To begin, prepare 30 microliters of staining buffer at 2x working concentrations for single color and FMO controls. Transfer the single color and FMO control staining buffer to the wells of a 96 well plate. Top up the volume with 20 microliters of FACS buffer.
To stain the sample, prepare 0.5 milliliters of 2x FACS staining buffer in 1.5 milliliter centrifuge tubes. Pipette 10 microliters of the cell suspension to single color and FMO control wells. Then, add 2x FACS staining buffer to the rest of the cell suspension and incubate.
Next, add 100 microliters of FACS buffer into the wells and two milliliters of the buffer into the sample tubes. Centrifuge at 500G for five minutes at four degrees Celsius. Aspirate the supernatant, then resuspend the cell pellet in viability buffer.
Add 300 microliters of proliferation media into 1.5 milliliter centrifuge tubes. These will act as collection tubes for cell sorting. After sorting the cells using FACS, centrifuge the collected cells at 800G for 10 minutes at four degrees Celsius.
Use a P1000 pipette to carefully remove the supernatant without disturbing the pellet. Resuspend the pellet in proliferation media to a final density of 100 cells per microliter. Seed the cell suspension into the wells of a 48 well tissue culture plate.
Finally transfer the plate to an incubator at 37 degrees Celsius under respective gas supplementation until cells are confluent. Confluent cells with typical fibroblast morphology were obtained within five days of culture.
