We all know, generally from bitter experience, that experiments don’t always work first time and that sometimes the little things that govern success are the things that get left out of that online protocol!
- Make sure the BrdU (5-Bromo-2´-Deoxyuridine) you add to you culture is fresh. The half-life of the reagent at 4°C is short, so don’t use one that has been in the ‘fridge for more than a day. I make up 1 ml aliquots of a 100X stock (1 mM) and store at –20°C. And, a good rule for Lab life, don’t trust anyone else’s preparation!
- To detect incorporated BrdU, the DNA needs to be unwound either by acid, alkali, heat or enzyme treatment. And this can be the tricky part – too little and the signal is low, too much and the DNA signal will suffer – it may need some preliminary experiments to get the conditions just right!
- If using acid denaturation be sure to wash at least 3 times after this step – even a small amount of acid left will be enough to denature your antibody and you won’t see any incorporation!
- EdU (5-ethynyl-2’-deoxyuridine) is great when you want to combine other fluorochromes, but beware that some fluorochromes [particularly proteins such as PE (Phycoerythrin) and APC (allophycocyanin)] do not survive the processing and fluorescence is lost. There is an easy solution to this – do the EdU staining before any surface staining.
- Remember that nucleotide incorporation only reflects cells in S phase when labelling time is short – if BrdU is left in the culture medium for any length of time, cells that were in S at the beginning of the labelling period may have progressed through to G2 or even back to G1 – bear this is mind when interpreting any data.
- If you are using a protein binding dye (e.g. CellTrace Violet or CFSE), make sure that there is no protein present in your labelling buffer as this will also bind to the dye. However, it is important at the end of the labelling period to add back a protein-containing buffer that will absorb any unbound dye. You also need to let the cells rest for about 60 mins, allowing the cells to kick out any remaining unbound dye. This will lead to uniform labelling and to a low coefficient of variation (CV) of the starting population – exactly what you are aiming for.
- Titrate the dye – don’t always go by manufacturers recommended concentrations. Cultured cells, in particular, are variable in size. These dyes bind either to the plasma membrane or intracellular proteins and the recommended concentration may not be saturating for bigger cells like it is for small cells like lymphocytes.
- Do check the intensity of fluorescence after labelling – the brighter it is the more divisions you will be able to detect.
- Check the viability of the cells after staining – the dye should not be cytotoxic.
- Check the CV of the labelled, but unstimulated, population. The lower the CV the better your chances are of seeing resolved division peaks. And be prepared to start again if the CV is too high.
- Always run an unstimulated control at each time point– that highest peak you see isn’t always undivided, all your cells may be proliferating. It’s a simple control and we all love those, right?
- Don’t forget to include a viability dye in your analysis, we always want to exclude the dead cells from analysis in these assays!
- Beware of dye transfer. This can be a particular problem with membrane-binding dyes in mixed cultures. Cells can easily transfer parts of their membranes to unstained cells. This will increase the CV of the labelling and again mean that division peaks are less easily seen.
- Finally, analyze your data correctly – don’t put a marker on your unstimulated population and say x% have divided, this is not true and you need to take into account the number of divisions. So, if possible. use a computer algorithm to deconvolute the data for you.
Both these approaches to measuring proliferation by flow cytometry are extremely powerful; taking a little care in sample preparation and being aware of some of the possible pitfalls will mean that your experiments will be more likely to work the first time!
Image Credit: Chris Potter