I'm having lots of trouble getting a CFSE protocol to work for cells. I harvest splenocytes and then put them in 1%FBS in PBS with 2.5micromole of CFSE at 15million cells per mL. After 9minutes in the incubator I add 5x the volume with cold 20% FBS in DMEM. I then put the tube in ice for 10 minutes and then wash three times with serum containing media. I then plate the cells at .5million/ ml at 4mL total in a 6 well plate with 5microgram/mL of Con-A.

I rarely see nice sharp peaks of the CFSE stain after collecting on Day 1, Day 2, Day 3, Day 4. The cells might be dieing...But why I'm not sure. The FITC peaks I see are very broad and don't show the nice sharp divisions. I've been working on this protocol for months now. Any ideas??

I use a minor modification of your protocol, not that I really think that is the problem. I incubate the cells with 5 micromolar CFSE in just PBS for 10 minutes at room temperature. Minor difference and I'm not sure that it accounts for your troubles.

I'm tempted to believe that the mixture of cell types you have is contributing to the spread in the peaks. Can you counter stain with another marker like CD3 to identify the sub-population your interested in in the final analysis? Staining PBMC from human blood I find that the surviving non-T cell population (monocytes mostly) after an MLR has a lower CFSE fluorescence than the undivided T-cells. If I didn't have CD3 in my analysis I would also have one big sloppy peak of CFSE expression. You might get some idea if I'm right by looking at your forward vs. side scatter profile and see if you can identify suspopulations of cells. Gate on the subpopulations and see if that sharpens your peaks.

yeah, i've tried double stain with CD3 PE and still get a blur of populations.

Are you discriminating live and dead cells with PI or some other such method?

Can you post a picture?

I haven't been discriminating dead cells. I will try that next though with 7-AAD.

I've tried both splenocytes and a T cell cell line. I'm thinking that the splenocytes have several cell types and divide at several rates. But this still doesn't explain the smear I see when I double stain with CD3 or when I stained a cell line with CFSE.

Here are some pictures of the results i get.

Your staining looks pretty good at Day 0, although I'm a bit surprised by how quickly it declines. I assume your showing me the results from the cell line? In our hands, even after 7 days in a murine MLR we still had a tight peak of undivided cells. It looks like all your cells are dividing, so that makes me think this is your cell line. Even with a tighter peak of initial staining, discriminating subsequent populations is less than perfect, as you can see below. The data is from our publication: Chen JC, Chang ML, Muench MO. A kinetic study of the murine mixed lymphocyte reaction by 5,6-carboxyfluorescein diacetate succinimidyl ester labeling. J Immunol Methods 2003;279:123-33.

At this point my best guess is that the spread of your populations is due to a combination of three things. One, your cells vary in size (FSC), probably due to their position in the cell cycle. Do your peaks get sharper if you gate only on the main cell population seen in your FSC vs SSC plot? Do the slightly larger cells have a higher autofluorescence and thus are broadening the peaks? Second, your day 1 results may actually represent two populations, those that have divided once and those that have divided twice. Again, your cells seem to be dividing at a rapid pace given how fast they are loosing staining. Third, even cell lines have a population of dead cells and discriminating these cells should help clean things up, but this won't change your results that much. I think in MLR it is very important as the stimulator cells and even the responder cells are likely to die at a high rate depending on the day of analysis.

(Edit: my figure didn't seem to attach, so if you look up the paper it is figure 3).

Gating on just the lymphocyte population doesn't change the peaks. They're virtually identical to the peaks I've shown you.
I'm going to go back to staining spleen cells and LN cells and double staining with PE-CD3 or something.

This is frustrating...I only want to hammer out this protocol, so that we can use it in the future for in vivo proliferation studies. Any ideas on how to go about this? I really appreciate your help.

I must admit I'm also running low on bright ideas.

Try staining spleen or LN cells as you stated and do an in vitro proliferation assay and stain for dead cells when you analyze. At this point I would avoid the cells lines if that is not what you want to do in the end anyway.

Although your initial CFSE staining looked OK, is there any chance your CFSE is old? We keep our aliquots at -20°C and with desiccant.

edit: I meant to say in vitro, not in vivo. Also, when you test this on splenocytes, culture some without stimulus. Unstimulated T cells should survive a number of days and stay CFSE bright.

I have not performed in vivo proliferation assays, although it sound relatively straight forward once you get it all working in vitro. I remember a paper being posted on these boards some time ago regarding the technique, but your probably aware of all the relevant papers before you started the project.

I have noticed that the amount of ConA you are using is 5microgramms per ml, we have found that 1 microgramm per ml is enough. And also it might help to vortex the cells 2 times during those 9 minutes labeling time. But these are only minor "cosmetics" but it might help.

Gibacht

actually, the 5 ug v/s 1 ug may not just be 'cosmetic' - 5 ug/ml Con A actually shows greater cell death than 1 ug/ml for primary cells (JLeuk Biol,2002,2005).
Your protocol is very similar to what I use, except mine is 2 uM final, and culture in 96-well plates at 100kcells/well. Your cells do seem to stain well initially, so no problem there. (I'm not very surprised if a cell line shows no/low 0 div peak)
Try reducing ConA conc or using ViViD Red/PI or other live v/s dead discriminitor.
Ensure you have a DDM module on (FSC-A v/s H),before FSC-A v/s SSC.
Your primary culture optimal profile should look like this (~70 h activation), with a peak ratio ~0.5. The fig is for just the basic DDM-gated cells, with no other SSC/live-dead gates applied.

Thanks, I'll try the 1ug/ml Con-A on splenoycytes and see if it helps.

Okay, so here are some plots from Day 4 of the CFSE stain at 1.25uM. I also lowered the con-A concentration to 1ug/ml. Still not pretty.

Having a lot of trouble attaching images...no formats work for me.

images

If I see it right, you seem to have picked the wrong gate from your FSC-SSC plot - its straddling the lo-high FSC population.
I don't know which cytometer and program you are using, but if it supports it, try acquiring/analysing cells in this order:
Set threshold to ~5000
Gate cells on the diagonal in the FSC-A/FSC-H plot (DDM) - use as count/collection gate, but acquire ALL cells
Gate the lymphocyte population from FSC-A/SSC-A
Apply compensation matrix.
Gate CD3+ cells
Apply CFSE analysis.

How is your compensation matrix like? It appears as if you do not have the proper compensation set up for spillover from FITC to PE.
CFSE is a rather terrible dye in that respect - it spills over in PE, PerCP-Cy5 (FL2, FL-3) to a considerable degree. Once that is straightened out, you should be able to see the division peaks in your CD3+ lymphocyte population much more clearly. Also, you will need to use a curve-fitting program - most times it cannot be done by eyeballing it - see the example I'd posted earlier.

The red population shown in PE vs FITC is the smaller gate from the scatter plot. I think the compensation is fairly good (PE and FITC is fairly distinct and no weird curvature is there.) I do admit that the FSC/SSC plot looks odd, but I seem to get that when using Con-A in cultures. I did do the FSC-H/FSC-A but it is not shown.