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4 mol (20% excess) of permanganate per mol of aldehyde is required to oxidize the alkene bond and the aldehyde group. Procedures for the laboratory scale treatment of surplus and waste chemicals (from an article launched to web by University of Geneva and really helpful): Formaldehyde is oxidized conveniently to formic acid and carbon dioxide by sodium hypochlorite. T us 10 mL of formalin (37% formaldehyde) in 100 mL of water is stirred into 250 mL of hypochlorite laundry bleach (5.25% NaOC1) at room temperature and allowed to stand for 20 minutes before being fl ushed down the drain. T is procedure is not recommended for other aliphatic aldehydes because it leads to chloro acids, which are more toxic and less biodegradable than corresponding unchlorinated acids. https:// www.unige.ch/sciences/chiorg/matile/12-ProceduresforLabTreat- mentofWasteChemicals.pdf Wolfgang Muss wij.muss@aon .at T u Apr 27


Formalin is historically ~40% formaldehyde in water and laced with 5-10% methanol to inhibit polymerization. T us, 10% “formalin” is 4% in formaldehyde. T at is the historically recommended concen- tration of the gas as a fi xative. 4% paraformaldehyde at 4ºC (buff ered or not) is also a very good treatment (10-20 min) for ‘freezing,’ muscle in thin tissues that are under various degrees of tension (e.g., in a partially to maximally distended urinary bladder), before they are immersed (or fi lled) with the same fi xative. Fixation at 4ºC is a very good fi xative for both routine and special histology. I stopped using formalin in the mid 1970’s, and substituted as follows. I usually make 20% HCHO from paraformaldehyde, and I can refrigerate it in 100 ml aliquots for over a decade without any polymerization. Formalin ought not to be used for anything in biological histology, unless the methanol is absent - which solution should not be called “formalin.” Fred Monson fmonson@ wcupa.edu Fri Apr 28


Microtomy: certifi ed thickness of sections A customer of mine has an ISO certifi cation of the EM lab coming. One of the problems is to reliably demonstrate the thickness of the 50 nm ultramicrotome slices and the semi-thick 500 nm slices in clinical workfl ow. Normally this is done in the fi eld by using the interference color of the slices in the water tray which is not very accurate and depends also on the color spectra of the illumination system, the resin system, the accuracy of the ultramicrotome, the user, room situation; what shall I say more...? Did anyone of you had this problem and how did you solve it? T is question is also posed to the manufacturers of ultramicrotomes, RMC and Leica. How do you handle this? Did anyone measure the thickness of the slices and how? T rough-focus-series in TEM? Some setup in SEM? Stefan Diller stefan.diller@t-online.de Fri Apr 28


Another thing to consider, is do you care about the thickness of the section (on water, grid, slide), or the thickness of the material removed from the block; i.e. the block advance (assuming the ultrami- crotome is cutting all the material off aſt er each advance). For our studies, the block advance, which can be related back to the original volume of tissue, is what we like to know. If you minimize compression (ultrasonic diamond knifes, diff erent angles of knife and/or harder resin), then the block advance and the section thickness on grid begin to converge. Ben Micklem ben.micklem@pharm.ox.ac.uk Fri Apr 28 I thought the original method to determine section thickness accurately was to shadow sections, at a known angle with the sections on mica (or similar), and then calculate the height (thickness) of the section by the length of the shadowed metal deposit and the known angle. Admittedly, this is a lot of prep work and not something one would want to introduce into a “routine procedure.” Lorenzo Menzel lmenz001@fi u.edu Fri Apr 28


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Work has been done to determine section thickness in a more exact way than using interference colors alone. One method is to use the minimal folds method using the TEM (Small, 1968). Imagine crinkling a piece of paper so a fold forms in the middle in the Z axis, which will be twice the thickness of the paper. T is fold can be measured on the TEM to get a relatively accurate thickness measurement. Ideally, measure multiple folds and get an average thickness. T is method is addressed nicely in the following paper: When using samples with mitochondria, you can also use the cylindrical shape of those to estimate section thickness (Fiala and Harris, 2001). T is is ideal if you have sections with very few wrinkles and was found to have an average thickness measurement very similar to that of the minimal folds method. Yet another method is to re-embed some sections you’ve cut and section them again at 90 degrees to measure the thickness (Bedi, 1987). Finally, laser confocal microscopes can be used to measure the section thickness with a reported accuracy of 1 nm (see Kubota et al. 2009). Connon T omas connon.thomas@mpfi .org Fri Apr 28


Immunocytochemistry:


Immuno-gold labeled cellular structures on the membrane We did immunogold labeling on MDA-MB-231 breast cancer cells grown on cover glass to see if the antigen would appear on the cell surface. T e cells were fi xed in 4% paraformaldehyde and 0.1% glutar- aldehyde in 0.1M cacodylate buff er pH 7.4. Immunogold labeling was performed before critical point drying and sputter-coating with Au/Pd. Images were taken on a Zeiss Sigma FE-SEM. Besides some scattered signals, a lot of gold particles are localized in one area. An example is here: https://goo.gl/photos/YPXZrwzNAukfsHBw7


Can you tell what the structure is? In addition, there were a lot of gold particles on the background/cover glass? Could you suggest ways of eliminating them? Zhuo Li zhuoli@coh.org T u Mar 23 T is is not easy to answer in just a few words. To evaluate the reliability of any immunolabeling, it is required to at least check the negative controls: specimens incubated without primary antibody. Based on those results one can then look at background and what to do about it. If the negative control is clean, you are dealing with labelling based on binding of the primary. Pre-adsorption of the primary can help provide answers as to whether what you observe is specifi c or non-specifi c. If the negative control is not clean, the labelling is the result of interaction between gold conjugate and specimen. In that case, incubation protocols and specimen conditioning are the fi rst thing to look at. Background, false positives can almost always be controlled. I will be happy to help with detailed suggestions if you would like, but since that might go into products and brands it would be better to do this off -list. Jan Leunissen leunissen@aurion.nl T u Mar 23 Some thoughts about this problem: 1. It seems, at least, two possibilities for your picture. One is the structure (with gold particles) could be extracellular matrix, which is usually ‘sticky’ for gold particles, or could contain the antigen. Another possibility is that it is a broken membrane and you are viewing the inner membrane, which might have more antigen, or be sticky. You may want to do a ‘control’ with full fi xation (glutaraldehyde + osmium), which provide you a better resolution of the structure. 2. Aldehyde group, especially that from glutaraldehyde is very ‘sticky’ to antibodies. Since you are not viewing the internal structure, I strongly recommend avoiding using glutaraldehyde, not even 0.1%. It may explain, at least in part, the heavy labeling on your cover glass. 3. Blocking with BSA, and/ or serum could reduce the non-specifi c background. 4. I recommend the sample be coated with carbon, instead of Au/Pd. You can then confi rm the gold particles with a backscatter electron detector (here is a reference I published many years ago - Localization of


www.microscopy-today.com • 2017 July


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