TRYPSINIZING AND SUBCULTURING CELLS FROM A MONOLAYER

A primary culture is grown to confluency in a 60-mm petri plate or 25-cm2 tissue culture
flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and
then reseeded into secondary cultures. The process of removing cells from the primary
culture and transferring them to secondary cultures constitutes a passage, or subculture.
Materials
Primary cultures of cells
HBSS (APPENDIX 2A) without Ca2+ and Mg2+, 37°C
Trypsin/EDTA solution (see recipe), 37°C
Complete medium with serum: e.g., supplemented DMEM (APPENDIX 2A) with 10%
to 15% (v/v) FBS (complete DMEM-10 or -15), 37°C

Sterile Pasteur pipets
37°C warming tray or incubator
Tissue culture plasticware or glassware including pipets and 25-cm2 flasks
or 60-mm petri plates, sterile
NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2
incubator unless otherwise specified. Some media (e.g., DMEM) may require altered
levels of CO2 to maintain pH 7.4.
1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering
cell monolayer once or twice with a small volume of 37°C HBSS without Ca2+ and
Mg2+ to remove any residual FBS that may inhibit the action of trypsin.
Use a buffered salt solution that is Ca2+ and Mg2+ free to wash cells. Ca2+ and Mg2+ in the
salt solution can cause cells to stick together.
If this is the first medium change, rather than discarding medium that is removed from
primary culture, put it into a fresh dish or flask. The medium contains unattached cells that
may attach and grow, thereby providing a backup culture.
2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer.
3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop
to dislodge cells. Check culture with an inverted microscope to be sure that cells are
rounded up and detached from the surface.
If cells are not sufficiently detached, return plate to warming tray for an additional minute
or two.
4. Add 2 ml 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse
cell layer two or three times to dissociate cells and to dislodge any remaining adherent
cells. As soon as cells are detached, add serum or medium containing serum to inhibit
further trypsin activity that might damage cells.
If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml
of cell suspension can be transferred into each fresh culture vessel.
5. Add an equal volume of cell suspension to fresh plates or flasks that have been
appropriately labeled.
Alternatively, cells can be counted using a hemacytometer or Coulter counter and diluted
to the desired density so a specific number of cells can be added to each culture vessel. A
final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures.
For primary cultures and early subcultures, 60-mm petri plates or 25-cm2 flasks are generally
used; larger vessels (e.g., 150-mm plates or 75-cm2 flasks) may be used for later subcultures.
Cultures should be labeled with date of subculture and passage number.
6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2
incubator.
If using 75-cm2 culture flasks, add 9 ml medium per flask.
Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is
thought to simulate the in vivo environment of cells and to enhance cell growth.
For some media it is necessary to adjust the CO2 to a higher or lower level to maintain the
pH at 7.4.
7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium
and adding fresh 37°C medium.
8. Passage secondary culture when it bcomes confluent by repeating steps 1 to 7, and
continue to passage as necessary.
Current