Hi Fei! Apologies I missed this question. For fluorescent proteins, you need to run a single color control that has mCherry. I am not aware of any mCherry beads. I would suggest if you have a cell line that expresses mCherry at a higher level, you can use this in lieu of the experimental sample. Sometimes I suggest to my investigators that come to the lab to have a cell line that expresses the fluorescent protein well and to freeze down aliquots so you can thaw/use as single color controls. Thanks! - Kathy

Hi Hadee, thanks for your question, it's a good one. Very often we don't see a binary positive a negative population in our samples but rather a shift above the control and often this is not particularly large. But we should be careful about normalising by division of a negative because we aren't comparing the same things. The positive signal will come from expression of a marker or fluorescent protein or probe but the negative signal will be autofluorescence primarily plus some background staining. There are some times when we do want to do this though; for example if we're looking at modulation of a level of a marker with some sort of treatment and then it is perfectly fine to calculate a ratio (or fold-increase/decrease) because we are then comparing apples to apples. When we have overlap of the negative and positive signals, so when our positive signal is weak ,it is sometimes better to take into account both the positive and negative samples and in these cases this is where the resolution distance metric can be useful. It is very similar to the staining index that we discussed in previous webinars and takes into account the difference between the median of the positive and negative populations but also uses the spread of the data (as measured by the standard deviation) of both of those populations. It is calculated as (MFI+ - MFI-)/Sqrt (SD-^2+ SD+^2). The bigger the number the greater difference between the two histograms. This is often used in imaging or imaging flow cytometry data. Hope that makes some sense!

Hi! I would suggest checking out our video on voltage optimization for a more in depth answer. kzfaq.info/get/bejne/jM6ljaeBu56ZeIk.html The ideal method would be to optimize your voltages for each of your detectors by carrying out a voltage walk up and determining where you get the best separation (with the positive signal all on scale) and using that setting for your experiment. This would take into account the MFI of the positive and the negative as well as the spread of the negative. We wouldn't necessarily need to decrease voltage just because a negative is at 10^3. - Kathy

I agree with Kathy. As long as the voltages were set correctly AND your data is on scale, it would be best to leave the signal were it is, as it will mean you have the minimum voltage that gives you maximum resolution. If the signal in your sample is offscale, there are two approaches: 1) if you're not looking at changes in expression levels then you can use less antibody (or if it's a control as is actually the case here, you can do this regardless as long as signal is still the same or higher than in the sample) or 2) you can lower the voltage slightly until data is on scale knowing that you may have lost some resolution. But it doesn't matter where the negative is really, as long as you have maximum separation and all signal on scale.

Thank you for the responses. We do 6color L/L panels between 2 canto II and a 10color canto I. So we use the Baseline SDden value from all 3 as well as positive bead MFI as part of a standardization procedure for the instruments. We try to put the unstained population 2.5x the background in the channel per BD.