Hello all,

I’m working on a project that requires the isolation of cells from mouse joint tissue to be used later on in flow cytometry. So far we’ve been able to get a few cells, but are having an issue obtaining any live ones. Does anyone have any suggestions for a protocol to follow, or any papers to read?

If helpful, our current protocol is as follows:

· Separate joint tissue from bone in small sterile cell culture dish by teasing the tissue away from the bone using 20-gauge syringe needles and dissecting microscope

· Digest the tissue by incubation in 5 ml DMEM culture media containing 0.2mg/ml collagenase and 100ug/ml DNase I

· Incubate cells, DMEM, Liberase, DNase for 1hr at 37C

· After incubation, disrupt any intact tissue by gently pipetting using the end of a 5-mL syringe or with a pipette (sucking up and down)

· Filter the cells through a 100 um cell strainer

· Pellet the cells through centrifugation (1000rpm, 10minutes)

· Discard media

· Wash the cells with 10mL of Cold 1XPBS.

· Centrifuge at 1000 RPM for 10 minutes

· Discard spent PBS

· Resuspend the pellet in 2mLs 1XPBS

· Count the cells by using 10uL Trypan blue + 10uL cell suspension (add 10uL of mix to cell counter).

We’ve tried changing filter size from 100 µm to 70 µm, and skipping the RBC lysis buffer, and it increased the number of cells obtained, but did not improve viability. Would decreasing the incubation time help, or perhaps pool joints from several mice together? Currently we have just been using joints from single mice until we have the protocol figured out.

Thanks!