NetNotes


structure once this is all over and we get back in the lab. Esteban Fer- nandez g.esteban.fernandez@gmail.com


Calibration with Microspheres Confocal Microscopy Listserver We did some extensive intensity calibration experiments with different


intensity green and red microspheres. Te green microspheres were 3.7% and 35% relative intensities with an intensity ratio of 10.6. We measured the intensity ratio with lots of different microscopes and lots of different lenses and very consistently got a ratio of 8. I can’t seem to figure this out at all. It means the bright beads must be a bit dimmer than expected or the dim beads a bit brighter than expected. Te calibration would have been done by TermoFisher—I would guess they do this by flow cytometry. Maybe it could just be that microscopy measures the intensities more accurately? Does flow just get one data point per sphere? Maybe the bright beads bleach relatively more than the dim ones when you are imaging on a CLSM? Any ideas are welcome! Claire Brown claire.brown@mcgill.ca


Just curious - were the measurements done in water? Scott Hen-


derson schenderson@scripps.edu Perhaps they used a different emission filter? Michael Model


mmodel@kent.edu Tere are a number of possible ways to measure bead fluorescence,


it would be helpful if you described your pipeline in detail. Are you segmenting beads by intensity and summing up the total fluorescence per bead? Or taking the average? Are you performing background sub- traction? As Patrick inquired, are the beads the same size? Te likeliest explanation, given that you’re consistently underestimating the ratio between them is that there is a fixed background in the image. For ex- ample, if the dim beads represent 100 counts absolute and the bright beads 1060 for a ratio exactly of 10.6, a background of 50 counts would give you an apparent ratio of 8 instead. To correct for this, you can either subtract the background or do a 3-point calibration with 0-in- tensity ROI’s of the same size as your beads are randomly distributed in the image background. Pavak Shah pavak@ucla.edu


Tey were done in CyGel from Biostatus Ltd. I guess there could


be some quenching from the medium. Te imaging was done on over 40 different microscopes with different magnifications, immersion medium and NA and instrument settings, which is why we used the intensity ratio. Te dim and bright beads were in the same sample though. Tey are the same size, but perhaps the brighter ones could heat up more? Not sure how much heating you might get with a green laser though. It could also be that flow obtains the max intensity at the middle of the sphere but in microscopy we measure the whole thing. However, I would think that any difference like that would disappear with the ratio. We did correct for background so that is not it. I guess we could also have the wrong lot information. We got them from a 3rd party not directly from the company. Claire Brown claire.brown@ mcgill.ca


Size of a Photobleach Point Confocal Microscopy Listserver When using the Bleach Point feature in Leica confocal systems


(I’m sure it also exists in the other brands), I don’t know how to measure the surface of the bleached area. I figure out the wavelength of the laser beam and the numerical aperture of the lens affects the size of that area, so would it be correct to use the resolution formula to


2020 July • www.microscopy-today.com


calculate it? Or am I mixing up things here? Xavier Sanjuan Samarra xavier.sanjuan@upf.edu


I tend to use the PSF generator in ImageJ/Fiji (http://bigwww.epfl.


ch/algorithms/psfgenerator/). You can take the full width at half maximal to indicate the bleach area. In x, y, and z. I tend to think of bleaching a volume rather than an area. I’m sure this isn’t perfect, as the energy isn’t equally spread within the PSF. If I’m teaching this, I use the plot profile on live view of a PSF, fix the axes ranges on the graph, and run through the z stack to show the energy distribution in different z slices. Or save a small (65×65×65 pixel) PSF with 50 or 100 nm xyz pixels and use ICY 3D view- er with a color map (LUT) like Parula. I always need to fix the metadata in ICY to the correct xyz dimensions. Dale Moulding d.moulding@ucl.ac.uk


For fastest photobleaching, it is generally best to use maximum


power for as brief a time as possible. Te more photons, the bigger the spot. Oxygen radicals and dye radicals diffuse short distances relative to PSF (nanometers, though activated tyramide radicals have a diffu- sion radius of ∼100 nm and can be restrained by increasing viscosity or additives. Cytoplasm has a higher viscosity and density than water, so the radical diffusion radius might be suppressed in live cells that are [over]expressing catalase). A(nother) good PSF calculator is avail- able free at SVI (https://svi.nl/NyquistCalculator). I liked this recent publication on the photophysics and photochemistry of fluorescence and generalization across the UV-Visible fluorophores: Aleksandr Ba- rulin and Jerome Wenger, 2020 J. Phys. Chem. Lett. 2020, 11, 2027–35. George McNamara geomcnamara@earthlink.net


For quantitative analysis of recovery aſter bleaching a spot with a


Gaussian laser beam (TEM00), the beam waist typically is used for the radius term. Tis is the width at which the normalized intensity of the beam drops to 1/e^2. To measure the intensity profile of the beam, you would take an image of the parked beam on a uniform, photostable sample, use ImageJ to get an intensity profile across it and fit a Gauss- ian to the profile. In principle, you could also do this from the inverted intensity profile of the bleached region in the first image acquired aſter bleaching, but if recovery is fast, this will be an underestimate of the “true” radius of the spot. Te FWHM is approximated by 2.355×the standard deviation of the Gaussian and the waist is about 1.7×FWHM. Te difference is important for quantitative analysis to obtain the trans- port characteristics of what you are bleaching because the area of the bleached region is significantly larger using beam waist than using FWHM. Kate Luby-Phelps kate.phelps@utsouthwestern.edu


Plexiglas may not scatter and absorb light the same as your


biological samples, but with SP2 AOBS and SP5 AOBS systems we bleached spots inside fluorescent Plexiglas and then went back and im- aged a Z-series of the bleached volumes to see focal plane and cones above and below. Two examples: https://www.flickr.com/photos/ mcammer/2605562876/ and https://www.flickr.com/photos/mcam- mer/2608080259/. Plexiglas previously worked well as a model for cell cultures, but we are having an issue now (ok, in February and maybe again someday…) that as we focus deeper in drosophila embryos us- ing a Bruker galvo miniscanner that there is a lot of scattering. At the surface of a zebrafish embryo we can target membranes specifically, but deeper in drosophila scatters too much for fine targeting. We have tried three different lenses and adjusting the collar of one empirically. Tis is at 405 and 470 nm. So spot size is very depth and sample dependent. Michael Cammer michael.cammer@med.nyu.edu


Tanks for all the feedback! I did not want this for a photobleach- ing experiment, we have been using a multiphoton laser at 830 nm to


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