Hi, great question! If you are using 2 spatially separated lasers and 2 different PMT arrays, colors that have the exact same emissions but are excited exclusively by different lasers would not have overlap. Examples of this are dyes like PECy7 and APCCy7. PECy7 is excited by blue or yellow lasers (not red) and APCCy7 is excited by a red laser (not blue or yellow). So on our instruments those 2 dyes, while having the same emission profile, require very little to no compensation. This is an ideal case, but most dyes are not so ideal in that they usually are a bit excited by multiple lasers.

@@ucmercedscifstemcellinstru711 Thanks for the super clear explanation! And yes, I've noticed that some fluorophores, such as PerCP, have very wide excitation spectra, and thus you can get some spectral overlap but due to "excitation overlap". For example, if using PerCP-Cy5.5 and Super Bright 702, even though PerCP is most efficiently excited with a 488nm blue laser, the 405nm violet laser will still cause some excitation of PerCP-Cy5.5 and that will bleed over to the Super Bright 702 channel (excited with the blue laser), since both Cy5.5 and SB702 have similar peak emissions.

You're starting to get into challenging territory with 14 colors. Much of your success will rely on good experimental design. For example, putting the high density antigens and good antibodies on the dim fluors and the low density antigens on the bright fluors. And try to avoid using co-expressed markers on colors that have a lot of spillover. Fluorofinder is a good resource for building your panel to minimize compensation. If you already have your panel set, I'd say it's best to go through each fluor one by one and make sure that it's brightest in the channel of interest by at least a log if possible. Adjust your voltages to make that happen. For example, if you're using your FITC single stain, make sure it's showing up brightest in the FITC channel. If not, you'll get greater than 100% compensation values and autocomp may give you issues. So you'll need to do 13 plots where you're looking at FITC versus everything else and make sure your signal is brightest for FITC. Then repeat for every other color, adjusting voltages as necessary. This will take some time, especially the first time you set it up. There are some tricks to set up the plots in the global worksheet to make your life easier during this process. I can explain further if that would be helpful.

Great!, Can you please elaborate on that?, we can also connect via email Praveen98481@gmail.com

@@ucmercedscifstemcellinstru711 Hi, thanks very much for this video, it is very helpful! I would also like to learn more about these "tricks" to set up the plots in the global worksheet. Could you please share these with me? Thank you!

@@ucmercedscifstemcellinstru711 thank you so much i am also facing a same problem. In my exoeriment i am using 10 flourophore and right now I am struggling with voltage setting and compensation so I need your more guidance if you possible. It will really help me to proceed further in my research where now i am stuck. My email id: jadhavsuchi@gmail.com

He does in this video: kzfaq.info/get/bejne/hNKKlbB8tb27fac.html

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You can always record all your single stain tubes and samples and then compensate later, even in 3rd party software like FlowJo. I always tend to compensate while I'm running single stains and prior to running samples so I can see my compensated data while I collect it. Then I can make sense of the data while it's running on the cytometer and make sure the staining panel looks correct and as expected.

1) yes, just make sure you have a positive and negative bead and the fluorochrome is the same as what you stained your sample with. Here's a bead product that binds to antibodies and is often used if cells are not available or you don't have a great positive control stain: www.thermofisher.com/order/catalog/product/01-2222-412) 2) The compensation matrix is saved in the fcs file and flowjo will automatically apply the compensation values you set on the instrument. You can always modify or remove the compensations in flowjo as well. To do this you can double click on the little grid symbol that appears next to your sample name in the flowjo workspace. 3) Yes, in the flowjo compensation editor you can modify the compensations no matter how they were performed on the instrument. Thank you for your questions.

Hi, using comp beads, what voltages would you setup? would you run unstained cells, setup voltages, then run comp beads without touching to voltages even if neg beads are higher than unstained cells? Am I correct here? Thanks

I'm not a fan of setting up voltages based on a negative control. The reason is without a positive control you don't know how bright your positive will be. Thus, you may set it up based on the unstained cells and then you put your sample on and find that the positive signal is off scale (voltage set too high) or does not give optimal separation (voltage set too low). So I always try to set up with single stained cells that are stained with whatever is in the actual panel. Of course, you don't always have that luxury (rare positive events, very little sample to work with, etc). When I have to use beads, I would use the positive beads to set the voltage so the beads are bright and you get good separation between positive and negative beads. Then I would check with unstained cells to see that the background is acceptable with that voltage setting. You're right that beads often have higher background than unstained cells. Also, it's important to know your instrument. For example, I know if I'm setting any PMT voltage outside of the 400-650 range on my LSR II that something is wrong.

many thanks for your reply. I always thought that unstained cells could be used to set approximately the voltages, then of course checking if beads were ok and adjust voltages if needed and re-check unstained cells are not out of scale. How do you check the range of the instrument? CST beads? voltage titrations? Thanks

When a baseline is performed with CST beads (only administrators can do this) the software will determine what it thinks are optimal PMT voltages for each channel based on where the channel is getting good linearity and low CVs. It does a voltage titration and some calculations to determine this. This is the default voltage value that you see for each channel when you make a new experiment. In general that's a good starting point for where to set voltages, but more often than not I find it's on the high end. I know the ranges of my channels more based on a lot of experience with adjusting voltages and seeing what's best with the fluorophores we use most.

I think you're right about getting crucified by the cytometry purists! But frankly, visually compensating is a practical solution that works well and is important in getting the concepts across to users, rather than a black box solution.