Cell Culture Techniques: How to Grow Cells in the Laboratory
Fundamentals of Cell Culture
Cells in culture require conditions that closely mimic their natural environment. The basic requirements include a sterile growth surface, a nutrient-rich liquid medium, a controlled atmosphere (typically 37 degrees Celsius, 95% humidity, and 5% carbon dioxide), and protection from microbial contamination. The CO2 is necessary because most culture media use a bicarbonate buffering system in which dissolved CO2 combines with sodium bicarbonate to maintain the pH at approximately 7.4. A phenol red indicator in the medium changes color to signal pH shifts: red at the optimal pH, yellow when too acidic (indicating metabolic waste accumulation or CO2 excess), and purple when too alkaline.
Cell cultures fall into two broad categories. Primary cultures are established directly from tissue samples and contain cells that closely resemble their in vivo counterparts. However, most primary cells have a limited lifespan in culture (the Hayflick limit, typically 40 to 60 population doublings for human fibroblasts) before they enter senescence and stop dividing. Cell lines are populations of cells that have acquired the ability to proliferate indefinitely, either through spontaneous mutation or deliberate immortalization. The HeLa cell line, derived from the cervical cancer cells of Henrietta Lacks in 1951, was the first human cell line established and remains one of the most widely used in research, having contributed to discoveries ranging from the polio vaccine to the molecular biology of cancer.
How to Culture Cells: Step by Step
Step 1: Prepare the Biosafety Cabinet and Materials
All cell culture work must be performed in a biosafety cabinet (BSC) to maintain sterility. Turn on the BSC at least 15 minutes before use to establish laminar airflow, then wipe down all interior surfaces and any items entering the cabinet with 70% ethanol. Gather all necessary materials, including pre-warmed media, sterile serological pipettes, culture vessels, and waste containers, before beginning work. Minimize hand movements in and out of the cabinet to avoid disrupting the protective airflow curtain. Wear gloves and a lab coat, and spray gloves with ethanol before reaching into the cabinet.
Step 2: Prepare Growth Medium
Growth medium provides the nutrients, hormones, and growth factors that cells need to survive and proliferate. Most mammalian cell culture uses a basal medium such as Dulbecco Modified Eagle Medium (DMEM) or RPMI 1640, supplemented with fetal bovine serum (FBS) at concentrations typically ranging from 5 to 20 percent. FBS provides growth factors, hormones, attachment factors, and lipids that are essential for most cell types. Additional supplements may include L-glutamine (an amino acid consumed rapidly in culture), non-essential amino acids, and antibiotics such as penicillin and streptomycin, though many researchers avoid routine antibiotic use because it can mask low-level contamination and promote antibiotic-resistant organisms.
Warm complete medium to 37 degrees Celsius in a water bath before adding it to cells. Cold medium can shock cells and reduce viability. Check that the medium color is the correct shade of red/orange (indicating proper pH) before use, and never use medium that has turned yellow (acidic) or purple (alkaline) without investigation.
Step 3: Seed Cells into Culture Vessels
When starting a culture from a cryopreserved stock, thaw the cryovial rapidly in a 37-degree water bath, transfer the contents to a tube of pre-warmed medium, and centrifuge gently to pellet the cells and remove the DMSO-containing freezing medium. Resuspend the pellet in fresh growth medium and count the cells using a hemocytometer or automated cell counter, mixing the cell suspension with trypan blue to distinguish live cells (which exclude the dye) from dead cells (which take it up). Seed cells at the density recommended for the specific cell type, typically between 5,000 and 50,000 cells per square centimeter for adherent cells. Place the culture vessel in a humidified incubator set at 37 degrees Celsius with 5% CO2.
Step 4: Monitor and Feed the Cultures
Examine cultures daily using an inverted phase-contrast microscope. Assess cell morphology (cells should appear healthy, with smooth membranes and typical shapes for their type), growth rate (cells should be increasing in number), and any signs of contamination (turbidity in the medium, floating debris, or unusual pH changes). Replace spent medium with fresh pre-warmed medium every 2 to 3 days, or more frequently for rapidly growing cell lines. When changing medium, aspirate the old medium carefully to avoid disturbing the cell layer, then gently add fresh medium along the wall of the flask to minimize shear stress on the cells.
Step 5: Passage Cells at Confluence
Adherent cells must be passaged (subcultured) when they approach confluence, the point at which they cover the entire growth surface. Most cell types should be passaged at 70 to 90 percent confluence; allowing cells to become over-confluent can alter their growth properties, gene expression, and responsiveness to stimuli. To passage, remove the old medium, wash cells gently with phosphate-buffered saline (PBS) to remove residual serum (which inhibits trypsin), add a thin layer of trypsin-EDTA solution (typically 0.05 to 0.25 percent trypsin), and incubate at 37 degrees for 2 to 5 minutes until cells detach. Monitor detachment under the microscope to avoid over-trypsinization, which can damage cell surface proteins. Add serum-containing medium to neutralize the trypsin, pipette to create a single-cell suspension, and reseed at a lower density into new flasks. A typical split ratio ranges from 1:3 to 1:10 depending on the cell growth rate.
Step 6: Cryopreserve Stocks
Maintaining frozen stocks of cells at early passage numbers is essential for experimental reproducibility and as insurance against contamination or incubator failures. To cryopreserve, harvest cells by trypsinization, count them, and resuspend at 1 to 5 million cells per milliliter in freezing medium consisting of 90% FBS and 10% dimethyl sulfoxide (DMSO). DMSO is a cryoprotectant that prevents ice crystal formation during freezing, which would otherwise rupture cell membranes. Aliquot the suspension into labeled cryovials, place the vials in a controlled-rate cooling device (such as a Mr. Frosty container filled with isopropanol) that cools at approximately negative 1 degree per minute, and store overnight at negative 80 degrees Celsius. For long-term storage, transfer vials to liquid nitrogen (negative 196 degrees Celsius), where cells can remain viable for decades.
Contamination Prevention and Detection
Contamination is the most common reason cell cultures fail. Bacterial and fungal contamination usually produces visible signs within a few days: turbid medium, pH changes, floating colonies, or filamentous growth. Mycoplasma contamination, caused by tiny bacteria that lack cell walls, is far more insidious because it produces no visible changes in the culture but can profoundly affect cell behavior, gene expression, growth rate, and experimental results. Studies have found that 15 to 35 percent of cell lines in research laboratories are contaminated with mycoplasma, making routine testing essential. PCR-based detection kits and commercial fluorescent staining methods (using DAPI or Hoechst stains that reveal mycoplasma DNA on the cell surface) are the most reliable detection methods.
Prevention is always preferable to treatment. Strict aseptic technique, including working in a properly maintained biosafety cabinet, wearing gloves, avoiding mouth pipetting, and never sharing media bottles between cell lines, is the foundation of contamination prevention. Quarantining new cell lines for 2 to 4 weeks and testing for mycoplasma before introducing them to the general culture area is a standard precaution in well-managed cell culture facilities.
Advanced Culture Systems
Traditional two-dimensional cell culture on flat plastic surfaces, while technically simple and widely used, fails to replicate many features of the three-dimensional tissue environment in which cells normally reside. Cells grown on flat surfaces adopt unnatural morphologies, gene expression patterns, and drug sensitivities that can limit the relevance of experimental findings. Three-dimensional culture systems, including scaffold-based cultures, spheroid cultures, and organoids, address these limitations by allowing cells to interact with each other and with extracellular matrix components in architectures that more closely resemble real tissues.
Organoids, self-organizing three-dimensional structures grown from stem cells or tissue progenitor cells, represent the most sophisticated current approach to recapitulating tissue architecture in vitro. Intestinal organoids, first developed by Hans Clevers and colleagues in 2009, form crypt-villus structures that closely mirror the organization of the real intestinal epithelium. Brain organoids, kidney organoids, and liver organoids have since been developed, providing powerful models for studying development, disease, and drug responses in contexts that two-dimensional cultures cannot replicate.
Successful cell culture depends on rigorous aseptic technique, appropriate growth conditions, regular monitoring, and proper passage and cryopreservation protocols, with advanced 3D systems and organoids increasingly bridging the gap between in vitro experiments and in vivo biology.