Cell Culture Basics: A Beginner's Guide

Cell culture is the technique of growing and maintaining cells outside their original organism, in a controlled artificial environment. For a new lab member, it is often the first hands-on skill to learn and the easiest one to get wrong in ways that are not obvious until weeks later, when a contaminated flask or a mishandled passage quietly ruins months of work. This guide covers the fundamentals every beginner needs before starting bench work: what cell culture actually involves, the equipment and media you will use daily, the aseptic habits that separate a clean culture from a contaminated one, how to subculture and count cells, and the biosafety rules that apply to the cells themselves. For the sterile-technique habits referenced throughout, see CASRAI’s companion guide to aseptic technique; for long-term storage of the lines you establish, see cryopreservation of cells.

What Is Cell Culture?

“Cell culture” broadly refers to any process of maintaining living cells in vitro (outside the body) under conditions that support their survival and, usually, their growth. Most new lab members work with one of three categories of material:

Primary cells — cells isolated directly from tissue (a biopsy, a blood draw, a dissociated organ) and cultured without genetic modification. Primary cultures are the closest to native biology but have a finite lifespan: they can typically only be subcultured a limited number of times before they senesce and stop dividing.
Cell strains — primary cells that have been subcultured but retain a finite lifespan and largely normal karyotype, as opposed to cell lines proper.
Immortalized (continuous) cell lines — cells that have acquired the ability to divide indefinitely, either spontaneously (as with many cancer-derived lines) or through deliberate immortalization (e.g., viral transformation, telomerase expression). Lines such as HeLa, HEK293, and CHO are examples most new lab members will recognize by name; they are easier to maintain and standardize than primary cultures but diverge further from native tissue biology the longer they are passaged.

Cultures are also described by growth pattern. Adherent cultures grow attached to the surface of a flask or plate and must be enzymatically or mechanically detached to be counted, passaged, or harvested. Suspension cultures (many blood-derived and hybridoma lines) grow freely in the media and are handled without a detachment step. Which type you are working with determines almost every procedure below.

Core Equipment You Will Use Every Day

A standard mammalian cell culture setup centers on a small number of pieces of equipment, each doing a specific job:

Biosafety cabinet (BSC) — nearly all routine cell culture work is performed in a Class II biosafety cabinet, which protects the culture from environmental contaminants and, depending on the cell source, protects the operator from the material being handled. See CASRAI’s guide to biosafety cabinet classes for how Class I, II, and III cabinets differ and when each is required.
CO2 incubator — maintains the temperature, humidity, and gas atmosphere most mammalian cultures need to grow. A standard set point is 37°C, roughly 95% relative humidity, and 5% CO2; the CO2 level is calibrated to the bicarbonate buffering system used in most standard culture media (see below), not an arbitrary figure, so it should not be changed without also reconsidering the media’s buffer.
Inverted microscope — used to check cultures daily for confluency, morphology, and early signs of contamination without removing the vessel from the incubator for longer than necessary.
Centrifuge — used to pellet cells after detachment or during media changes for suspension cultures. See CASRAI’s guide to centrifuge rotor balancing for the safety practices that apply here.
Water bath — used to pre-warm media and reagents (typically to 37°C) before they contact cells, since cold media can shock and detach adherent cultures.
Hemocytometer or automated cell counter — used to determine cell concentration and viability before plating or passaging (see the counting section below).

Aseptic Technique: The Non-Negotiable Skill

Every step above depends on rigorous aseptic technique — the set of practices that keep a culture free of bacteria, fungi, yeast, and other cells. CASRAI’s dedicated aseptic technique guide covers this in depth; for cell culture specifically, the habits that matter most on day one are:

Work inside the biosafety cabinet’s sterile field, keeping hands and objects away from the front air intake grille and avoiding movements that disrupt laminar airflow.
Spray gloves and the outer surface of any item (bottle, pipette, tube) with 70% ethanol before it enters the cabinet.
Never open more than one cell line’s vessel in the cabinet at the same time, and decontaminate the work surface between different cell lines, not just between sessions.
Use a dedicated, filtered pipette tip or serological pipette for each reagent and each cell line — never reuse a tip or pipette across bottles or lines.
Check media and reagent bottles for cloudiness, color shift, or particulate before use; a compromised reagent bottle can contaminate every culture it touches.

Culture Media: What Is Actually in the Bottle

Growth media supplies the nutrients, salts, and signaling factors a culture needs. Most beginner protocols use a commercially formulated basal medium — common examples include DMEM (Dulbecco’s Modified Eagle Medium) and RPMI-1640 — supplemented with:

Serum, most often fetal bovine serum (FBS), typically added at around 5-10% of the final volume. Serum supplies growth factors, hormones, and attachment proteins that most basal media lack on their own.
Antibiotics (e.g., penicillin-streptomycin), commonly added to routine culture as a contamination safety margin — though many labs and cell-line providers now discourage routine antibiotic use in cultures intended for sensitive downstream assays, since it can mask low-level contamination rather than prevent it.
A buffering system — most standard media rely on a sodium bicarbonate/CO2 buffer, which only functions correctly inside a CO2-controlled incubator at the correct CO2 percentage; media intended for bench work outside the incubator (e.g., during a long procedure at the biosafety cabinet) is often additionally buffered with HEPES to hold pH stable in ambient air.

Complete media should be stored according to the manufacturer’s instructions (typically refrigerated, protected from light) and pre-warmed before use — but not held at 37°C for extended periods when not in use, since some components (notably glutamine) degrade with prolonged warm storage.

Daily and Weekly Maintenance: Feeding and Observing Cultures

Between passages, a culture needs regular attention. Daily, most labs visually check each active culture under the inverted microscope for confluency (the percentage of the vessel surface covered by adherent cells, or density for suspension cultures), morphology (cells should look consistent with how that line normally appears — rounding, granularity, or debris can signal stress or contamination), and media color (phenol red, the pH indicator in most standard media, shifts from red/pink toward yellow as the culture acidifies with growth, and toward purple/pink if it becomes too alkaline). Media is exchanged on a schedule appropriate to the line’s growth rate — faster-growing or more metabolically active lines need more frequent feeding — replacing spent, depleted media with fresh, pre-warmed media without disturbing the adherent monolayer.

Subculturing (Passaging): Keeping Cultures Healthy

Subculturing — commonly called “passaging” or “splitting” — means transferring a portion of an established culture into fresh media and, usually, a fresh or larger vessel, before the existing culture becomes overgrown. For adherent cultures, the standard sequence is:

Remove and discard spent media.
Rinse the monolayer with a calcium- and magnesium-free buffer (commonly PBS) to remove residual serum, which inhibits the detachment enzyme.
Add a dissociation reagent — most often trypsin-EDTA — and incubate briefly (typically a few minutes at 37°C) until cells round up and detach, checked periodically under the microscope rather than left on a fixed timer, since over-exposure to trypsin damages cell-surface proteins and viability.
Neutralize the trypsin with serum-containing media (serum proteins inhibit trypsin activity) and gently resuspend the detached cells.
Count the cell suspension (see below), then seed a known number or fraction of cells into new vessel(s) with fresh media at the split ratio the protocol for that line calls for.

Suspension cultures skip the enzymatic detachment step entirely: cells are simply diluted with fresh media, or a fraction of the culture is removed and replaced with fresh media, once density reaches the target range for that line.

Two records matter every time a culture is passaged: the passage number (how many times the line has been subcultured since it was thawed or established) and the split ratio used. Passage number matters because most cell lines drift — genetically, phenotypically, or in growth behavior — the longer they are kept in continuous culture, and many labs set a maximum passage number past which a line is discarded and a fresh vial thawed from frozen stock rather than used for experiments.

Counting Cells and Checking Viability

Before plating an experiment or passaging at a defined ratio, cells are counted and their viability assessed. The traditional method uses a hemocytometer — a specialized microscope slide with an etched counting grid of known volume — combined with the trypan blue exclusion assay: trypan blue dye is excluded by cells with an intact plasma membrane (viable cells appear clear/unstained under the microscope) but freely enters cells with a compromised membrane (dead or dying cells stain blue). Counting a defined number of grid squares and applying the hemocytometer’s known chamber volume yields a cell concentration; the ratio of unstained to total cells yields a viability percentage. Many labs now use automated cell counters that apply the same trypan blue principle (or an image-based/fluorescence-based equivalent) with less manual counting error, particularly for higher-throughput work.

Recognizing and Preventing Contamination

Contamination is the most common way a beginner loses a culture, and it comes in two very different forms:

Visible contamination — bacteria, yeast, and fungi typically announce themselves quickly: sudden media turbidity or cloudiness, a rapid pH shift (media turning yellow faster than expected), visible particulate, filaments, or a film on the media surface, and a sour or off odor when the vessel is opened. Once confirmed, a visibly contaminated culture is normally discarded (autoclaved or chemically decontaminated per institutional biohazard waste procedures) rather than “rescued,” and every other culture and reagent that may have shared the same cabinet session should be checked.
Mycoplasma contamination — the more insidious problem, because mycoplasma (a genus of cell-wall-lacking bacteria) does not cause visible turbidity, an obvious pH shift, or a distinct odor; a heavily mycoplasma-infected culture can look completely normal under a standard light microscope while still altering cell growth, metabolism, and experimental results. Because it is invisible on inspection, mycoplasma is detected with dedicated testing — PCR-based assays or enzymatic luminescence kits — and many labs and journals now require routine, periodic mycoplasma testing of any line used in published work. Surveys of cell culture facilities over the years have repeatedly found mycoplasma-positive cultures at a meaningful, non-trivial rate, which is why testing is treated as routine rather than optional in a well-run lab.

A related but distinct risk is cross-contamination — one cell line overtaking or contaminating another, usually from shared reagents, non-dedicated pipettes, or simultaneous handling of multiple lines in the same cabinet session. Misidentified and cross-contaminated cell lines have been a long-documented problem in the literature; the International Cell Line Authentication Committee (ICLAC) maintains a public register of known misidentified/contaminated lines, and short tandem repeat (STR) profiling — recommended by resources such as ATCC — is the standard method for confirming a line’s identity. Rigorous aseptic technique (one line open at a time, dedicated consumables per line, cabinet decontamination between lines) is the primary defense against both mycoplasma and cross-contamination, since both usually trace back to a lapse in the habits covered above.

Biosafety Considerations for Cell Culture

Not all cell culture carries the same biosafety classification. Well-characterized, long-established immortalized lines with no known infectious agents are typically handled at Biosafety Level 1 (BSL-1). Primary human cells, primary cells of unknown infectious status, and any culture involving human blood, body fluids, or unfixed human tissue are, by default, handled at Biosafety Level 2 (BSL-2) under CDC/NIH Biosafety in Microbiological and Biomedical Laboratories (BMBL) guidance and, in the United States, under OSHA’s Bloodborne Pathogens Standard (29 CFR 1910.1030), because the infectious status of primary human material generally cannot be assumed negative without testing. In practice this means routine use of a certified biosafety cabinet for any aerosol- or splash-generating step (which most cell culture procedures are), and appropriate PPE — see CASRAI’s guide to PPE selection for glove, eyewear, and lab coat guidance that applies alongside biosafety-cabinet use. New lab members should confirm the assigned biosafety level and any institutional biosafety committee (IBC) protocol requirements for their specific cell line or tissue source before starting work, rather than assuming BSL-1 by default.

Freezing Cells for Long-Term Storage

Once a culture is established, it is standard practice to freeze down a bank of early-passage stock rather than keep a single line in continuous culture indefinitely — continuous passaging accelerates genetic drift and raises contamination risk the longer a line stays active on the bench. CASRAI’s companion guide, Cryopreservation of Cells: Basic Protocol and Best Practices, covers the freezing protocol itself — cryoprotectant selection, controlled cooling rate, and long-term liquid-nitrogen storage — in full; the short version for a new lab member is that a low-passage master stock, frozen soon after a line is received or established, is what lets a lab recover from a lost or contaminated culture without starting over from an external source.

Common Beginner Mistakes to Avoid

Skipping the pre-cabinet wipe-down or gown-up. Aseptic technique is a routine, not an occasional precaution — lapses compound over repeated sessions even when no single lapse causes an obvious problem.
Letting cultures overgrow before passaging. Cultures left past their intended confluency or density can become nutrient- and space-limited, changing growth behavior and, for some lines, reducing post-passage viability.
Over- or under-trypsinizing. Watching cells detach under the microscope, rather than trusting a fixed timer, avoids both incomplete detachment and membrane damage from prolonged enzyme exposure.
Handling multiple cell lines in the same cabinet session without decontaminating between them. This is the single most common route to cross-contamination.
Skipping or delaying mycoplasma testing because the culture “looks fine.” Visual inspection cannot rule out mycoplasma; only a dedicated assay can.
Poor labeling and record-keeping. Every vessel should be labeled with the cell line, passage number, and date; every passage should be logged. This is what makes a passage-number cutoff or a contamination trace-back actually possible later.

Frequently Asked Questions

What is the difference between a primary cell culture and a cell line?

Primary cells are isolated directly from tissue and have a limited number of times they can be subcultured before they senesce. A cell line (or continuous/immortalized line) has acquired the ability to divide indefinitely, either spontaneously or through deliberate immortalization, and can be passaged far more times without losing the ability to grow — though it will still drift genetically and phenotypically over many passages.

What is the difference between adherent and suspension culture?

Adherent cells grow attached to the surface of the culture vessel and must be enzymatically or mechanically detached (typically with trypsin-EDTA) to be counted, passaged, or harvested. Suspension cells grow freely dispersed in the media and are handled by simple dilution or partial media replacement, without a detachment step.

How often should I passage my cells?

There is no single universal schedule — it depends on the specific line’s growth rate and the target confluency or density set by the protocol for that line. As a general principle, adherent cultures are typically split before they become fully confluent (commonly in a moderate-to-high confluency range well short of 100%), since overgrowth changes cell behavior and can reduce viability at the next passage.

How do I know if my culture has mycoplasma contamination?

You generally cannot tell by eye. Mycoplasma does not cause the visible turbidity, pH shift, or odor that bacterial or fungal contamination typically does, and an infected culture can look completely normal under a standard light microscope. Routine, periodic testing with a PCR-based or enzymatic luminescence assay is the only reliable way to detect it, which is why many labs and journals require documented mycoplasma testing before a line’s data can be published.

What biosafety level applies to cell culture work?

It depends on the cell source. Well-characterized immortalized lines with no known infectious agents are commonly handled at BSL-1. Primary human cells, cells of unknown infectious status, and anything derived from human blood, body fluids, or unfixed tissue are, by default, handled at BSL-2 under CDC/NIH BMBL guidance and OSHA’s Bloodborne Pathogens Standard, since infectious status generally cannot be assumed negative without testing. Confirm the assigned level and any IBC protocol requirements for your specific material before starting.

Why do I need to freeze down cell stocks instead of just keeping one culture going?

Continuous passaging accelerates genetic and phenotypic drift and increases cumulative contamination risk the longer a line stays actively growing on the bench. Freezing a low-passage master stock soon after a line is received or established lets a lab discard a drifted, aged, or contaminated working culture and recover a fresh, early-passage replacement instead of losing the line entirely or re-sourcing it externally.