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 without Ca²⁺ and Mg²⁺ at 37°C
• Trypsin/EDTA solution
• Complete medium with serum with 10% to 15% (v/v) FBS
• Sterile Pasteur pipettes
• 37°C warming tray or incubator
• Tissue culture plasticware or glassware, including pipettes and 25-cm² flasks or 60-mm petri plates, sterile

Procedure

1. Remove all medium from primary culture with a sterile Pasteur pipette. Wash the 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 the bottom of the 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 2.
4. Add 2 mL 37°C complete medium. Draw cell suspension into a Pasteur pipette and rinse cell layer 2 or 3-times to dissociate cells and 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 hemocytometer 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 stimulate the in vivo environment of cells and 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 becomes confluent by repeating steps 1 to 7, and continue to passage as necessary.