Cell culture passaging protocol describes a general procedure for adherent cell lines and suspension mammalian cells in culture.
Definition of Subculture
Subculturing in tissue culture, also known as “cell splitting,” is required to offer fresh nutrients and growing space to continually expanding cell lines on a regular basis. The frequency of subculturing in tissue culture and the split ratio, or cell density plated, will be determined by the properties of each cell line. The line may be lost if cells are broken too frequently or at too low a density.
If cells are not divided frequently enough, they may exhaust the media and die, or in mixed cell cultures, a different type of cell may be selected. In general, once the proper timing and split ratio for a cell line has been determined, it should be utilised consistently for that line, with slight modifications only when absolutely necessary. subculturing in tissue culture techniques can be used to select for certain qualities in the cell lines transported. Changing the culturing routine could cause the cells and the scientist to become confused. Subculturing in tissue culture entails removing the growth media, cleaning the plate, dissociating the adherent cells enzymatically (e.g., with trypsin), and diluting the cell suspension into fresh media.
It is usually not essential to eliminate the leftover enzyme when using serum and the split ratio is large (e.g., 1:100). If the culture is kept in serum-free media, however, a suitable protease inhibitor, such as soybean trypsin inhibitor, must be used to neutralise the enzyme. Other considerations necessitate the adoption of alternative procedures. Even if serum is present, it is recommended to wash the cells following enzyme treatment if the cell line is carried at a very low split ratio. To prevent breaking the monolayer, all solutions added to dishes/flasks with adherent cells should be pipetted down the side of the dish or flask.
Adherent Cell Subculture
Figure 1: Subculturing Adherent Cells
- Assess the degree of confluency and validate the absence of bacterial and fungal contamination using an inverted microscope.
- Remove the spent media and wash the cell lines monolayer with PBS that is free of Ca2+ and Mg2+. If the cells are known to attach aggressively, repeat the wash step (Figure 1).
- Pipette roughly 1 ml of trypsin/EDTA per 25 cm2 of surface area onto the cleaned cell monolayer. To cover the monolayer with trypsin, rotate the flask. Return the flask to the incubator for 2-10 minutes after emptying the excess trypsin.
- Using an inverted microscope, inspect the cells to confirm that they are all unattached and floating. To liberate any remaining adhered cells, lightly tap the flasks on the side.
- To inactivate the trypsin, resuspend the cells in a tiny volume of new serum-containing media. Remove 100-200 L and count the cells. Use a trypsin inhibitor, such as soybean trypsin inhibitor, to inactivate trypsin in cells cultivated in serum-free medium.
- Transfer the required number of cells to a new labelled flask containing pre-warmed media and incubate according to the cell line’s instructions.
Suspension Cell Subculture
Cell cultures in flasks or spinners that develop in suspension can be maintained by diluting an aliquot of the suspension into fresh growth media.
- Flasks (75 cm2)
- Growth medium
- Using an inverted phase contrast microscope, examine cultures. Bright, spherical, and refractile cells should be seen in the exponential growth phase.
- Hybridomas can be quite sticky, and detaching the cells requires a moderate knock on the flask. EBV-transformed cells can form huge clumps that are difficult to count, and the large clumps’ centres may be non-viable.
- Take a tiny sample of the cells (100-200L) from the cell suspension and count them. Calculate cells/mL and re-seed the necessary number of cells into freshly prepared flasks by diluting the cells without centrifugation.