NetNotes


Edited by Thomas E. Phillips University of Missouri


phillipst@missouri.edu


Selected postings from the Microscopy Listserver from July 1, 2015 to August 31, 2015. 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: cells on coverslips


I have one user who wants to do a TEM study of cultured neurons on a coverslip. She wants to preserve elongated nerve processes. We cannot go with suspension culture and make its pellets. Can anybody help me here? Ravi T akkar ravi.thakkar369@gmail.com T u Jul 16 I have had success looking at drosophila testes that have been adhered to a coverslip and then processed into resin for TEM. I followed a method by J P Chang where you process the samples normally (I used a glass petri dish to contain the solutions with something on the bottom to hold up the coverslip and make it easier to pick up), infi ltrate on a shaker plate and then embed on a mold that I made from silicon. To separate the coverslip from the resin I dipped the sample in liquid nitrogen briefl y and didn’t notice any changes in the tissue. T e paper is: Chang J.P. (1971) “A new technique for separation of coverglass substrate from epoxy-embedded specimens for electron microscopy,” 37, 370–377. You can also process as normal, infi ltrate in the glass petri dish on a shaker plate and then use BEEM capsules to embed. You put some resin (don’t fi ll it right up) in the BEEM capsule and then carefully invert it onto the area of interest and carefully place in the oven. T e capsules can be quite easy to tip over. T ey can be separated using liquid nitrogen also. Jordan Taylor j.w.taylor@massey.ac.nz T u Jul 16 No problem, you can fl at-embed the cultured cells on the coverslip. Process the coverslips + cells as you would if you had pellets. You can probably use just glutaraldehyde, no formalin, since these are spread cells. I’d also add 1% monomeric tannic acid to the glut and/or OsO 4 to help preserve the membranes. Do the resin infi ltration with a nutator or the like, but try to limit the degrees of tilt (and slow speed), so you don’t get the fl uid everywhere. Go through to 100% resin as usual, then cut the bottom & cap off of a BEEM capsule, invert over the cells, Carefully fi ll with 100% resin – don’t get air bubbles! - and polymerize. Aſt er polymerization, drop the coverslip + BEEM capsule in LN 2 , and the capsule with pop off . Trim and section. Phil Oshel oshel1pe@cmich.edu Fri Jul 17


Specimen Preparation: wear pattern of used sputter target


We have Denton Desk II Sputter coater. Attached is the image of the glowing sputter target. Should it be replaced? https://www.fl ickr. com/photos/97321550@N08/19760866061/in/dateposted-public/ Ravi T akkar ravi.thakkar369@gmail.com T u Jul 16 T is is the normal glow pattern of a Denton Desk II in use. We’d need to see a photo of the target itself when not it use. Top open, looking directly at the target to get a good image. If there are any holes in the target, especially in the area of the plasma annulus, the target is toasted and needs to be replaced. Phil Oshel oshel1pe@cmich.edu Fri Jul 17


With a dubious target of 57 mm, I, with fresh gloves, remove it and lay it on a vertically oriented dissecting scope lamp. In a darkened


50


room, one can easily detect pinhole defects in the anulus. Fred Monson fmonson@wcupa.edu Fri Jul 17 T e other replies have been very good—light will help show a pinhole in the target when viewed from the other side in a dark room. I wanted to add a few points that might be helpful for other deposition systems also. For thicker targets, a profi le gauge oſt en used by woodworkers to duplicate a profi le can be used to help determine the depth of the wear-groove. T is gauge has a row of pins that slide when pushed and match the profi le of the item being pushed against. T is is helpful in harder-to-reach targets if the gauge can fi t. Oſt en the mounting plate for the target is made of another material (stainless steel or copper) and this will show up in EDS or other analyses when the deposited fi lm is analyzed if the wear track has broken through the target suffi ciently. Finally, a good metals reclaimer (and sometimes the target supplier) will be willing to pay for the remaining high purity metal target “scrap.” T is may help in purchasing the replacement target. Allen J. Hall ajhall@prairienanotech.com Fri Jul 17


Specimen Preparation: protozoa


What is the current thinking regarding conventional SEM of protozoa? I have some Trichomonas tenax fi xed in 2.5% glutaraldehyde that need processing for SEM (and TEM) and I was hoping to use my standard procedure through ethanol/HMDS. Will this be adequate, or will critical point drying yield superior results? Also, what is the best method of transition? Does the sample need fi ltering, and how do I process the fi lter? Would they adhere to a subbed coverslip, or will they all just wash off ? If using HMDS, can I process them in a centrifuge tube, and just sprinkle the dried sample onto a sticky tab? We have some microporous specimen capsules that I use for critical point drying, but fear the protozoa are too small, and there is not enough sample, to eff ectively retrieve them aſt er processing. If anyone does regular processing for TEM and has a reliable protocol for resin embedding protozoa, that would also be very useful! Any suggestions would be welcome. Natalie Allcock nsa2@leicester.ac.uk Fri Jul 17 I fi lter protozoa onto a membrane fi lter (pore size determined by the critters, but anywhere from 0.45 µm to 8 µm). First, sputter coat the fi lters on both sides, so you fi lter the beasts onto a conductive surface. T en add fi x (1–1.25% glutaraldehyde); obviously in your case, just fi lter the fi xed critters. ethanol:HMDS series 2:1, 1:1, 1:2 then 3 X 100% HMDS. I fi nd most protistans dry best at 60°C. You can CPD the fi lters with critters, but some (sometimes most) of them may come off in the CPD and be lost. Other times, this works fi ne. Just be sure you know which side of the fi lter your critters are on - the simplest way is to look at the pattern on the fi lter support, as it will be embossed on the fi lter. You might also try drying from tert-butyl alcohol. T ere’s an article about this in the May 2014 issue of Microscopy Today. I never use microporous capsules. I fi nd they shed particles and clog CPD valves. TEM: try processing with whatever your usual protocol is.


doi: 10.1017/S1551929515000929 www.microscopy-today.com • 2015 November


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