good fixation of the cells without extraction of the cytoplasm, etc. Tom Bargar tbargar@unmc. edu Mon May 24 I have had good success with the OTO method—osmium
thiocarbohydrazide osmium of Seligman et al. 1966. You won’t believe how black your tissues turn! See Willingham and Rutherford 1984 J Histochem Cytochem 32:455–460
http://www.jhc.org/cgi/ reprint/32/4/455.pdf for a good start. Tom Phillips phillipst@
missouri.edu Mon May 24 We use reduced osmium as a secondary fixative for membranes.
Works especially well on cultured cells. Pat Kysar pekysar@ucdavis. edu Mon May 24
Specimen Preparation: stain variability on LM sections I don’t do a lot of semi-thin sectioning, but a recent request has
led me to the following question: Why would two (different animals) of the same thick (0.25 µm) semi-thin sections show different color hues when stained identically? Te stain is Richardson’s (1% methylene blue, 1% azure II, 1% borate) stained for 30 sec on a hot plate then water rinsed and sealed with Permount. One of the two sample sets shows more purple than blue? Any thoughts will be greatly appreciated. M Delannoy
delannoy@jhmi.edu Mon May 1 My first comment is that 0.25 µm is pretty thin for a semi-thin.
I usually use 0.5 µm. Unless you are carefully selecting sections by their interference colors while floating on water, I doubt they are all uniformly 0.25 µm and this might alter staining patterns. In addition, the degree of cross-linking of the resin will alter the staining properties so if there are different batches of plastic resin used for embedding or they were heated down in the oven at different shelf levels and the top of the oven was hotter, you may get differences in staining. 30 sec seems short for staining—I might go longer at a lower temperature to get a more even end result but my experience would suggest it is subtle differences in the resin that caused the difference. Differences in fixation duration might also impact staining. Tom Phillips
phillipst@missouri.edu Mon May 17 One reason is that methylene blue oxidizes in solution to a
mixture of azure A, azure B, and other thiazine dyes. Te mixture is uncontrolled, but this is done deliberately for some polychrome staining. What are the differences between your samples? Species, tissue, . . . ? If just different individuals, and all else is the same, I’d look to section thickness and subtleties of embedding, fixation and the like. Philip Oshel
oshel1pe@cmich.edu Mon May 17 Tanks to all of you who responded to my question so quickly,
I received many helpful insights and suggestions. I would like to summarize for you the responses. 1. Methylene Blue oxidizes in a mix of Azure A and B and other thiazine dyes. It is uncontrolled a my occur over time. 2. pH differences of the stain, if the sections are of appreciable volumes, pH differences of the two sections may have altered the pH of the stain. 3. Differences in pre and post staining drying times and position on the hot plate. 4. Permount contains organic solvents which elute basic stains from sections. I overcame everything by matching the interference colors, staining as soon as the sections dried, (30 sec staining), rinse, dry and controlled position on the hot plate. I mounted immediately in Permount, and then recorded the images immediately. Tis seemed to work perfectly. Michael Delannoy
delannoy@jhmi.edu Wed May 19
Specimen Preparation: picric acid I am trying to find information about the chemistry of picric acid fixation. Specifically, I want to know what functional groups are cross
www.microscopy-today.com • 2010 September
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