I am trying to count the numbers/percentages of lymphatic endothelial cells in different mouse adipose tissue depots. These cells are theoretically CD31+CD45-Lyve-1+Podoplanin+, with the expression of the last two markers (Lyve-1 and podoplanin) varying across LECs of different origins. My problem is the difficulty in setting gates and “finding” the cells of interest (which are only a few), due to the “dim” staining of both Lyve-1 and podoplanin and due to the autofluorescence as well. The dim staining (population shift and no clear + and – populations) was also seen using a commercially available LEC cell line for FACS analysis.

I am using a usual collagenase/70μm cell strainer protocol to prepare the single-cell suspension. Fc block is being used. Staining buffer contains 1% FBS as well. Please find the example graphs attached. Take into account that the Alexa488- and Podoplanin-isotypes are single stained (that mean there has been no gating on CD31+, I know it was not the best thing to do…)

For this experiment Podoplanin-Alexa488, Lyve-1-PE, CD31-APC and 7-AAD were used. I have checked for unspecific binding using isotype controls and this is not my problem (MFI of cells stained with isotype control is the same with the MFI of the internal negative control, i.e. the negative cells in a single stained sample). I have tried setting my negative/positive gates using the isotype controls (although I am aware of the whole discussion in the literature that this is not correct), but this made of course no difference anyway, as I have no significant unspecific binding.

My plan is to try with FMO controls, to improve gating, and CD45, in order to gate on non-leukocytes, and improve my compensation using beads, although I am not sure that it will make a huge difference.