NetNotes


could be an attractive purchase. Avoiding over-tightening works as well, whether with the pencil or the proper tool. I’ve never broken a pencil in LN2


, but surely it’s not the intended application. Michael Elbaum


michael.elbaum@weizmann.ac.il I found that a very nice replacement for the tool to handle the pin


grid boxes are Pilot pens. Tey actually work better than the TFS tool. Ruben Diaz pindusito@gmail.com


I cool down the metal part very well with liquid nitrogen before


using it. When the metal part becomes super cold, the metal holds the tip of the grid box cover well. When turning, I also push down the tool. If I do these there is no turning of the metal part without unscrewing the cover. Reika Watanabe reika.watanabe.c@gmail.com


Tips on Polishing Paraffin Wax Samples Microscopy Listserver I have the opportunity to do some very cool EBSD work on a


fossilized eggshell (probably almost entirely calcite), which has come to me embedded in paraffin wax. I find that my normal polishing process is not going very well. I’m curious if anyone on this list has had experience with polishing a sample in paraffin, or if there is a safe way to remove paraffin from the equation entirely and proceed with a more traditional epoxy. Te eggshell fragment looks cohesive, so I’m optimistic that it would survive a gentle removal of paraffin. Tank you all for your time and expertise! Omero Felipe (Phil) Orlandini omero.orlandini@jsg.utexas.edu


I would remove the paraffin with a solvent, classically xylene


was used in pathology but nowadays there are alternative solutions. Stephane Nizet nizets2@yahoo.com


Te eggshell should survive removing the paraffin with no problem.


I’ve done this with small crustaceans: fix, process to paraffin and embed, carve away the unwanted wax and crusty bits to expose the internal structures, de-embed to EtOH, CPD, and image in the SEM. Samples were fine. Might be an idea to practice on a chicken eggshell first. Te protein matrix would be the most likely part to be affected by the treatments, and this should be less of an issue with fossilized eggshell. Any protein leſt should be robust. Polishing: I assume you’re using a flat- lap. If so, the advice I was given was to make a figure 8, oriented vertically, so: lap wheel O, polish direction 8 (not on its side). And, if you’re doing EBSD, you’ll want to ion-mill your sample. Mechanical polishing won’t give the best results for best EBSD. Phil Oshel oshel1pe@cmich.edu


Infiltration Issues with Chlamydia-Infected HeLa cells Microscopy Listserver I was wondering if anyone out there has experience working with


Chlamydia infected cells and could offer some advice? I am having infiltration issues that seem to be localized only to the inclusion. It seems that there is material present in the inclusion that is difficult to embed. Does anyone have an idea of what this material might be, or any alterations to the processing protocol that might improve the infiltration? Papers I have found that use TEM to visualize Chlamydia infected cells use standard processing protocols, and we are using a fairly standard protocol here as well: Fix with 2% GA, 2% PFA in 0.1M cacodylate, 1% OsO4


post-fix, ethanol dehydration,


PO, 1:1 PO:resin overnight, and embedding the next day. Our resin of choice is EMBed 812, medium hardness. Any advice would be appreciated, thank you! Nicholas Conoan nicholas.conoan@unmc.edu


64


Back in the dark ages (early 1980s), when I did my work for


Brunham and Peeling, we worked mainly with cell monolayers. Te problem is the structure of the Chlamydial bodies. In particular, the elementary bodies. Tey have a very thick cell wall, and the internal structures are very condensed. Tis makes them hard to infiltrate. Te reticulate bodies, on the other hand, are a dream. My solution was to use acetone dehydration. Propylene oxide is not necessary if you are dehydrating with acetone, and in my experience, membranes were not leached out with the propylene oxide step. Tis also works with cell pellets. Te only thing I cannot attest to is the success you would have with a tissue granuloma taken from an infected case, as I never worked with one of those. By the way, with acetone dehydration, rather than ethanol/propylene oxide, you do not need the uranyl acetate en bloc fixation to retain membranes. Leaching of the membranes seems to be associated with the propylene oxide, at least in my experience. Plus, you don’t need the propylene oxide, thereby getting rid of a very noxious and flammable agent. Paul R. Hazelton paul.hazelton@umr.umanitoba.ca


Are you using vacuum infiltration? Jerry Jasso jfasso493@gmail.com Another option would be to use microwave-assisted processing and


embedding (https://pubmed.ncbi.nlm.nih.gov/24357357/). Igor Kraev igor.kraev@open.ac.uk


Seconding and acknowledging Prof. Hazelton’s post/opinion on


the parameters to consider with these delicate ‘critters’. A research paper that might help, and contains some slightly more detailed information (including acetone dehydration and intermediate fluid-infiltration) is Bradley et al., https://www.int-res.com/articles/dao/4/d004p009.pdf. From my knowledge and experience, I would add to Prof. Hazelton’s proposals the following considerations/thoughts: (1) prolong the processing time in most steps (especially fix, washing, infiltration);


(2) aſter osmication for 3-5 hrs (4°C; 2 hrs at RT in fume hood!), wash in appropriate buffer 2-3x for 5min) followed by at least 1x 10min 50% EtOH or Acetone;


(3) incubate in 1% paraphenylene diamine (PPD) in 70% EtOH or 70% Acetone. PPD in combination with OsO4


‘mordants’


the specimen and retains lipids and ‘matrix’ substrate that is usually eluted by the standard dehydration protocol;


(4) wash at least 2-3 x (15min each) in 70% EtOH or 70% acetone (until most of the brownish ‘PPD-bleeding’ in the washing fluid is removed);


(5) proceed with a further step of 70% EtOH pure or 70% acetone, and then continue with dehydration as usual (80, 90, 96, 96% each 5-15min @RT, 100%, 100% (EtOH or acetone) at least 10-15min each @RT; fluid:


(6) Intermediate resin mixture (hardener/catalyst/


accelerator added) 1:1 only, at least for 2-3 hrs (@RT, specimen rotator);


(7) pure resin: 2-3 times; (8) Polymerization: classically, I used a 3-step polymerization: (water vapor-free polymerization ovens): 24 hrs @ 37°C, 24 hrs @ 45°C, 24 hrs @ 65°-70°C. Wolfgang Muss wij.muss@aon.at


Phenol Red in the Imaging Medium Confocal Listserver I have heard that the reason Phenol red is avoided in cell culture media


used for imaging is that it quenches fluorescence in some (at least the green) spectrum. I cannot find any reference about this. Does anyone know of a paper showing this? Tanks. Sylvie Le Guyader sylvie.le.guyader@ki.se


www.microscopy-today.com • 2021 November


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