NetNotes


on a piece of junk on the slide, then you will have a region that will be out of focus. Ironically, clinicians wouldn’t care, since they could still make their diagnosis; but students seem to care a great deal when the scan isn’t absolutely perfect. As for fluorescence, one thing I found out pretty quickly when


demoing scanners 3 or 4 years ago was that many people use non- hardening mountants for fluorescence. Some scanners (3D Histech for example) have the slides vertical in the racks, and overnight the coverslips slipped (you risk gooing up the innards). Horizontal mounting of slides may be crucial - it’s hard to get your customers to change their specimen prep habits (and I believe the non-hardening mountants are better for many fluorescence tissue applications). Tat narrows the field considerably. Te most challenging part of fluorescence slide scanning for us is


finding the tissue. Tis is one weakness of the Zeiss system. It’s impor- tant to outline the tissue very accurately so that there are no empty frames around the tissue - fluorescence slide scanning is slow so empty frames waste time, but more importantly they can affect stitching for regions of the slide. Zeiss likes to show off their algorithm that lets you outline the tissue on the glass with a sharpie, but this hasn’t worked for us since the tissue is nearly invisible (if they have properly matched the RI with the mounting media) and you just can’t do it accurately enough. We then installed a 2.5× objective for a prescan with RAC contrast which in principle should work, but there have been some bugs with that. So we are leſt with cranking up the contrast on a bright- field macro camera image and guessing for the region. I think a proper means for finding the tissue is crucial for fluorescence. I would love to hear what other users’ experience has been in this regard. When you demo slide scanners, don’t fall prey to the “in the


interest of time” ploy. It is MUCH easier to scan a small region in the middle of a piece of tissue, compared to entire slides where you have to deal with how well the tissue finder works, focuses, and stitching. I suggest setting up some slides to run over lunch and come back and see how it did. James Jonkman James.Jonkman@uhnresearch.ca


We have two slide scanners in our core (Hamamatsu Nanozoomer


and Zeiss Axioscan), which are heavily used (many times nearly 24/7), and I have demoed a number of other systems. My favorite system hands down is the Nanozoomer. In terms of image quality, scan speed and ease of use I believe it is the clear winner among the field (at least for bright- field scanning). I have trained hundreds of people on both of our systems and while I could fully train someone to use the Nanozoomer for basic bright-field use in probably less than 15 minutes, it takes a full hour or more as well as significant “assisted use/retraining” time commitments for the Axioscan for each new user. Now, some of that is because we pre- dominantly dedicate our Nanozoomer for brightfield and the Axioscan for fluorescence (though they can technically each do both modes) but a lot of it is because of the complexity of setting up the slides in Zen and issues with locating/specifying tissue sections, etc., as has been men- tioned. Also, our Nanozoomer is going on 10 (!) years old and still going strong. I have word from Hamamatsu reps that the original Nanozoom- ers are still in the field (∼15 yrs old). It’s a good sturdy design, and as long as you don’t have “abnormal” slides (broken, coverslips hanging off, or God forbid wet mounted) it does what it is supposed to with very few issues. Denise Ramirez denise.ramirez@utsouthwestern.edu


Paraffin Samples in a Scanning Electron Microscope


Microscopy Listserver We would like to validate a method with SEM (meaning that another preparation method more suited to SEM cannot be considered)


64


that requires the analysis of paraffin blocks in a SEM. I fear that this will melt the paraffin. Have any of you had experience with observa- tion of paraffin-embedded material in a SEM? Optimally we would like to perform EDX analysis on the paraffin block surface, which would require enough kV to see Al and Si (I don’t know the energy lines off the top of my head). Do you think that the electron beam could heat the paraffin up to the melting point? Would it be possible to circumvent this by coating the sample with a “thick” layer of conductor? Stephane Nizet nizets2@yahoo.com


First, create a flat surface on the block of tissue. Preferably, you


can section into the block with a standard microtome. Alternatively, carve the block down with a scalpel blade. Second, remove the paraf- fin using standard histological techniques like removing the paraffin wax from paraffin sections. Xylene or a substitute will do it. Tird, gradually transition from xylene and through an ethanol series to 100% ethanol. Finally, critical point dry your tissue block, mount, and coat in the usually ways for SEM. View the flat or carved surface with the SEM. I can’t speak to EDS using this method. Jan Factor jan.factor@purchase.edu


Although I appreciate any hints to avoid the problem, I may not


be able to process the paraffin block like sectioning and dissolving because I need the full sample to be able to compare and correlate with another method. Any treatment would require a new validation to ascertain that it does not impair the comparison and I would like to avoid that. For example, if I dissolve the paraffin I may lose the position of the particles. So my primary question remains open: will the paraffin melt under the electron beam? I saw that Al and Si peaks are under 2keV in EDX, so using 5 kV should work. Would working in low vacuum possibly decrease the heating of the sample? Would it still be possible to perform EDX analysis in low vacuum? Stephane Nizet nizets2@yahoo.com


Yes, the paraffin will very likely melt under the beam. Even


“hard” paraffin melts at 65 degrees C, and the sample will be hot- ter than that where the beam impacts the sample. Besides melting the paraffin, this will re-arrange the sample’s ultrastructure and likely cause the elements of interest to migrate. Te paraffin will also weaken (by absorption) or possibly obscure the X-ray signal from the sample. Plus, it will contaminate the SEM’s chamber and condense oil on the snout of the EDS detector (since it’s cold). So, yes, you have to de-embed the sample. De-embedding can produce excellent results; I’ve done this with both sections and for internal anatomy of small crustaceans by hand-carving the critters while in paraffin. Further note: You’ll want to make sample holders with a hole


in them, so the area of interest for EDS bridges the hole. Tis will decrease the X-ray background significantly. Drill a hole through a brass or (better) carbon/polymer cylinder. Coat with Aquadag or some such conductive carbon paste. (But do a background EDS on this - I’ve found high levels of phosphorus in some lots of Aquadag.) Keep in mind SEM stubs are almost all aluminum, so you’ll get an aluminum X-ray signal whether or not there is Al in your sample. Control for this. For Al and Si, you’ll want to use 4–5 kV. Te other question is: how liable are Al and Si in your samples?


For some elements in tissues, like Na, K, Mg, and the like, one has to use cryofixation/processing/cryoSEM, as these elements are other- wise lost during processing. Tis may be the case for Al and possible (though not likely) for Si. Meaning a negative result may not be truly negative, just an artifact of preparation. Te more preparative steps, the more likely the elements can be lost. If they are bound to some


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