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Selected Area Electron Diffraction Microscopy Listserver Could someone please explain to me the principle of how a selected


area diffraction pattern is formed inside the TEM column? I understand the principles by which diffractograms from x-ray powder diffraction (XRD) are obtained: X-ray wavelengths, inter-planar spacings and constructive/ destructive interferences. However, I simply cannot wrap my head around the SAED pattern and how are they obtained? How is it possible to get a pattern from a 2D material such a sheet of graphene? Shouldn’t there be at least two layers to generate constructive interference (hexagonal dot pattern)? I imagine the beam hitting the graphene sheet perpendicularly while the sheet is perfectly horizontal on the TEM grid. Te scatter points (atoms) would scatter the electrons in all directions equally without a particular preference, yet we still see intensity maxima in the shape of a hexagonal pattern. I realize there must be a visual/logical flaw in how I imagine this process. Any help explaining/visualizing the real situation would be welcome. I’ve gone through materials online; however, it seems the explanations lack visual representation and are very trigonometry heavy which is not helping me to picture the situation in my head. Aruna arunasme@gmail.com


With one exception, your understanding of the scattering/


diffraction process is correct. Te one exception is the assumption that the thin graphene specimen on the grid is one atom layer thick. Not true. If you see a sharp, low background, hexagonal pattern on the screen the sample is at least 10 nm thick, which translates to 30 or so atomic planes thick. Te atoms in each plane scatter the electrons in all directions. All of these result that most scattering directions are canceled out. Only those scattering directions that satisfy Bragg’s Law will allow for the passage of a diffracted beam through the specimen, which results in the observed diffraction pattern. Ron Anderson anderson20@tampabay.rr.com


I suspect that you are picturing X-ray diffraction in reflection. In that case, you are looking at


the planes parallel to the surface


and, indeed, one plane won’t give diffraction. In TEM, we work in transmission mode. Each atom becomes a scattering center. You can think of this like Young’s slit experiment. Tere are 2 slits and the waves from each slit interfere to form a diffraction pattern. In TEM, the atoms are scattering centers and act in a manner analogous to the slits. Hendrik Colijn colijn.1@osu.edu


I would say you should treat a 2D crystal as a phase grating. You


can find quite a lot about diffraction by phase gratings on the web and the book on Fourier optics by Goodman also covers it. Here’s an attempt at an explanation: If every point of the graphene sheet gives rise to a spherical wave (Huygens construction) then the relative phases of these waves will be different depending on whether a point contains an atom or not. You can only get constructive interference in directions where the Huygens waves are in phase. Tose directions are the diffraction spots. In the direction of the spots the Huygens waves come from the atoms interfere constructively with another and so do the waves coming from the gaps between the atoms. Tere are examples of single-layer crystals that give rise to electron diffraction. Most membrane protein crystals are only periodic in 2 directions and they diffract very nicely. 3D-periodicity or multi-layers are not necessary for diffraction. I wouldn’t be surprised if a single graphene layer also gives a visible diffraction pattern. Philip Köck koeck@kth.se


Mounting Tissue Sections on Slides Confocal Listserver When I train people to use the microscopes in the facility, 90% of the


tissue section samples are mounted on the glass slide with lots of bubbles, so I am trying to convince people to mount the tissue section directly onto the


2020 September • www.microscopy-today.com


cover-slip. But many people told me the tissue won’t stick on the coverslips. Would you mind sharing how labs mount tissue sections? Are tissue sections mounted directly onto the coverslips? Do you charge or coat the coverslips so the tissue section will stick better for staining? Erika Wee wee@cshl.edu


Having done a little bit of histology myself, I can say that it’s a lot


easier said than done to get tissue sections cut and placed on a coverslip! If they have issues with bubbles, I feel like that would be true whether they place the tissue on the slide or the coverslip. Is your bubble issue with cryosections or paraffin sections? Tat might help some of us with limited histology experience give some other troubleshooting advice. Rhonda Reigers Powell rhondar@clemson.edu


Te trick for me has always been to put a small drop of mounting


media on top of the sample on the slide, and then using a pair of #5 forceps (or any ultrafine forceps) to hold onto one side of the coverslip. Holding the coverslip vertically relative to the slide, lower the coverslip until the side opposite the forceps is resting on the slide, and then gently lower the side with the forceps until the media first wets the coverslip. Continue to gently lower the coverslip, allowing the mounting media to wick along the coverslip. Tis way, no air will get trapped under the coverslip. In short, gradually lower just one side of the coverslip so that the mounting media can wet along the length of the coverslip, and you will almost never get air bubbles. Ben Smith benjamin.smith@berkeley.edu


Tanks very much for your quick response. Our histology core


usually mounts sections onto glass slides, but the students working on post-staining and mounting used large coverslips to cover many tissue sections, and they used uneven mounting media causing the bubbles. Some labs here are doing their own cryosections and they have told me the cryo-sectioned tissue will not stay on coverslips. Tis is fine if they are using 10× or 20× objectives for whole-section imaging, but for RNA-FISH or for higher resolution imaging, mounting the section to the coverslip becomes more critical. Erika Wee wee@cshl.edu


Could this be due to the slides being charged, while the coverslips are


not? Most of the slides I have used are charged to encourage the tissue to stick. Otherwise some sort of surface preparation or pre-coating may be necessary to promote adhesion. Craig Brideau craig.brideau@gmail.com


Tere are definitely times when I wish tissue would adhere


directly to the coverslip, but as you said, the issue is typically that there is no charge on the coverslip to help the tissue stick. Even if it sticks initially, it’s likely to fall off during processing/staining steps. Plus, the fragility of the coverslip means it might break and then you have other issues. Plus, if you actually try cryo-sectioning, you start to realize how much skill it takes to get the tissue on the slide, and you have a much bigger surface area target on a slide than a coverslip. People who do this successfully have completely earned my respect! If the bubbles are the main issue, I would have them practice their coverslip technique, even with no tissue on the slide. I’ll add that I use the back of a pipet tip to gently and carefully squish out any bubbles before I seal the coverslip or allow the sample to dry in something like ProLong Gold. Rhonda Reigers Powell rhondar@clemson.edu


If you have access to a plasma cleaner, putting your slides with


sections in a plasma for 30 sec or so just before cover-slipping makes the slides and sections super-hydrophilic and the mounting medium wets like a treat. I found this trick quite helpful for minimizing bubbles, especially those that love to form right over the section. Tobias Baskin baskin@bio.umass.edu


Coverslips can be charged or subbed, just as are slides. Several methods have been mentioned in this thread, others may be found in the


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