For some cell cultures, especially valuable ones, it is common practice to maintain a two-tiered frozen cell bank: a master and a working cell bank. The working cell bank comprises cells from one of the master bank samples, which have been grown for several passages before storage. If and when future cell samples are needed, they are taken from the working cell bank. The master cell bank is used only when absolutely necessary. This ensures that a stock of cells with a low passage number is maintained, and avoids genetic variation within the culture.
Check that cells are healthy, not contaminated, and have the correct morphology.
Change the medium 24 h before freezing the cells.
Adherent and suspension cell cultures should not be at a high density for freezing. We recommend freezing cells when they are in the logarithmic growth phase.
Adherent cultures: harvest the cells by trypsinization, resuspend in medium containing serum, centrifuge at 200 x g for 5 min, and then resuspend cells in freezing medium (see table Freezing medium) at a density of 3–5 x 106 cells/ml.
Suspension cultures: centrifuge the cells at 200 x g for 5 min, and resuspend in freezing medium at a density of 5–10 x 106 cells/ml.
IMPORTANT: Freezing medium containing DMSO is hazardous and should be handled with caution.
Transfer 1 ml of the cell suspension (approximately 3–5 x 106 adherent cells or 5–10 x 106 suspension cells) into each freezing vial. Label vials with the name of cell line, date, passage number, and growth medium.
Tip: It may also be useful to note the cell density in the freezing vials before storing. This enables determination of the cell density that provides optimal recovery after thawing.
Place freezing vials in racks and transfer to a polystyrene box (with walls approximately 15 mm thick) lined with cotton wool. Store box in a –80°C freezer overnight.
The next day, quickly transfer the vials to a liquid nitrogen chamber, making sure that the vials do not begin to thaw.
Growth medium (RPMI, DMEM, etc.) containing 10–20% FBS and 5–20% glycerol or DMSO
Trypan blue staining provides a method for distinguishing between viable (i.e., capable of growth) and nonviable cells in a culture. This staining method is based on “dye exclusion”: cells with intact membranes exclude (i.e., do not take up) the dye and are considered viable.
Harvest the cells, either by trypsinization (adherent cell cultures) or by centrifugation at 200 x g for 5 min (suspension cell cultures). Resuspend the cells in an appropriate volume of pre-warmed growth medium to give a cell density of at least 106 cells/ml.
Add 0.5 ml 0.4% (w/v) trypan blue (see table Trypan blue) and 0.3 ml PBS or Hank’s balanced salt solution (HBSS; see tables 1x PBS and 1x HBSS) to 0.1 ml of the cell suspension. Mix thoroughly, and let stand for 1–2 min. Alternatively, add 0.4 ml trypan blue directly to 0.4 ml of cells in growth medium.
At least 106 cells/ml are required for accurate counting.
Count the stained and unstained cells using a hemocytometer (see Cell counting using a hemocytometer). Blue-stained cells are nonviable and unstained cells are viable.
No. of viable cells/ Total no. of cells = % viability