I recently cultured some HeLa cells as practice as I am relatively new in the lab.
Day 1 (Friday): Started with fast-thawing cells in a water bath (37 deg Cel). Pipette cells in to a 15-cm culture dish of 28 mL DMEM culture medium. Left to incubate over the weekend.
Day 4 (Monday): This was what I observed . Aspirated the cell suspension. Wash with PBS and then trypsinize (coupled with incubation 37 deg Cel) followed by neutralizing with DMEM. Spin down 1.2k rpm for 4 min in swing bucket centrifuge. Followed by transferring to a 10-cm culture dish containing 6 mL DMEM. Allowed to incubate overnight.
DMEM used was consistent.
Would like to know if I am on the right track? I deviated quite a lot from the standard protocol especially when it comes to the amount of PBS, trypsin and DMEM used when transferring the cells from 15-cm dish to 10-cm dish. I used 10 mL PBS to wash the cells in the 15-cm dish before aspirating it and adding 8 mL of trypsin. Subsequently, I used 10 mL of DMEM for neutralization. Attaining a total of 28 mL in the 15-cm dish. I then pipetted half, 14 mL into a 15 mL Falcon tube for centrifugation to pellet the cells before discarding the supernatant and then later transferred cells into a 10-cm dish of 6 mL of DMEM.