Cell Culture Basics: An Introduction to Growing Cells
By Dr. Zubair Khalid, DVM, MS, PhD ·
Cell culture is the process of removing cells from an animal or plant and growing them in an artificially controlled environment, typically a glass or plastic vessel containing a liquid nutrient medium. This technique allows researchers to study the physiology, biochemistry, and genetics of cells in isolation from the complex systemic influences of a whole organism. Since its development in the early twentieth century, cell culture has become a cornerstone of molecular biology, enabling breakthroughs in vaccine development, drug screening, cancer research, and regenerative medicine. The ability to maintain living cells outside the body, observe their behavior directly, and manipulate their environment with precision has transformed our understanding of life at the cellular level.
What Is Cell Culture?
At its core, cell culture involves the ex vivo maintenance and propagation of cells under controlled conditions of temperature, pH, nutrient supply, and sterility. The cells are provided with a culture medium—a liquid formulation containing amino acids, sugars, vitamins, inorganic salts, and often serum or defined growth factors—that mimics the extracellular fluid of the body. The culture vessel is placed in an incubator that maintains a constant temperature (typically 37°C for mammalian cells) and a humidified atmosphere containing 5% CO₂ to buffer the pH of the medium through the bicarbonate buffering system.
The importance of cell culture in research and medicine cannot be overstated. It provides a simplified, reproducible system for studying cellular processes such as signal transduction, gene expression, and apoptosis. It allows the production of biological molecules like monoclonal antibodies, hormones, and enzymes. It is essential for the safety testing of pharmaceuticals and cosmetics, and it underpins the development of cell-based therapies. Without cell culture, much of modern biotechnology and molecular medicine would simply not exist.
History of Cell Culture
The foundations of cell culture were laid in the late nineteenth and early twentieth centuries. In 1885, Wilhelm Roux maintained a piece of chick embryo neural plate in warm saline for several days, demonstrating that tissue could survive outside the organism. The true pioneer, however, was Ross Granville Harrison, who in 1907 cultured frog embryonic nerve fibers in a hanging drop of lymph, observing the outgrowth of axons. This experiment proved that cells could grow and differentiate in vitro and is widely regarded as the birth of cell culture.
The next major milestone came in 1912 when Alexis Carrel, a French surgeon, claimed to have maintained chick heart fibroblasts in culture for decades. Although his methods were later questioned—the long-term survival was likely due to the addition of fresh embryo extract containing new cells—Carrel's work established the principle that cells could be kept alive for extended periods. The development of antibiotics in the 1940s, particularly penicillin and streptomycin, made routine cell culture practical by controlling bacterial contamination. In 1952, George Gey established the HeLa cell line from a cervical carcinoma, the first immortalized human cell line, which remains a workhorse in laboratories worldwide. The subsequent development of defined media formulations, such as Eagle's Minimal Essential Medium in 1955, and the introduction of laminar flow biosafety cabinets in the 1960s, brought cell culture into the mainstream of biological research.
Why Grow Cells in the Lab?
Growing cells in the laboratory offers several distinct advantages over studying whole organisms. First, it provides a controlled environment: the researcher can precisely manipulate the chemical composition of the medium, the oxygen tension, the temperature, and the physical substrate on which cells grow. This allows the isolation of specific variables and the study of their effects in a way that is impossible in a whole animal, where homeostatic mechanisms constantly compensate for perturbations.
Second, cell culture is scalable. A researcher can start with a single vial of frozen cells and expand them into millions of cells for biochemical analysis, or scale up to industrial bioreactors for the production of therapeutic proteins. Third, cell culture reduces the need for animal experimentation, aligning with ethical considerations and the "3Rs" principle of Replacement, Reduction, and Refinement. Fourth, cultured cells can be genetically manipulated with relative ease, allowing the introduction of reporter genes, knockdown of specific transcripts, or expression of mutant proteins to study their function. Finally, cell culture enables high-throughput screening: thousands of compounds can be tested for their effects on cell viability, proliferation, or signaling in multi-well plates, a scale that is impossible in animal models.
Types of Cell Cultures
Cell cultures can be broadly classified based on their origin and lifespan. The two main categories are primary cultures, derived directly from tissue, and immortalized cell lines, which can proliferate indefinitely. A further distinction is made between adherent cultures, which grow attached to a surface, and suspension cultures, which grow freely in the liquid medium.
Primary vs. Immortalized Cells
A primary culture is established by dissociating cells from fresh tissue, typically using enzymatic digestion with trypsin or collagenase, followed by mechanical disaggregation. These cells closely resemble their in vivo counterparts in terms of gene expression and function, making them valuable for studies of normal physiology. However, primary cells have a finite lifespan: most mammalian cells can undergo only a limited number of divisions—typically 20 to 60 population doublings in culture—before they enter a state of irreversible growth arrest known as replicative senescence. This limit, first described by Leonard Hayflick in 1961, is due to the progressive shortening of telomeres, the protective caps at the ends of chromosomes, with each cell division.
Immortalized cell lines, in contrast, have acquired the ability to divide indefinitely. This can occur spontaneously, as in the case of HeLa cells, or through deliberate genetic manipulation, such as the introduction of the telomerase reverse transcriptase gene (hTERT) or viral oncogenes like the SV40 large T antigen. These cells bypass senescence and can be propagated indefinitely, providing a consistent and reproducible experimental system. However, immortalization is often accompanied by genetic and phenotypic drift, meaning that cell lines may no longer accurately represent the tissue from which they were derived. For example, many widely used cancer cell lines have accumulated numerous mutations and chromosomal abnormalities over decades of passage. For detailed guidance on handling these cells, see Primary Cell Culture Guidelines and the broader discussion of Primary and Secondary Cell Culture.
Adherent vs. Suspension Cultures
Most cells derived from solid tissues, such as fibroblasts, epithelial cells, and endothelial cells, are adherent: they require a surface to attach to and will not grow in free suspension. In the laboratory, they are cultured on treated plastic dishes or flasks, where the surface has been modified to carry a net negative charge, promoting cell attachment through electrostatic interactions and the deposition of extracellular matrix proteins such as fibronectin and collagen. Adherent cells spread out on the surface, adopt a flattened morphology, and divide until they form a confluent monolayer covering the dish.
Suspension cultures are used for cells that naturally grow in the bloodstream or lymphatic system, such as lymphocytes, or for cells that have been adapted to grow without attachment, such as many hybridoma and Chinese hamster ovary (CHO) cell lines used in biopharmaceutical production. Suspension cells grow as single cells or small clumps floating in the medium. They are easier to scale up than adherent cultures because they do not require a large surface area; they can be grown in spinner flasks or Mammalian Cell Culture Bioreactor systems with agitation and aeration. The choice between adherent and suspension culture has practical implications for passaging, transfection efficiency, and downstream processing.
Essential Equipment and Supplies
Establishing a cell culture laboratory requires specialized equipment designed to maintain sterility and provide a controlled environment. The basic infrastructure includes a biosafety cabinet, a CO₂ incubator, an inverted microscope, and a supply of sterile plasticware and reagents.
Biosafety Cabinets
The biosafety cabinet (BSC), also known as a laminar flow hood, is the most critical piece of equipment for maintaining sterility. It provides a sterile work area by filtering incoming air through a high-efficiency particulate air (HEPA) filter that removes 99.97% of particles larger than 0.3 micrometers, including bacteria, fungi, and dust. There are several classes of BSCs; for most cell culture work, a Class II, Type A2 cabinet is used. This type protects both the user and the culture by directing a portion of the filtered air downward over the work surface and exhausting the remainder through a second HEPA filter back into the laboratory.
The cabinet creates a sterile environment through unidirectional airflow, but it does not sterilize objects placed within it. All items must be sprayed with 70% ethanol before being placed in the hood, and the hood itself should be UV-irradiated for 15–30 minutes before use and wiped down with ethanol afterward. The operator must work at least 15 cm inside the front edge of the hood to avoid disrupting the airflow barrier, and movements should be slow and deliberate to prevent turbulence that could draw contaminants into the work area.
Incubators and CO2 Control
Mammalian cells require a constant temperature of 37°C and a humidified atmosphere containing 5% CO₂. The CO₂ incubator provides these conditions. The CO₂ is essential because it dissolves in the culture medium and reacts with water to form carbonic acid, which in turn dissociates to bicarbonate and hydrogen ions. This equilibrium maintains the pH of the medium at approximately 7.4, provided the medium contains sodium bicarbonate as a buffer. The incubator monitors the CO₂ concentration with an infrared sensor and injects CO₂ from a compressed gas cylinder as needed.
Humidity is maintained by a water pan at the bottom of the incubator, which typically holds sterile distilled water. High humidity prevents evaporation of the medium, which would otherwise concentrate salts and alter osmolarity. The incubator should be cleaned regularly with 70% ethanol or a copper-based disinfectant to prevent fungal and bacterial growth. Some incubators also offer options for oxygen control, which is important for hypoxic studies or for cells that require lower oxygen tension.
Other essential equipment includes an inverted microscope for observing cells in culture vessels, a hemocytometer or automated cell counter for determining cell numbers, a centrifuge for pelleting cells during medium changes, and a water bath for warming media to 37°C before use. All plasticware—flasks, dishes, pipettes, and tubes—must be sterile and certified for cell culture use, as must all media and reagents.
Cell Culture Media and Supplements
The culture medium is the lifeline of the cells, providing all the nutrients, growth factors, and environmental buffering they need to survive and proliferate. The formulation of the medium is critical, and different cell types have different requirements.
Basal Media
Basal media are defined formulations that provide the essential nutrients: amino acids, vitamins, inorganic salts, glucose, and a buffer. The most common basal media include Dulbecco's Modified Eagle's Medium (DMEM), RPMI-1640, and Minimal Essential Medium (MEM). DMEM, for example, contains 25 mM glucose, 4 mM L-glutamine, and 1 mM sodium pyruvate, along with a mixture of amino acids and vitamins. The glucose concentration is important: high-glucose DMEM (25 mM) supports the rapid growth of many cell lines, while low-glucose DMEM (5.5 mM) is used for cells that are sensitive to glucose-induced metabolic stress.
L-glutamine is a critical component because it serves as an energy source and a precursor for protein and nucleotide synthesis. However, it is unstable in solution and degrades to ammonia and pyrrolidone carboxylic acid over time. Many laboratories now use stable alternatives such as GlutaMAX, a dipeptide of L-alanine and L-glutamine that is more resistant to degradation. The medium also contains a pH indicator, typically phenol red, which turns yellow under acidic conditions and purple under alkaline conditions, providing a visual check on the health of the culture.
Serum and Serum-Free Media
Serum, usually fetal bovine serum (FBS) at a concentration of 5–10%, is a complex mixture of proteins, growth factors, hormones, lipids, and trace elements that supports cell attachment, proliferation, and survival. FBS contains platelet-derived growth factor (PDGF), fibroblast growth factor (FGF), insulin, transferrin, and many other bioactive molecules. It also provides attachment factors such as fibronectin and vitronectin, which promote cell adhesion to the culture surface.
However, serum has significant drawbacks. It is undefined, meaning its exact composition varies from batch to batch, leading to variability in experimental results. It can also introduce contaminants such as mycoplasma, viruses, or prions, and it raises ethical and cost concerns due to its bovine origin. For these reasons, serum-free media have been developed. These formulations replace serum with defined components: recombinant growth factors, insulin, transferrin, selenium, and specific lipids. Serum-free media are essential for the production of therapeutic proteins, where the presence of animal-derived components is undesirable, and for studies where the effects of specific growth factors need to be isolated. Many cell lines, particularly CHO cells used in biopharmaceutical production, have been adapted to grow in serum-free suspension culture.
Aseptic Technique and Contamination Control
The single most important skill in cell culture is the maintenance of aseptic technique—a set of practices designed to prevent the introduction of microorganisms into the culture. Contamination is the most common cause of cell culture failure, and it can arise from the operator, the environment, or contaminated reagents.
Sterile Work Practices
The fundamental principle of aseptic technique is to minimize the exposure of cells to non-sterile surfaces and air. All work is performed in a biosafety cabinet that has been running for at least 15 minutes to purge the air. The operator wears a laboratory coat, gloves, and often a face mask to prevent the shedding of skin cells and respiratory droplets into the culture. Gloves are sprayed with 70% ethanol before entering the hood, and all items placed in the hood are likewise sprayed.
Pipetting is performed with sterile serological pipettes or micropipette tips, and the pipette is never allowed to touch non-sterile surfaces. Bottles and flasks are opened only inside the hood, and their caps are held in the hand rather than placed on the hood surface. The necks of bottles are flamed briefly with a Bunsen burner or alcohol lamp to create an updraft that prevents airborne contaminants from falling into the opening, although this practice is less common in modern laminar flow hoods where the airflow provides sufficient protection. Work should be organized so that all manipulations are done quickly and efficiently, and the hood should be cleaned with 70% ethanol before and after each session.
Detecting Contamination
Contamination can be bacterial, fungal, or mycoplasmal. Bacterial contamination is usually visible within 24–48 hours: the medium becomes turbid, the pH drops (indicated by a yellow color change of phenol red), and the cells may detach or die. Fungal contamination, often from airborne spores of Aspergillus or Penicillium, appears as fuzzy or filamentous growth on the surface of the medium, sometimes with a musty odor. Mycoplasma contamination is far more insidious: these bacteria lack a cell wall, are only 0.2–0.3 micrometers in diameter, and do not cause visible turbidity or pH change. They can alter cell growth, metabolism, and gene expression without any obvious signs. Mycoplasma contamination is detected by PCR, by staining with a DNA-binding dye such as Hoechst 33258, or by culture on selective agar. Routine testing every 3–6 months is recommended.
If contamination is detected, the affected culture should be autoclaved immediately to prevent spread to other cultures. Antibiotics such as penicillin-streptomycin can be added to the medium to prevent bacterial contamination, but they are not a substitute for good aseptic technique, and their routine use is discouraged because they can mask low-level contamination and select for resistant strains.
Passaging and Maintaining Cells
Cells in culture do not grow indefinitely without intervention. As they proliferate, they consume nutrients, produce waste products, and eventually fill the available surface area or volume. To keep the culture healthy and in the logarithmic growth phase, cells must be passaged (also called subcultured) at regular intervals. This involves transferring a small number of cells to a new vessel with fresh medium.
Cell Counting
Before passaging, it is essential to know the number of viable cells. The standard method is to use a hemocytometer with the trypan blue exclusion assay. Trypan blue is a dye that is excluded from live cells with intact membranes but enters dead cells, staining them blue. A 10 µL sample of the cell suspension is mixed with an equal volume of 0.4% trypan blue, and the mixture is loaded onto the hemocytometer. The cells in the four corner squares of the grid are counted under a microscope, and the cell concentration is calculated using the formula:
Cells per mL = (average count per square) × dilution factor × 10⁴
The dilution factor accounts for the mixing with trypan blue (usually 2), and 10⁴ is the volume correction factor for the hemocytometer chamber. Viability is expressed as the percentage of live cells relative to the total count. For a more detailed explanation of this procedure, see Calculate Cell Viability. Automated cell counters, which use image analysis or electrical impedance, are faster and more reproducible but require the same trypan blue principle.
Cryopreservation and Thawing
Long-term storage of cells is achieved by cryopreservation in liquid nitrogen at −196°C. Cells are suspended in a freezing medium containing 10% dimethyl sulfoxide (DMSO) as a cryoprotectant, which prevents the formation of ice crystals that would damage the cell membrane. The cells are cooled slowly—typically at −1°C per minute—using a controlled-rate freezer or a passive freezing container placed at −80°C overnight, then transferred to liquid nitrogen for storage.
Thawing must be done rapidly to minimize ice crystal damage. The frozen vial is placed in a 37°C water bath with gentle agitation until only a small ice crystal remains, then the contents are transferred to a tube containing pre-warmed medium. The DMSO is toxic at room temperature, so the cells must be centrifuged, the supernatant removed, and the cells resuspended in fresh medium before being placed in a culture vessel. The day after thawing, the medium should be changed to remove residual DMSO and dead cells. For a step-by-step protocol, see Cell Passaging.
The passaging procedure for adherent cells is as follows:
- Aspirate the spent medium from the culture vessel.
- Rinse the cell monolayer with phosphate-buffered saline (PBS) without calcium and magnesium, to remove residual serum that would inhibit trypsin.
- Add a small volume of 0.25% trypsin-EDTA solution (enough to cover the monolayer, typically 1–2 mL for a T75 flask).
- Incubate at 37°C for 2–5 minutes, tapping the flask gently to dislodge the cells. Check under the microscope until the cells have rounded up and detached.
- Add fresh medium containing serum to inactivate the trypsin. The serum contains α1-antitrypsin, which inhibits the protease.
- Gently pipette the cell suspension to break up clumps, then count the cells.
- Seed the desired number of cells into a new vessel with fresh medium, typically at a split ratio of 1:4 to 1:10, depending on the growth rate of the cell line.
Applications of Cell Culture
Cell culture has a vast range of applications across biomedical research, pharmaceutical development, and clinical medicine. The ability to grow cells in a controlled environment has enabled discoveries that would be impossible in whole organisms.
Drug Testing
Cell culture is the primary platform for drug discovery and development. Before a compound is tested in animals, it is screened against panels of cell lines to assess its cytotoxicity, efficacy, and mechanism of action. High-throughput screening (HTS) uses 96-, 384-, or 1536-well plates to test thousands of compounds simultaneously against a target cell line. The readout is often a measure of cell viability, such as the MTT assay, which measures the reduction of a yellow tetrazolium salt to purple formazan crystals by mitochondrial dehydrogenases in living cells. The amount of formazan is proportional to the number of viable cells and is quantified by spectrophotometry. This assay and its variants are described in detail in Mtt Assay Cell Viability.
Cell culture is also used to test the safety of pharmaceuticals, cosmetics, and chemicals. The Ames test uses Salmonella bacteria to detect mutagenic compounds, while mammalian cell lines such as the mouse lymphoma L5178Y line are used to detect chromosomal damage. The 3T3 neutral red uptake assay is used to predict acute oral toxicity, reducing the need for animal testing.
Stem Cell Research
Stem cell research relies heavily on cell culture techniques. Embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs) are cultured under defined conditions that maintain their pluripotency—the ability to differentiate into any cell type of the body. This requires a feeder layer of mouse embryonic fibroblasts or, more commonly today, a defined extracellular matrix such as Matrigel, along with a medium containing basic fibroblast growth factor (bFGF) and transforming growth factor beta (TGF-β) pathway modulators.
The ability to differentiate stem cells into specific lineages—neurons, cardiomyocytes, pancreatic beta cells, and others—has opened new avenues for disease modeling and regenerative medicine. Patient-derived iPSCs can be generated from skin fibroblasts or blood cells, differentiated into the affected cell type, and used to study disease mechanisms in a dish. This approach is particularly valuable for neurological and cardiac diseases where obtaining primary tissue is difficult. The quality control of these cultures, including the verification of pluripotency markers and karyotypic stability, is critical and is discussed in Culture for Stem Cell Quality Control.
Beyond these, cell culture is essential for vaccine production (e.g., the cultivation of viruses in Vero or MDCK cells), the production of recombinant therapeutic proteins (e.g., monoclonal antibodies in CHO cells), and the generation of tissues for transplantation.
Common Pitfalls and How to Avoid Them
Beginners in cell culture often encounter a predictable set of problems. Understanding these failure modes and their causes can save months of frustration.
Contamination
Contamination is the most frequent and devastating problem. The signs are often subtle at first: a slight change in medium color, a small dark spot under the microscope, or a drop in cell viability. By the time contamination is visible to the naked eye, the culture is usually beyond saving. The best approach is prevention: rigorous aseptic technique, routine testing for mycoplasma, and the quarantine of new cell lines until they are confirmed clean. It is also wise to maintain a separate stock of frozen cells as a backup, so that a contaminated culture can be discarded and replaced without losing the cell line.
Cell Line Cross-Contamination
Cell line cross-contamination occurs when one cell line is accidentally mixed with another, usually through the use of shared reagents or pipettes. The most famous example is the contamination of hundreds of other cell lines by HeLa cells, which grow so aggressively that they overgrow and replace the original cells. This problem is so widespread that it is estimated that 15–20% of cell lines in use are misidentified. The consequences are severe: experimental results are invalid, and published data may be irreproducible.
Prevention requires strict adherence to single-use pipettes, the use of separate media bottles for each cell line, and the routine authentication of cell lines by short tandem repeat (STR) profiling, a DNA fingerprinting technique. STR profiling should be performed when a new cell line is received, before freezing a master stock, and periodically thereafter. The International Cell Line Authentication Committee (ICLAC) maintains a database of known misidentified cell lines and recommends that all cell lines be authenticated at least once a year.
Other common pitfalls include allowing cells to become overconfluent, which can cause contact inhibition and changes in gene expression; using expired or incorrectly prepared media; failing to pre-warm media to 37°C, which can shock the cells; and using the wrong trypsin concentration or incubation time, which can damage cell surface proteins. The key to success is consistency: follow the same protocol every time, document everything, and pay attention to the subtle signs that indicate the health of the culture.
Frequently Asked Questions
What is cell culture?
Cell culture is the process of growing cells outside their natural environment, typically in a laboratory vessel containing a nutrient-rich liquid medium. The cells are maintained under controlled conditions of temperature, pH, and humidity, and they can be derived from either animal or plant tissues.
Why is cell culture important?
Cell culture is important because it provides a controlled, reproducible system for studying cellular processes, testing drugs, producing biological molecules, and developing cell-based therapies. It reduces the need for animal experimentation and enables high-throughput screening of compounds.
What are the basic requirements for cell culture?
The basic requirements are a sterile environment (provided by a biosafety cabinet), a controlled incubator (37°C, 5% CO₂, humidified), a suitable culture medium with nutrients and growth factors, and sterile plasticware. Aseptic technique is essential to prevent contamination.
What is the difference between primary cells and cell lines?
Primary cells are derived directly from tissue and have a finite lifespan, typically undergoing 20–60 population doublings before senescing. Cell lines are immortalized and can divide indefinitely, either spontaneously or through genetic manipulation. Primary cells are more representative of the tissue of origin, while cell lines are more convenient and reproducible.
How do you prevent contamination in cell culture?
Contamination is prevented by strict aseptic technique: working in a biosafety cabinet, spraying all items with 70% ethanol, using sterile pipettes and media, wearing gloves and a lab coat, and testing cultures regularly for mycoplasma. Any contaminated culture should be autoclaved immediately.
What does passaging cells mean?
Passaging, also called subculturing, is the process of transferring a small number of cells from an existing culture to a new vessel with fresh medium. This is done when the cells have reached a certain density (typically 70–90% confluence) to keep them in the logarithmic growth phase and prevent overgrowth.
What is the purpose of serum in cell culture media?
Serum, usually fetal bovine serum, provides a complex mixture of growth factors, hormones, attachment factors, and nutrients that support cell proliferation and survival. It is undefined and variable between batches, which is why serum-free defined media are increasingly used for specific applications.
Key Takeaways
- Cell culture is the ex vivo maintenance of cells under controlled conditions, enabling the study of cellular processes in a reproducible and scalable system.
- Primary cells have a finite lifespan, while immortalized cell lines can divide indefinitely; both have distinct advantages and limitations.
- Adherent cells require a surface to attach to, while suspension cells grow freely in the medium; the choice affects passaging and scale-up strategies.
- Aseptic technique is the most critical skill in cell culture; contamination by bacteria, fungi, or mycoplasma is the leading cause of experimental failure.
- Passaging involves detaching adherent cells with trypsin, counting viable cells, and reseeding at a defined density; cryopreservation in liquid nitrogen allows long-term storage.
- Cell culture underpins drug discovery, vaccine production, stem cell research, and the manufacture of therapeutic proteins.
- Cell line misidentification and cross-contamination are serious problems that require routine authentication by STR profiling.