Cell passaging, also known as subculturing, is a fundamental and routine technique in cell biology laboratories. It is the process of transferring cells from a crowded “parent” culture vessel to a new vessel containing fresh growth medium. This procedure maintains the cells in an optimal growth state, allowing the population to continue proliferating exponentially. By diluting the cell suspension and providing a renewed environment, passaging prevents the culture from entering a non-growing state and dying.
Why Cell Cultures Require Subculturing
Cell growth in a culture vessel is limited by finite resources and space. As cells divide, they rapidly consume nutrients, such as glucose and amino acids, dissolved in the culture medium. Depletion of these resources means the environment can no longer support sustained cell proliferation.
Cells also produce metabolic byproducts, such as lactic acid, which accumulate in the medium. This accumulation of waste products lowers the medium’s pH, creating a toxic environment that inhibits growth and can lead to cell death. Without intervention, the culture enters the stationary phase, ceasing growth and jeopardizing the cell line’s viability.
For adherent cells, the physical limit is the vessel’s surface area. Once these cells cover the entire substrate, they exhibit a phenomenon called contact inhibition, where cell-to-cell contact signals them to stop dividing. Passaging a culture before it reaches maximum density, typically 70% to 90% confluency, is necessary to keep the cells in the exponential growth phase.
Key Steps in the Passaging Procedure
The methodology for passaging differs based on whether the cells are adherent or suspension cells. Adherent cells must first be detached from the culture vessel surface before transfer. This process begins by removing the spent growth medium and washing the cell monolayer with a balanced salt solution, such as phosphate-buffered saline (PBS), to remove residual medium components that could interfere with the detachment agent.
A dissociation enzyme, most commonly trypsin, is then added to the flask and incubated briefly at 37°C. Trypsin is a protease that breaks down the proteins anchoring the cells to the plastic surface. Once the cells appear rounded and loosely detached, fresh culture medium containing serum is added. The serum neutralizes the trypsin activity, preventing the enzyme from damaging the cells.
The resulting cell suspension is collected into a centrifuge tube. The cells are pelleted by low-speed centrifugation, typically at 150–300 xg for a few minutes. This step concentrates the cells, allowing the researcher to aspirate and discard the old medium and residual enzyme. The cell pellet is then gently resuspended in a controlled volume of fresh medium, creating a single-cell suspension ready for counting and seeding into new vessels.
Suspension cells, which grow floating in the medium, do not require an enzymatic detachment step. Passaging these cultures involves collecting the cell suspension, centrifuging it to form a pellet, and then resuspending the cells in a calculated volume of fresh medium for dilution. This straightforward dilution process results in little to no growth lag phase when seeded into the new flask, unlike the trauma associated with the proteolytic enzyme dispersal needed for adherent cells.
Calculating Split Ratios and Cell Density
Before seeding new culture vessels, it is necessary to quantify the cells in the suspension to ensure the new culture starts at an optimal density. This is done by taking a small sample of the cell suspension and counting the number of viable cells using an instrument like a hemocytometer or an automated cell counter. This measurement yields the current cell concentration in cells per milliliter (cells/mL).
The next step involves calculating the appropriate split ratio to maintain the culture. A split ratio, such as 1:5 or 1:10, represents the factor by which the cell population is diluted into the new culture vessel. A 1:5 split, for example, means that one part of the cell suspension is added to four parts of fresh medium, effectively diluting the cell concentration by five times.
This ratio is chosen based on the cell line’s doubling time, which is the amount of time it takes for the population to double in number. A faster-growing cell line with a short doubling time will require a higher split ratio (e.g., 1:10) to prevent it from reaching confluency too quickly, while a slower-growing line may only need a 1:3 split. The goal is to select a ratio that ensures the cells will be ready for the next passage in a predictable timeframe, often every three to four days, while remaining in the exponential growth phase.
Maintaining Cell Line Integrity
Successfully maintaining a cell line requires rigorous attention to quality control. Aseptic technique is paramount during all passaging steps to minimize the risk of microbial contamination from bacteria, fungi, or mycoplasma. Working within a sterile environment, such as a biosafety cabinet, and sterilizing all reagents and tools are standard practices to protect the culture from foreign organisms.
Continuous passaging over an extended period can lead to genetic drift, which is the accumulation of subtle changes in the cell line’s phenotype or genotype. To mitigate this gradual change, researchers meticulously track the passage number, which is the number of times the cells have been subcultured. It is recommended to use cells with a low passage number, often below 10 splitting cycles, for experiments to ensure consistent and reliable results.
To ensure a continuous supply of genetically consistent, low-passage cells, cryopreservation is routinely performed. This involves freezing a large number of healthy cells in a cryoprotectant solution, such as dimethyl sulfoxide (DMSO), and storing them long-term in liquid nitrogen at -196°C. This creates a cell bank, which serves as a stable, viable backup stock that can be thawed when needed, safeguarding against contamination and genetic variation.