Subculturing Adherent Cells
For passaging your own cell line, we recommend that you closely follow the instructions provided with each product you are using in your experiments. The consequences of deviating from the culture conditions required for a particular cell type can range from the expression of aberrant phenotypes to a complete failure of the cell culture.
- Culture vessels containing your adherent cells
- Tissue-culture treated flasks, plates or dishes
- Complete growth medium, pre-warmed to 37°C
- Disposable, sterile 15-mL tubes
- 37°C incubator with humidified atmosphere of 5% CO2
- Balanced salt solution such as Dulbecco’s Phosphate Buffered Saline (DPBS), containing no calcium, magnesium, or phenol red
- Dissociation reagent such as trypsin or TrypLE™ Express, without phenol red
- Reagents and equipment to determine viable and total cell counts such as Countess® Automated Cell Counter, Trypan Blue and hemacytometer, or Coulter Counter® (Beckman Coulter)
|This video explains why, when and how to passage cells grown in both adherent and suspension cultures. This includes cell dissociation, counting cells, determining optimal seeding density and preparing new culture vessels for passaged cells.|
All solutions and equipment that come in contact with the cells must be sterile. Always use proper sterile technique and work in a laminar flow hood.
- Remove and discard the spent cell culture media from the culture vessel.
- Wash cells using a balanced salt solution without calcium and magnesium (approximately 2 mL per 10 cm2 culture surface area). Gently add wash solution to the side of the vessel opposite the attached cell layer to avoid disturbing the cell layer, and rock the vessel back and forth several times.
Note: The wash step removes any traces of serum, calcium, and magnesium that would inhibit the action of the dissociation reagent.
- Remove and discard the wash solution from the culture vessel
- Add the pre-warmed dissociation reagent such as trypsin or TrypLE™ to the side of the flask; use enough reagent to cover the cell layer (approximately 0.5 mL per 10 cm2). Gently rock the container to get complete coverage of the cell layer.
- Incubate the culture vessel at room temperature for approximately 2 minutes. Note that the actual incubation time varies with the cell line used.
- Observe the cells under the microscope for detachment. If cells are less than 90% detached, increase the incubation time a few more minutes, checking for dissociation every 30 seconds. You may also tap the vessel to expedite cell detachment.
- When ≥ 90% of the cells have detached, tilt the vessel for a minimal length of time to allow the cells to drain. Add the equivalent of 2 volumes (twice the volume used for the dissociation reagent) of pre-warmed complete growth medium. Disperse the medium by pipetting over the cell layer surface several times.
- Transfer the cells to a 15-mL conical tube and centrifuge then at 200 × g for 5 to 10 minutes. Note that the centrifuge speed and time vary based on the cell type.
- Resuspend the cell pellet in a minimal volume of pre-warmed complete growth medium and remove a sample for counting.
- Determine the total number of cells and percent viability using a hemacytometer, cell counter and Trypan Blue exclusion, or the Countess® Automated Cell Counter. If necessary, add growth media to the cells to achieve the desired cell concentration and recount the cells.
Note: We recommend using the Countess® Automated Cell Counter to determine the total number of cells and percent viability. Using the same amount of sample that you currently use with the hemacytometer, the Countess® Automated Cell Counter takes less than a minute per sample for a typical cell count and is compatible with a wide variety of eukaryotic cells. See the protocol on Counting Cells with a Hemacytometer.
- Dilute cell suspension to the seeding density recommended for the cell line, and pipet the appropriate volume into new cell culture vessels, and return the cells to the incubator.
Note: If using culture flasks, loosen the caps before placing them in the incubator to allow proper gas exchange unless you are using vented flasks with gas-permeable caps.
- Passage insect cells at log phase. However, if your insect cells are strongly adherent, you may passage them at confluency or slightly after when they are starting to pull away from the bottom of the flask because they will be easier to dislodge.
- Densities lower than 20% confluency inhibit growth. The healthiest cells are those taken from log phase cultures.
- CO2 exchange is not recommended for insect cell culture.
- Maintain insect cells at 27°C in a non-humidified environment. Cells can be maintained at room temperature on the bench top if protected from light or in a drawer. However, a 27°C controlled environment is recommended.
- Use media specifically formulated for insect cell growth.
- Insect cells attach very tightly to substrates under serum-free conditions and require additional effort to detach. To dislodge the cells, you may need to give the flask one quick shake using a wrist-snapping motion. To avoid contamination, always tighten the cap before this procedure.
Caution: We do not recommend shaking the flask vigorously, because it may result in damage to the cells.