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and monitored their growth. T e growth rate of the treated plants was obviously much higher than the non-treated plants, but the student wants to extend her project to see if there are any microscopic diff er- ences that she could visualize at the tissue level of the plants using a light microscope. Aside from building stomatal peels, I don’t have much experience with plant microscopy, especially not when it comes to hormone detection. Does anyone know of something she could look for? Beth Dixon bdixon@rafos.org T u May 10 T is sounds like fun. You can cut hand sections with a razor blade and look at them in brightfi eld (or fl uorescence if you have that ability). Many plant tissues are auto-fl uorescent. If you have access to a drawer with stains, you can try staining. Lots of these dyes stain plant tissues diff erentially, and this can simply provide more contrast for you (in either brightfi eld or fl uorescence). It doesn’t matter really what exactly they stain (and in many cases that won’t be well known). I am a little curious about the control your student used? Plants do have steroid-type hormones, but they are not exactly like those of animals. And certainly some steroid hormones don’t do anything to plants. I wonder if there are other materials in the inhaler “juice”? I would be happy to correspond with you offl ine about that if you like. Tobias Baskin baskin@bio.umass.edu T u May 10


Specimen Preparation: TEM artifact


We are embedding kidney samples that have been stored in paraformaldehyde/glutaraldehyde/phosphate buffer fix for a day to weeks using immersion fixation. I am seeing what looks like classic lead precipitate—dense round spheres over the tissue after staining with uranyl acetate/lead citrate. However, when we look at unstained sections, the artifact is also present everywhere on the tissue, not on the resin area of the section, only the tissue: mitochondria, nuclei, etc. Please comment on what is causing this. Sue Van Horn susan. vanhorn@stonybrook.edu Sat May 19 T is reminds me of “salt and pepper” precipitates due to your


fi xation—generally, processing of the tissue. NB: phosphate buff er (PB) and molarity of working solution? Too rapid or less careful dehydration (i.e., for example, dehydration out of buff er washes post-paraformal- dehyde or paraformaldehyde/glutaraldehyde or glutaraldehyde with 70% ethanol (for sure 76% ethanol) may cause (sometimes huge) PO 4 -precipitation within tissue. Not knowing about your using OsO 4 [if yes: in PB too?) as secondary fi xative and aſt erward. It would be interesting to see your standard processing protocol just to follow all the steps done until examination of the grids. Such precipitates also would also happen aſt er processing as before + uranyl acetate en bloc tertiary fi xation (= a pre-staining option) without applying rigorous washing tissue specimens before with the maleate buff er method/ sequence to get rid of the whole phosphates. Worst case would be (but for sure you can discriminate between microorganisms like bacilli or bacteria from long-storage PO 4 or other “ion” precipitation in specimen in primary fi xative solutions) if some detrimental alteration of your kidney specs happened during storage. Naturally, it would be of benefi t to see a typical micrograph/digital image of the precipitates (i) aſt er conventional double staining (uranyl acetate-lead citrate) as compared to (ii) unstained ultrathin sections from same specimen. Wolfgang Muss wij.muss@aon.at Sat May 19


I ran into this problem several years back. I changed almost every variable that I could: water source, fi ltration of buff ers and fi xes, new vendors for chemicals, etc. Aſt er an exhaustive search for the culprit, I found (through EDS on a new Hitachi TEM) that osmium was the major off ender! Do you use a post-fi x in OsO 4 ? If so, you will fi nd that adding 0.8–1% potassium ferricyanide as a chelating agent may solve your problem. T e solution will be bright yellow, like uranyl acetate; it will still act as an oxidizer (tissue will be black at the end of


2018 September • www.microscopy-today.com


an hour); you may process as usual. It took me forever to fi gure it out (it seemed like forever anyway...), and I’m hoping to save you time and aggravation. Debra Townley debrat@bcm .edu Fri May 25 I had a similar problem a few years ago. I contacted the listserver for suggestions and got the following from Ann Ellis. She was a wonderful source of help and information. It worked for me so give it a try. Ann Ellis email from archive: T e pepper sounds like the same old thing we have had from time to time over the years with glut and osmium fi xation. Traditional buff er washes will not solve the problem. I published a paper way back in Stain Technology (1979) 54:282–85. We ran out of reprints twice since every pathologist and his brother wanted one. I don’t have any more, or I would send you one. T e salvage method is simple. Cut new sections and pick up on nickel grids. Oxidize the sections with 1–2% (wt/vol) freshly prepared periodic acid for 5–10 minutes. Wash the grids several times with deionized water. [With nickel grids you can make the grids wash themselves by setting them on a magnetic stirrer at low speed.] Post stain as usual with uranium and lead. More importantly, I do have some ideas about how to prevent the problem from happening again. In my many years of doing cytochemical localization, I washed the tissue in buff er wash, which contained 0.5-1.0% (vol/vol) DMSO. T is removed the aldehyde and protected the enzymatic activity. In the last several buff er washes before localization procedures, I added 0.1 M glycine to the buff er wash. T is has been recommended for years for removing unbound aldehydes to improve immunolabel. I have never seen the osmium pepper in any of those preps. A while back I was going through the list server archives, and Randy Tindall had a post about putting a small amount of beta-mercaptoethanol in the buff er washes to prevent this problem. It probably works similar to the DMSO and glycine. Ann Debby Sherman dsherman@purdue.edu Fri May 25


SEM:


rotary pump’s gas ballast I was replacing mist fi lters for our JEOL SEM’s rotary pumps recently when it occurred to me that I never ballasted the pumps. In my previous job, I used a lot of mass spectrometers, and we ballasted the rotary pumps once a week. Mass spectrometers handle a lot of solvents, while SEMs don’t. So I’m guessing that we almost never have to ballast rotary pumps for SEM. Am I correct? Our SEM has Variable Pressure capability, but we don’t analyze wet or moist samples frequently. If anybody has any thought on this subject, I would appreciate it. Tsutomu “Shimo” Shimotori shim0102@umn.edu Wed May 9 We always ballasted TEM rotary pumps once a week, back in the day when they used fi lm. Wouldn’t seem to be necessary with an SEM, unless you use LV mode a lot with dirty specimens. I’m not sure that it would hurt, but in changing RP oil in our SEM once a year, I don’t see any problems with the used oil coming out of the pump. We use LV mode occasionally as well, but common sense regarding sample size and amount of outgassing materials shouldn’t make this an issue. Jim Ehrman jehrman@mta.ca Wed May 9


SEM:


vacuum problem and desiccant replacement We have a Zeiss 1450EP SEM that is having an issue with the vacuum. It gives a vacuum error message; it doesn’t want to recognize the vacuum hardware. Has anyone else experienced this problem with their scope? Any advice would be greatly appreciated. If we should just order a black wreath, please let us know that, too. We took the panels off the scope and noticed a fi lter on the side of the scope. It contains an amber-colored substance that we’re assuming is similar to Drierite. Does anyone know if we can recharge this fi lter by drying it in an oven?


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