OpenFlow: Experimental Voltage Optimization in Diva

When setting up a flow cytometry experiment, voltage optimization is required to ensure you are resolving your populations of interest to the
best of your ability. In this Open Flow session, we will review in depth how
to carry out a voltration of single color cells and how to determine optimal voltages through statistical analysis. Diva acquisition software
tools and tricks will also be reviewed.

[@woosherry4036]

Thank you very much. Now feel pretty confident about setting up our BD machine.

[@lifesrealityis]

Thanks for these tutorials! Questions at around 57-58mins the presenter gated the weakly PE positive cell population (P4) and changed the color and then said look backwards at the P/ P2 graphs. To verify the cell population wasn't cut off in the singlet gating. But by definition of it being in the P3 wasn't it obviously was not cut off? thanks again!

[@RuiGardner]

Hi! Thanks for the question. In this case, Kathy wasn't trying to verify what was cut off in the singlet gating, but the opposite. She was verifying what wasn't excluded. To be more precise, that dimmer positive population she gated happened to be some aggregates that hadn't been excluded. But visualizing the colored population in the doublet exclusion plot she saw that we were not gating out all the doublets/aggregates. Once she removed those from the singlets gate, that dimmer population (P4) disappeared. A nice way of using backgating to verify if we have cleaned all our data appropriately. It's one of several tools to improve the quality of the data.

[@sjjordan1981]

If I'm designing a 12-color panel looking at rarely expressed cell surface markers and intracellular cytokines, what is the best way to compensate the voltages? Should I purchase highly expressed markers using identical fluorophores (e.g. CD3, CD4). Using the targets I'm studying likely won't generate nice populations to calculate the stain index.

[@DerekDaviesCytometry]

Hi Stephen, Your question is a good one and one that does come up - we always say that the first rule of compensation is that you need to use the same fluorochrome in your controls and you do in your samples. So don’t substitute FITC with Alexa488 for example. However, that fluorochrome doesn’t have to be attached to the antigen you use in your samples. In the case of FITC, it will be the same no matter what it is attached to and no matter which vendor it comes from. So you can certainly substitute a more highly expressed marker. One caveat though is that as you are using an intracellular marker in your panel, the CD3-FITC would need to be treated in the same way ie used on Fixed and permeabilised cells. This will control for any effects that sample prep introduces. Although you don’t mention any, this doesn’t apply to tandem dyes. We can’t nix and match those as each will have their own unique fluorescence emission profiles (the ratio of the donor to acceptor emission will vary). Remember that this will help you optimise the voltage but you would still have to ensure that you are using the cytokine Ab at the correct titre, you can’t use a substitution for that.

[@fabiankeppler315]

It seems you focus on BD DIVA for this course. Why would you determine optimal PMTV manualy when you have used CS&T and have run a baseline for all parameters? What with PMT drift? What do you consider linear range? It's not the end of the axis (or 246K) I feel you understated the importance not to change the PMT voltages yourself, If you use CS&T. It is hard enough to convince users that the instrument has been calibrated for optimal performance. Autofluorescence does not change the sensitivity of the PMT nor the resolution does it? If your signal is off-scale when using fluorochromes then this has another root cause, not? How will you compare data from different users when every user has his/her own PMT settings? How would you explain titration of Ab's when you also need to optimize the PMT? Why look at log when trying to figure out "off-scale", linear scale is more useful not?

[@OpenFlowCytometry]

Hi Fabian. Thanks so much for your comment/questions. CST voltages are determined by placing the dim peak at an MFI that is 10x the SD of electronic noise. This does not always equate to the best voltage to use for our experiments. What is your question regarding drifts in PMT voltage? There will be variations from day to day, yes. If there are researchers that are looking to compare MFI voltages from day to day for longitudinal studies, it is not sufficient to go based off of CST voltages. You would have to utilize application settings or use hard dyed beads to set target values for the channels you are utilizing. Linearity is determined by the instrument, and you can see the linear range of each detector when you are in the CST program. It is important that users understand that it is okay to change voltages and how to optimize the PMT voltages for their experiments. The CST voltages are not always going to be ideal. When you calculate the stain index for your specific cell type with your specific single color cells, you can see differences in where you obtain optimal separation. If your signal is off scale, it can be that the voltage is too high, that you may need to titrate down your antibody, etc. I would not suggest for different users to compare their data unless they have run some type of standardization across their assays, with the method I'd mentioned above using target values. If data is off scale, you can see this both in linear or log. If we are looking at (non cell cycle) fluorescent data, it is most always done in log so we keep it in log and within the linear range of the detector. Hope this helps!

[@Mariiamdov]

I have a naive question maybe.. I am setting compensations with beads (i know its better to use cells) but often its time consuming or not possible to obtain cells. When i set the voltages, inittially I look at my unstained sample. I check that peaks of all the other flurochromes on the unstained lie at least 10^2. For example, wehn I run an unstained, BV605 is usually at 10^4 for me. So in this case I have to reduce the voltage on the unstained. Does this makes sense or is this an innacurate way to do it? Secondly, when I run my single stained controls, I just check that the positive peak is highest and hopefully not overlapping with any other colours. If its not then I dont touch the voltage. My stainings are not looking great though. I am wondering if I should pay more attention to having a better separation between the negative and positive peaks in the single stained compensation control? I find for some colours I cant get a good separation, especially PB where the neg and positive dont separate well at all.

[@RuiGardner]

Hi Maria, I'm happy to chime in. Setting the right voltages/gains is crucial for a successful run in a typical flow cytometer. That should be the first thing to do when you run your experiment for the first time. Placing the negative around 10^2 is not an optimal way to find the right voltages, and it stems historically from the old analog instruments where people would set the voltages to place the negative population in the first quadrant. It worked for most of the cases, but it was an empirical assessment, and nowadays we know that at best it leads to suboptimal gains for many of your channels (as you are experiencing). There are several ways to calculate the minimum voltage that will give you maximum resolution (separation). Besides our videos, I encourage you to take a look at this poster: www.researchgate.net/publication/329103887_Evaluation_of_Multiple_Techniques_for_PMT_Optimization, where they compared different methods. I would suggest using the one from Anna Brooks and Simon Monard, where they use the negative/unstained cells (this is important, you need to use the cells that you'll be measuring/staining) and then you can either spike it with beads that fluoresce in all channels and are excited by all lasers, or just use your single stain controls as long as you know they have good separation. Depending on the size of your panel, this can take a whole morning/afternoon, since you need to measure the medians and SDs of the negative pop in each channel and use something like Excel to calculate the stain index. Once you plot stain index vs voltages, you should obtain a sigmoid curve for each channel, and you choose the minimum voltage that will give you the maximum separation (right when the plateau starts). Once you have your instrument setup with the right voltages, you should titrate your antibodies, again using the stain index, to find the best concentration (too much antibody not only is a waste of money, but it also impacts resolution negatively, since the increase in background unspecific staining affects separation). Once you do this, you're ready to acquire the whole panel. It seems a lot of work, but you do this once per instrument and per cell/tissue/sample type. The benefits of doing these things properly from the start are night and day. Hope this helps. Let us know if you have any further questions. All the best. Rui

[@junaidrehmani9829]

it may sound silly but I am having a hard time understanding, how adjusting the area scaling factor helps us with setting cells ON scale for FSC.

[@DerekDaviesCytometry]

The Area Scaling Factor (ASF) ensures that the Height and Area signals from each event are in proportion. In theory we should probably adjust this for every cell type but if you are chiefly using the same cells that may not be necessary. Once the ASF is set (so that the population in FSC-A v FSC-H falls on the diagonal), use the voltage to set the population to be in the correct place on the plot Hope that helps! Derek.

[@OpenFlowCytometry]

Agreed with everything that Derek just said and wouldn't change a thing! To help get a visual explanation of this, I recommend the following video. It is a quick watch and may be helpful. kzbin.info/www/bejne/jKKxnpqtj9d9aa8 - Kathy

[@junaidrehmani9829]

@@OpenFlowCytometry Thank you ! Kathy and Derk, much appreciated. One more question, Then should we technically see a big single blot that increases in size with the number of events. Why I am saying this is cos we are looking at uniform cell sub type (CD4+ T cells). That arises the question why do we see a Diagonal?

[@DerekDaviesCytometry]

@@junaidrehmani9829 When we are looking at doublet discrimination we use Area v Height and, as above, these should be in proportion so the single cells will fall along the diagonal. For other measurements we use the area signal and for immunofluorescence staining, if we are looking at two markers, we will see a cloud rather than a diagonal as the variation in marker expression is usually not related. If looking at CD4 epxression there will be a natural biological variability in the number of antigenic sites as well as variability introduced by sample prep and the cytometer so we see a spread of data, something we will always get.

[@kyshark1]

@@junaidrehmani9829 did you even bother to watch the BD video that Kathy suggested? It clearly explains the relationship that you are struggling to comprehend.

[@matini-hd]

Thank you so much for these wonderful videos. I am, also, kind of having issues setting a 17 colors 🙈 panel. I have a, maybe silly question: how is it connected the antibody dilution with the method you describe in this video. Shall i titrate my antibody and check for the stain index for each titration and then choose the right titration at its correct stain index? Another thing I did not understand is why do you plot PE vs BV421?

[@DerekDaviesCytometry]

Hi Stamatia, I would normally recommend starting by optimising the voltage for the channel by using the lowest dilution - this should give the brightest positive signal (slight caveat, check that there is not too much background staining by running an unstained control). At that voltage then run your titration samples and I normally derive a Stain Index for each point which will allow the optimal titre to be established. The reason we plotted BV421 against PE is that sometimes the positive population is easier to see in a dot-pot than a histogram especially when the signal is low and/or the cells are highly autofluorescent; we don't always have to use a histogram and cam derive the stats e need for the stain index from either a histogram or bivariate plot.

[@matini-hd]

@@DerekDaviesCytometry Dear Dr Davies, thanks very much for coming back to my question. I will have to try this and see how it does actually work in my hands. there are still things I struggle though: shall I adjust the brightest signal at 10^5 by changing the voltage, in other words to get it on scale? And then, if I have 7 or 8 dilutions of my antibody shall I go through all this process to define the right concentration on the correct voltage (Voltage walk from 200 - 700V)? It is very confusing. I have a 17 colour panel and I cannot see how dealing with that.....

[@OpenFlowCytometry]

@@matini-hd Hi Stamatia! When starting out, if you are doing an antibody titration you can also load your sample that has the highest concentration of antibody. Make sure that the signal for the positive is on scale by adjusting the voltage. If in Diva, the first tick mark after 10^5 is typically where I want my brightest signal to be. Then you can be confident that all of your antibody titrations will be on scale. Run all samples at this voltage and then calculate your optimal antibody titer. From there, you can do a voltage walk to determine the optimal voltage to get best separation with the antibody concentration you are using. Does this help?

[@OpenFlowCytometry]

This is coming from Kathy, by the way!

[@matini-hd]

@@OpenFlowCytometry Hi Kathy, you actually made my day much nicer! Yes it helps a lot and makes total sense now. I m on it next week and fingers crossed it will work out the way I want! Many thanks to you and the team for this great videos and supplementary explanations! 😊 Big hug from Heidelberg-Germany!