I am interested in comparing the fluorescence intensity of stem cells in two different developmental stages of transgenic embryos that express nuclear localized mCherry. I will be imaging fixed and cleared whole mount embryos on a Zeiss 780 with Zen Black using a 20x 0.8NA Plan-Apochromat objective and trying to quantitate post-collection using Image J. As I see it, my main concern is making sure the mCherry signal is always within the linear range and avoiding any saturated pixels. I plan on setting the confocal conditions (laser power, detector gain, etc), using the range indicator to avoid saturation, on my presumptive highest expressing embryo, then simply re-using the settings for the remaining embryos. I am concerned about the changes in brightness as the Z stack proceeds deeper into the tissue, but perhaps a consistent approach with both stages will mitigate this imaging difficulty. We typically don’t quantitate pixel brightness based on our images so I am unsure how to collect them in the best possible way. Any suggestions or past experiences would be appreciated. Thank you!
Hi @Baileyellowdog, and welcome to microforum.
This is a pretty big topic! There are entire courses dedicated to this sort of thing … so it’s going to be impossible to give a complete answer in a forum post. There’s also a lot of sample/experiment-specific considerations that are hard to answer in a generic or universal way, so i’d hate to give you the impression that “if you follow these steps, you’ll be fine”. but here are some thoughts:
- you’ve hit the nail on the head for the most important things to remember when quantifying images of different samples: setup your imaging parameters so that you are neither saturating (nor under-saturating with an intensity of 0) in any pixel. Then don’t change anything as you image different samples and different fields of view. If you move to a condition that is brighter and you start saturating, throw it all out and start again.
- When collecting images for quantification, try to detach yourself from “how pretty it is”. Remember that even modern point scanning confocal microscopes have much worse detectors with more limited dynamic range compared to modern camera based detection systems… so it can behoove you to make some compromises for the sake of better signal-to-noise ratio:
- consider opening up the pinhole. you’ll collect more light (higher SNR) at the cost of contrast. (whether you really needed that contrast depends on the sample/experimental question)
- increase your pixel size. You’ll get a higher SNR at the expense of spatial sampling (“digital resolution”). In many cases, you don’t really “need” the spatial resolution provided by the default/nyquist sampling settings, and the compromise is sometimes worth it for quantification: so think about what you’re trying to resolve. If you’re just drawing a region around the whole cell and taking the average of it, you didn’t need much resolution at all!
- collecting more photons is always better: so (in a perfect world), turn down the gain of your detector, turn up the laser, increase dwell time, increase frame averaging, etc… This of course comes at the cost of bleaching, so if you’re doing z-stacks, you have to fidget with this (before you start your experiment) to find the best settings.
- consider whether you actually needed those z stacks? if your question is fundamentally 3d… then duh but remember that you don’t have to take a stack just because you’re on a confocal
- consider whether you really need a confocal microscope at all! Widefield microscopy will usually provide superior SNR, at the expense of contrast. Whether you “needed” that contrast depends on how much out of focus fluorescence you have in your sample. When quantitating, remember that the the intensity of an object is very sensitive to focus in a confocal, but much less so in widefield (which can be a good thing for some experiments).
- try to avoid pre-bleaching your sample by “looking at it” before collecting your “real image” (for instance, to focus, etc…). If possible, setup your positions using some other channel, or transmitted light.
- don’t ever try to compare intensities across channels (i.e. don’t attempt to make any meaningful intensity comparisons of your mCherry channel to an EGFP channel)
- don’t forget that samples can themselves sometimes be fluorescent (autofluorescence). So do the proper (no label) controls to determine how much baseline fluorescence you’re working with.
I’m sure I’ve left lots of important stuff out (as mentioned, it’s a huge topic)… so others may want to chime in with some more pointers…
Let me add a couple things to Talley’s excellent advice.
- Don’t forget about applying flat-field corrections. If the Zeiss 780 is within a core facility, you should be able to ask them for help with shading correction.
- Correct for spherical aberration
Thank you for your thoughtful recommendations! There are several things that I will need to consider in my microscope set up. I did a small pilot experiment and did in fact, have to throw out all the images when a slightly brighter sample showed up late in the day! So, you are spot on with that caveat! Thanks again.
Talley is right, this is a big topic! Here are a couple more things to consider, especially if the two developmental stages you’re looking at are quite different in size or structure:
Fixation and clearing can have an effect on intensity of FPs, and they might have a slightly different effect when performed on different developmental stages. There’s a chance that differences in intensity you detect might be due different effects of fixation and clearing on the FP itself. The best way to control for that would be to identify a marker that has the same intensity in both developmental stages - ideally something that’s been well-characterized in live embryos that haven’t been fixed or cleared. Then, perform the fixation/clearing at the experimental stages, and compare the intensity in the images.
Intensity decreases with depth into a sample, due to spherical aberration as noted above, but also due to light scattering and absorption. Clearing should prevent a lot of scattering and absorption, but if the stem cells you’re imaging are at very different depths in the embryo at the different stages you’re imaging, there may still be significant depth-dependent changes in intensity. The best case scenario is to position and/or section the embryos so that the stem cells you want to measure are as close as possible close to the coverslip, or at least at a similar depth between the stages.
Make sure you select the 16 bit option instead of the default which for some strange reason is 8 bit on a Zeiss system.
Low light mandatory for development. I would move to a light sheet system if it is available.