Hi Aditi Mahajan

Using different antibodies with the same fluorochrome will not give you any definitive answer. For example, when you run samples through the flow cytometer, how will you decide which cell is positive for CD73, CD90 and CD45 if all are FITC positive? all of your cells will be positive for CD73, CD90 and CD45 based on your fluorochrome when in reality ADSCs should not have many CD45 positive population.

You should have each of the antibodies conjugated with different fluorochrome and ideally does not have too much spectral overlap. you can check spectra for each of the fluorochrome available on this website https://spectrum.cytekbio.com/

For controls, you should definitely have one unstained control to give the flowcytometer background fluorescence. Run single stain controls to get the spectral signature for each antibody. If you do not have too many cells to spare for the single stain controls, you can use compensation beads. Also a viability control is essential to consider only live cells.

if you are using a traditional cytometer, you will have to compensate for each fluorochrome, if you have a spectral cytometer, you can spectral unmix the fluorochromes. If you are running multiple fluorochromes on a traditional cytometer, you can run Fluorescence minus one (FMO) controls to help you with compensation. if you are unsure that antibody clone works perfectly, you should also run Isotype control, which will also act as your internal negative control.

it is very important you design your panel appropriately to get correct results, you can use any supplier websites to design your panel. For example, https://www.thermofisher.com/order/panel-builder/#!/

refer to this paper for reference on the type of antibody conjugated fluorochromes, Article Evaluation of Human adipose-derived stromal cell behaviour f...

Hope this helps

All the best with your experiments!

Ritihaas Surya Challapalli Thank you so much for such detailed explanation. I would like to clear a few things. I do not add all the antibodies in a single tube. Instead, I use 5 separate tubes, one of which is unstained (so I have my unstained control for gating), two tubes which contain only FITC labelled antibodies (that's my single stain control) and two tubes which contain one FITC and one PE labelled antibodies.

I only wish to know what could possibly be the positive control here? And how important is isotype control for this experiment? I do not have any isotype antibodies so what could the possible alternative to isotype control be?

Hi Aditi - just to make sure I've got the experimental setup right, you're looking to run the following five tubes:

1: unstained control

2: CD73-FITC

3: CD90-FITC + CD105-PE

4: CD34-FITC

5: CD45-FITC + HLA-DR-PE

To answer your question directly, without redesigning your panel at all, the bare minimum controls you need are an unstained tube (which you already have) and a single-colour stain for each colour you are using - in this case, FITC and PE - to set up compensation or unmixing (depending on your cytometer). Acquire each single colour control, either run compensation on your cytometer or afterwards in whatever software you are using to analyze, and then run your samples.

A true positive control in this experiment would be cellular material that you know for sure express the marker. This can be pretty easy for some of the markers you have (peripheral blood cells, for example, will definitely contain cells expressing CD45 and HLA-DR), but may be impossible for some others. In lieu of this true positive, you should think about doing a titration curve for each antibody. In your case, you'd need to prepare six serial dilutions (one for each antibody) on cells that you suspect express the marker (better if you know they do but we can't always be sure). So if you usually stain all of these at 1:100 dilutions, you should prepare a series of 1:25, 1:50, 1:100, 1:200, and 1:400 (for example) of each antibody and single-stain with them. Acquire all of these samples (a total 6 antibodies x 5 dilutions = 30 samples for this titration) and then see, for each antibody, which dilutions gives you the best separation between positive and negative cells. If you can't see a positive population at all, you need to adjust your titration and try again for that antibody, or change your cellular material or antibody (some clones are just bad, this is how you find that out). But once you have a titration that gives you the optimal separation between positive and negative cells, you can be more confident that seeing "no signal" in a real experiment means the cells don't express the marker, rather than you just can't detect it (so not a perfect positive control but better than nothing).

The alternative to an isotype is Fluorescence Minus One (FMO) controls for tubes 3 and 5. These are control tubes in multicolour staining where you stain with all the colours minus one. Normally, this is critical for good gating around possible spread from other channels, BUT in your case, where there are only max two colours stained together, this is not as important. Still, if you have lots of sample AND are worried that CD90 and CD105 might be hard to draw good gates, it would be worth doing for them. For example, if CD105 makes a big smear, it might be good to have the FMO so you can know for sure what "CD105 negative" looks like (i.e. the tube you didn't add CD105 antibody to) and then gate anything positive off of that.

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Okay, so that's the answer to your direct question. However, there's a few oddities I wanted to ask about in your panel design, since it might be possible to improve on this a bit.

1. Can you add a viability dye to your panel?

Dead cells autofluoresce in many channels, and so can make it very challenging to gate appropriately. You should strongly consider adding a viability dye (something as simple as DAPI, Propidium Iodide, or 7-AAD would all work fine here and not add any complications to your staining, depending on what colours work on your cytometer) so you can gate out dead cells. If you absolutely can't purchase/trade for any of these reagents, at least turn on the most violet detector you have (more violet usually means more autofluorescence) and then gate out cells that have small FSC, higher SSC, and noticeable violet fluorescence without adding a stain (not a great solution but might help in a pinch).

2. Why are you only using FITC and PE?

If at all possible, can you redesign this panel to use 7 colours (one for each of your markers and one for a new viability dye, see above)? This would cost more money in reagents but would dramatically reduce your sample needs (since you can then stain and run one sample instead of five) and will also make a much richer dataset (right now, you can't know if your CD34+ cells are also CD73+, for example - having it all in one panel would let you answer questions like that). And if seven is too much for your cytometer, even trying to knock it down to two or three tubes by combining some into new colours would be better, and would save you a ton of time and reagents in the long run (and make your data better). If you let me know what kind of cytometer you have and what colours are available, I'd be happy to help with this.

Sorry for the dissertation of an answer, and let me know if you have any more questions or if anything there doesn't make sense. Good luck!