Colony formation assays (CFAs), also known as clonogenic assays, evaluate the ability of a single cell to proliferate into a colony of 50 or more cells, reflecting its long-term growth potential.

Colony formation assays are used to indirectly assess cell viability after treatments such as those involving radiation or cytotoxic drugs, revealing therapeutic impact and growth potential.

In the 1950s, Theodore Puck and Philip Marcus developed the colony formation assay to study the effects of radiation on mammalian cells¹. Their work introduced the concept of reproductive cell death, laying the groundwork for cancer therapy research. Today, scientists rely on colony formation assays in numerous aspects of life sciences research, from oncology to toxicology and genetics.

Principles of the colony formation assay

Colony formation assays test the clonogenic potential of cells – their ability to survive and proliferate into a large colony of ≥50 cells from a single progenitor cell.

A colony is a cluster of at least 50 cells, reflecting about 5-6 divisions from a single progenitor. Puck and Marcus established this threshold, which remains a standard due to several key reasons. First, with typical mammalian cell cycles of 18-24 hours, a colony of 50+ cells confirms sustained proliferation. Second, colonies of this size are easily visible after staining, ensuring reliable counting. Finally, biologically, the threshold indicates that cells have overcome arrest or crisis points.

Colony formation assays reflect a cell’s long-term proliferative capacity, distinguishing it from short-term viability assays that measure only immediate metabolic activity or membrane integrity. This matters because surviving cells do not always retain the ability to proliferate. Studies show that short-term assays often overestimate cell survival, underscoring the need for long-term reproductive assessment through colony formation assays².

Colony formation assays also identify reproductive cell death, when cells look viable but cannot divide, an essential concept in cancer therapy, where stopping tumor repopulation is critical. By counting colonies, the impact of radiation or drugs on a cell’s long-term survival can be assessed, making colony formation assays a gold standard for evaluating treatment impact.

Reagents and materials

Cell culture media: Cell culture media is essential for maintaining cell viability and promoting continuous cell proliferation throughout clonogenic assays. Researchers rely on a balanced mix of amino acids, vitamins, minerals, carbohydrates, growth factors, and hormones. Any imbalance can significantly reduce colony formation³.

The specific type of culture medium used (eg, modified Eagle’s medium, minimum essential medium, and serum-free keratinocyte medium) depends on the particular cell line being cultured, as each cell type has distinct nutritional requirements.

To further promote cell growth and attachment, the media is typically supplemented with 10% fetal bovine serum (FBS). Reducing serum to 5% can lower colony formation in epithelial cells⁴. The addition of antibiotics such as penicillin and streptomycin prevents contamination, though long-term use may slightly reduce colony yield in sensitive lines⁵. Fresh L-glutamine at 2–4 mM boosts energy supply and improves colony formation.

In assays involving non-adherent or hematopoietic cells, semi-solid methylcellulose media can be used to prevent cell movement and ensure each colony arises from a single progenitor. The addition of cytokines such as erythropoietin, granulocyte-macrophage colony-stimulating factor (GM-CSF), and interleukin-3 (IL-3) supports specific lineages of erythroid cells, granulocyte-macrophage progenitors, and multipotent progenitors, respectively⁶.

The choice of medium directly affects assay outcomes⁷. Plating efficiency (PE) can vary across different media for the same cell line. Media composition also influences colony size, morphology, and post-treatment survival. These factors make careful media selection essential for accurate and reproducible colony formation assay results.

Trypsin and EDTA: Trypsin and EDTA effectively create viable single-cell suspensions, especially from adherent cells. Trypsin is a serine protease that cleaves adhesion proteins at lysine and arginine amino acid residue sites, disrupting interactions between cells and the surrounding matrix⁸. EDTA enhances this process by chelating calcium ions essential for adhesion. Together, they improve dissociation accuracy over mechanical methods, ensuring clump-free suspensions for reliable colony counting⁷.

The effectiveness of trypsin depends on concentration and formulation. Commonly, labs use 0.05% for sensitive lines and 0.25% for robust or dense cultures. Most trypsin solutions contain 0.02-0.53 mM EDTA and are prepared in calcium- and magnesium-free phosphate-buffered saline (PBS) (pH 7.2-7.4); trypsin loses activity below pH 6.8.

Incubation time and temperature critically influence cell viability. Typically, cells are incubated with trypsin for 2–10 minutes at 37°C. Fibroblasts require 3-5 minutes, epithelial cells 5-8 minutes, and strongly adherent lines up to 10 minutes. Lower temperatures cut enzyme activity in half, while prolonged exposure at 37°C can degrade membrane proteins, reducing cell viability⁹.

To prevent damage, promptly neutralize trypsin using a serum-containing medium. FBS provides inhibitors such as α1-antitrypsin and α2-macroglobulin. A 2:1-5:1 ratio of serum to trypsin ensures full inactivation. Incomplete neutralization risks cell loss and lower colony yield¹⁰.

Optimizing trypsinization improves colony formation. Exposure time to trypsin can be tailored by cell type for higher viability, verified via trypan blue or flow cytometry¹¹. Avoid overexposing cells to trypsin, as its proteolytic activity can degrade cell surface proteins. This degradation disrupts cell function and impairs downstream processes¹².

Fixatives and staining solutions: Effective fixation and staining are essential for preserving colony structure and enabling accurate counting in clonogenic assays. Fixatives improve colony retention by cross-linking proteins and nucleic acids, halting metabolic activity, and enhancing adherence to culture plates. Stains bind to cellular components, increasing contrast, facilitating visualization, and reliably quantifying colony formation.

The following table presents the preparation of fixative and staining solutions in colony formation assays.

Fixative and staining solution choices should align with specific experimental goals. PFA better maintains cellular structure, while methanol offers speed and is preferred for applications such as nuclear staining.

The choice between crystal violet and Giemsa depends on the experimental objectives:

• Colony counting: Crystal violet offers clearer boundaries and performs better with automated systems, while Giemsa delivers 88-92% accuracy along with morphological detail⁷.
• Procedure complexity: Crystal violet requires a simple, single-step process, whereas Giemsa involves a longer, multi-step protocol.
• Information yield: Crystal violet is sufficient for basic quantification and size measurements, while Giemsa provides insights into cellular differentiation and morphology.

Other essential materials and equipment

Successful colony formation assays require proper equipment, sterile techniques, and precise material handling.

Culture vessels: The choice of culture vessels directly affects colony formation and quantification. Multi-well plates are normally used for adherent cells; 6-well plates are standard, while 12- and 24-well plates serve smaller samples. Smaller wells can reduce colony size due to limited nutrients and increased contact inhibition⁷.

Surface treatment and material composition also matter. Tissue culture-treated surfaces, modified to promote cell attachment, are essential for adherent cells. Non-treated surfaces support suspension cultures or spheroid formation. Polystyrene remains the most common vessel material due to its optical clarity, while specialty polymers are used for sensitive cells needing specific surface chemistry.

Liquid handling tools: Accurate liquid handling ensures assay reproducibility. Serological pipettes are used for bulk transfers, and motorized controllers are used to ensure consistency. Micropipettes, ranging from 0.5-1000 µL, allow precise cell seeding. Single-channel pipettes suit small-scale work, while multi-channel options enable high-throughput applications. Calibration is key to maintaining seeding precision. Sterile filtration devices, such as syringes and bottle-top filters, reduce contamination and remove particulates from reagents and media.

Centrifugation and tubes: Centrifugation and tube selection influence cell viability. Centrifuges with temperature control and speeds from 200-1500 x g are standard, with swinging bucket rotors preferred for pelleting. Sterile, cytotoxic-free conical or microcentrifuge tubes are used for cell collection. Single-use, gamma-irradiated tubes minimize contamination risk.

Sterility and contamination control tools: Maintaining a sterile environment is critical. A Class II biosafety cabinet with HEPA filtration and UV sterilization ensures asepsis. Sterile techniques can be practiced using 70% ethanol, sterile lab coats, gloves, and minimal movement to reduce airflow disruption. These precautions are vital for assays that run over 1-3 weeks.

Environmental control through a CO₂ incubator supports optimal cell growth. Standard conditions include 37°C ± 0.2°C and 5% CO₂ to maintain a medium pH between 7.2-7.4. Even minor temperature fluctuations can impact colony formation. Humidity levels should remain between 95-98%, maintained by a water pan containing copper sulfate to deter microbial growth.

Microscopy and imaging tools: Microscopy is essential for colony monitoring and analysis. Inverted microscopes with phase contrast allow for the observation of early colony growth without disturbing cultures. Stereomicroscopes enhance colony visibility through oblique illumination and support manual counting. Digital systems with high-resolution cameras and software enable documentation and automated analysis.

Cell counting tools: Accurate cell counting before plating is crucial. Manual counting using a hemocytometer and trypan blue remains common. Automated systems reduce operator variability and speed up viability assessments. Flow cytometry offers advanced cell population analysis, including viability, cell cycle status, and surface marker expression, particularly valuable for heterogeneous samples.

Colony quantification tools: Colony quantification marks the assay’s endpoint. Manual tools include grid-marked overlays and counting pens, with colonies containing over 50 cells typically considered. Automated image analysis software can assess colony number, size, and morphology with high-throughput compatibility, processing up to 96-well plates with accuracy matching manual methods.

Quality and calibration tools: High-quality equipment underpins assay reliability. Calibrated tools and documented maintenance schedules reduce variability. Standard operating procedures and performance validations with controls and reference cell lines further enhance consistency and accuracy across experiments.

Colony formation assay procedure

Cell preparation: A single-cell suspension should be prepared from the cell line of interest. For adherent cells, trypsinization should be used for harvesting. The culture medium should be removed, cells washed with PBS, and a trypsin-EDTA solution added. The cells should then be incubated at 37°C until detachment occurs. Trypsin activity should be neutralized with culture medium, and the suspension should be pipetted to ensure homogeneity. The viable cell count should be determined using a hemocytometer or an automated cell counter. Cell viability should be assessed through trypan blue exclusion.

Cell seeding and plating: The optimal number of cells to seed should be determined. For untreated control cells, a target of 20-150 colonies per well in a 6-well plate should be aimed for. Optional: Seeding density for treated cells can be adjusted based on the anticipated survival fraction. Serial dilutions should be performed, and cells should be seeded into culture plates with the appropriate medium. The plates should be gently swirled to ensure even distribution. Prior to plating, it should be ensured that all cells are present as single cells through thorough mixing and adequate dilution. After plating, the plates should be examined under a brightfield microscope to confirm that only single cells are present.

Incubation: Incubate culture plates at 37°C in a humidified atmosphere with 5% CO₂. Incubation duration varies from 1-3 weeks. Ensure at least six cell divisions for the untreated control conditions. Periodically check plates under a microscope, change culture medium every 3–4 days, and maintain stable incubation conditions.

Fixation and staining: The medium should be gently aspirated at an angle, and an equal volume of pre-warmed fresh medium should be added. The plates must be returned to the incubator within 10 minutes to maintain stable conditions.

Colonies should be fixed once each of them contains 50 or more cells. Fixation should be performed in a well-ventilated area. The culture medium should be removed, cells washed with PBS or saline, and the fixative solution added to cover the colonies. Common fixatives include PFA, methanol, glutaraldehyde, and formalin.

For staining, the fixative solution should be removed, and the cells washed with PBS or distilled water. The staining solution should then be added to cover the colonies. Common staining solutions include crystal violet and Giemsa. After staining, the plates should be gently washed, air-dried at room temperature, and stored prior to colony counting.

Refer to the following table for more details on common fixing methods.

Analyzing and quantifying colony formation

Traditional manual counting: Microscopes such as bright-field, inverted, and stereomicroscopes are often used for manual counting. Stereomicroscopes offer better visibility of colony edges, often resulting in more accurate counts than standard inverted microscopes.

Manual colony counting, using a permanent marker to mark the single cells and tally them simultaneously, remains common. To reduce counting errors on large plates, they are often divided into sections like quadrants¹⁶. Accurate counting requires trained personnel, standardized protocols, and periodic validation. Some labs use two observers to compare and average counts for greater accuracy.

Limitations of manual counting:

• Time-consuming: Counting over 100 colonies can take 5-10 minutes per plate, leading to hours of work in experiments with multiple conditions (eg, several different drug concentration).
• Prone to error: Visual fatigue and subjective interpretation reduce accuracy, especially when colony density exceeds 200 per 60-mm dish⁷.

Automated image analysis: Automated colony counting systems may transform the quantification process in colony formation assays by addressing key limitations of manual methods. The automated process involves three main steps:

1.     Image acquisition: Systems capture digital images using flatbed scanners (≥600 dpi), digital cameras on stands, or automated microscopes with motorized stages.

2.     Image processing: Software analyzes images through background subtraction, threshold adjustment, and colony identification based on set parameters.

3.     Data analysis: The software quantifies colony numbers, size distribution, and morphological features.

Some systems detect about 97% of colonies identified by expert manual counting, with advanced algorithms reaching over 99% concordance¹⁷.

Various software packages are available, often integrated with compatible hardware. They range from fully automated high-priced systems¹⁸ to free tools such as ImageJ from the National Institutes of Health, which are widely used by researchers given their widespread adoption, availability of free plugins, and the support around the software¹⁹.

Key metrics and calculations

Plating efficiency: PE measures a cell’s ability to form colonies under optimal, untreated conditions. It is calculated as:

PE (%) = (Number of colonies formed/Number of cells seeded) × 100

This value reflects the proportion of seeded cells that successfully grow into colonies in control conditions.

PE acts as a baseline for assessing treatment effects. Consistent PE values suggest reliable techniques and healthy cultures. Different cell lines have characteristic PE values that reflect their clonogenic potential. Several factors influence PE:

• Cell handling: Poor pipetting or mechanical stress can reduce PE⁷.
• Growth medium: Lowering serum from 10% to 2% negatively affects PE.
• Cell density: Overcrowding lowers PE due to contact inhibition. Sparse seeding may reduce PE due to reduced paracrine signaling²⁰.

A high PE value indicates optimal conditions and healthy cells, whereas a low PE may reflect stress, poor technique, or suboptimal conditions.

Survival fraction (SF): SF measures the proportion of cells that maintain reproductive ability after treatment, normalized to the untreated control.

Use the standard formula to calculate SF:
SF = (Colonies after treatment/Cells seeded) ÷ (Plating efficiency/100)

Or

SF = (Colonies after treatment/Cells seeded) × (Control cells seeded/Control colonies formed)

Example:

Control: 100 cells seeded; 70 colonies formed (PE = 70%)

Treatment: 200 cells seeded; 70 colonies formed (PE = 35%)

SF = (70/200) ÷ (70/100) = 0.5 = 50%

This result means the treatment reduced clonogenic capacity to 50% of the control.

SF values reveal how treatments affect cell survival and are essential for generating dose-response curves, calculating IC₅₀ values, and assessing sensitivity. In radiobiology, researchers often fit SF data to the linear-quadratic model SF = e^{(−αD − βD²)}. It helps estimate radiosensitivity parameters α and β²¹. Relative clonogenic survival assesses clonogenic assays by comparing the survival of treated cells to that of control cells.

The following table lists key formulas for colony formation assay analysis:

Applications of the colony formation assay

Assessing treatment efficacy in cancer research: The clonogenic assay is widely used for evaluating cancer treatment effectiveness. In radiation therapy, colony formation assays help generate survival curves that show how the surviving fraction (SF) changes with increasing radiation dose (D). These curves follow the linear-quadratic model:

SF = e^{(−αD − βD²)}, where α represents lethal damage from a single radiation event and β represents damage from two separate events.

The α/β ratio, which reflects tissue radiosensitivity.

In chemotherapy, colony formation assays can be used to assess drug potency using IC₅₀ values, the concentration required to reduce colony formation by 50%. Colony formation assays can also be used for evaluating treatment combinations²².

Toxicology: The colony formation assays demonstrate long-term cellular toxicity caused by various chemicals.

Cadmium, an environmental toxin, increases oxidative stress and triggers apoptosis through the mitochondrial pathway in rainbow trout hepatocytes²³. Exposure to cadmium at concentrations of 2, 5, and 10 μM activates caspase-3, caspase-8, and caspase-9, ultimately leading to cell death²³.

Colony formation assays can also be used to evaluate the toxicity of nanomaterials. In MDA-MB-231 breast cancer cells, biosynthesized silver nanoparticles (AgNPs) exhibited an IC₅₀ of 8.7 μg/mL²⁴. In contrast, fungal-derived AgNPs had a higher cytotoxic concentration (CTC₅₀) of 260 μg/mL in human gingival fibroblasts, indicating that normal cells tolerate these nanoparticles better than cancer cells²⁵.

Genetic studies to evaluate gene functions related to growth and survival: Researchers have used colony formation assays to clarify gene function, especially in the context of CRISPR/Cas9 studies.

In B16F10 melanoma cells, knocking down PTGS2 with CRISPR/Cas9 slowed cell proliferation²⁶. Colony formation assays were used to confirm this effect, identifying PTGS2 as a potential therapeutic target for melanoma.

To validate tumor suppressors, scientists deleted the p107 gene in a murine small cell lung cancer (SCLC) model using CRISPR/Cas9. Colony formation assays showed that the loss of p107 accelerated tumor growth, produced fewer but larger tumors, and led to earlier metastasis. This highlighted p107’s role as a tumor suppressor²⁷.

Stem cell biology: The colony formation assay offers key insights into stem cell behavior, particularly self-renewal and potency.

One study classified epidermal stem cells based on their colony-forming ability, identifying three keratinocyte types²⁸. Holoclones formed large, fast-growing colonies with 100% secondary colony formation. Meroclones and paraclones produced smaller and abortive secondary colonies, respectively. Another study used colony formation assays to quantify the stem cell potency of human embryonic stem cells and pluripotent stem cells²⁹.

Pharmacological screening: Colony formation assays support efficient drug screening by enabling high-throughput and precise synergy detection.

Using a 96-well format, researchers screened over 1,000 compounds per week with automated image-based colony analysis³⁰. Colony formation assays were more valuable compared to short-term assays in detecting drug synergy³¹. The 50% colony inhibition combination index provided robust interaction metrics, while colony growth patterns helped refine treatment schedules.

Using optimized colony forming assay kits can significantly improve accuracy and reproducibility.

FAQs

What types of cells can be studied using the clonogenic assay?

Any adherent cell that can proliferate under control conditions can be studied using the colony formation assay, as these cells can attach to the culture surface and divide to form colonies over time. Commonly studied cells include cancer cells, which help assess tumorigenic potential and treatment response; stem cells, which allow evaluation of self-renewal and differentiation capacity; and primary epithelial cells, which provide insights into tissue regeneration and cellular behavior under different conditions. The assay’s versatility makes it a valuable tool across oncology, regenerative medicine, and toxicology research.

How can you ensure reproducibility and accuracy in results?

You can ensure reproducibility and accuracy in clonogenic assays by standardizing cell seeding density, which helps maintain consistent colony growth across experiments. They maintain uniform culture conditions, such as temperature, CO₂ levels, and media composition, to prevent variability in cell behavior. They also prepare single-cell suspensions carefully by fully dissociating cell clumps, ensuring each colony originates from a single cell. To minimize subjectivity and human error, they use automated or blinded colony counting methods, which provide objective and consistent measurements across samples.

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