In cell culture, passaging is the process of sub-culturing cells. It is usually done to produce large numbers of cells from pre-existing ones. Instances where it is followed include vaccine production labs and clonal expansion.
Adherent (sticky) mammalian cells are typically grown in a petri dish (a plate) or tissue culture flask, with growth media (e.g. FBS + DMEM), in an incubator at 37°C with 5% CO2 and a tray of water in the bottom for humidity. In the case of RAW 264.7 or HeLa cells, a 10%-full (10% confluent) plate will reach 100% confluency in two or three days. If no evasive action is taken, the nutrients will be depleted and the cells will die shortly thereafter, thus necessitating passaging in order to maintain the culture. During passaging, the growth media is removed, and the cells may be washed with phosphate buffered saline (PBS) (salt water), followed by the addition of trypsin to detach the cells from the bottom of the plate. Trypsin functions optimally at 37°C, so the plate is incubated for five minutes or more. Care is taken not to overexpose cells to trypsin, as it can damage important cell surface proteins, channels and receptors. The plate is removed from the incubator and PBS or growth medium may be added (the proteins present in growth medium essentially absorb the action of trypsin which is ultimately deactivated through natural degeneration) and the cells and medium are mixed with a pipettor (triturated). An appropriate number of cells in suspension is then transferred to new plates, fresh medium is added to each plate, the new plates are put in the incubator, and the cycle begins again.
For maximum yield, cells are kept less than 100% (log phase of growth) but more than 10% confluent. Cells may die if they are too few or much too crowded.