What is Dr. Zubair Khalid's research focus?

Dr. Zubair Khalid specializes in molecular virology, mRNA vaccine development, and computational biology, with a focus on avian pathogens like IBDV and Avian Reovirus.

Where is Dr. Zubair Khalid currently working?

Dr. Zubair Khalid is a Postdoctoral Research Associate at the University of Maryland (UMD), specifically within the Department of Animal and Avian Sciences.

Mycoplasma Contamination PCR: Detection and Prevention

By Dr. Zubair Khalid, DVM, MS, PhD · 2026-08-11

Introduction to Mycoplasma Contamination

Mycoplasma contamination is one of the most persistent and underappreciated problems in cell culture laboratories. Unlike bacterial or fungal contamination, which typically produces visible turbidity or pH changes within hours to days, mycoplasma infections can persist silently for months, altering cellular physiology without any obvious signs. PCR-based detection has become the gold standard for monitoring these infections because it is rapid, sensitive, and capable of detecting multiple species simultaneously.

What Are Mycoplasmas?

Mycoplasmas are the smallest self-replicating prokaryotes, belonging to the class Mollicutes. They lack a rigid cell wall, a feature that distinguishes them from all other bacteria and renders them intrinsically resistant to cell-wall-targeting antibiotics such as penicillin and streptomycin—the very antibiotics most commonly added to cell culture media. Their genome is remarkably small, ranging from approximately 580 to 1,400 kilobases, and they have evolved as obligate or facultative parasites that depend on host cells for many nutrients, including sterols and amino acids.

The absence of a cell wall also explains their pleomorphic morphology; mycoplasmas can appear as coccoid bodies, filaments, or rings, and they pass through 0.22 µm filters commonly used to sterilize media. This filter-passing ability is a major reason they evade routine contamination controls. In cell culture, the most frequently encountered species are Mycoplasma arginini, M. hyorhinis, M. orale, M. fermentans, and Acholeplasma laidlawii, with M. fermentans and M. hyorhinis being particularly common contaminants of laboratory cell lines.

Why Contamination Matters

Mycoplasma contamination is not a benign nuisance. Infected cultures exhibit a wide range of phenotypic alterations that can invalidate experimental results. Mycoplasmas compete with host cells for amino acids, nucleotides, and other precursors, leading to reduced cell proliferation and altered metabolic profiles. They also hydrolyze arginine via the arginine deiminase pathway, depleting this essential amino acid from the medium and causing growth arrest in many cell types. Some species, particularly M. hyorhinis, can induce chromosomal aberrations and modulate gene expression, while others produce nucleases that degrade DNA and RNA in the culture supernatant.

The most insidious aspect of mycoplasma contamination is its invisibility. Infected cultures often show no turbidity, no pH change, and no obvious cytopathic effect. Researchers may continue experiments for weeks or months with contaminated cells, producing data that are irreproducible or biologically meaningless. A 2015 meta-analysis of published studies estimated that between 5% and 30% of all continuous cell lines worldwide are contaminated, with some repositories reporting rates above 30%. This makes routine monitoring essential, and PCR has emerged as the most practical detection method for most laboratories.

Principles of PCR Detection

The Polymerase Chain Reaction amplifies specific DNA sequences through repeated cycles of denaturation, annealing, and extension. For mycoplasma detection, the goal is to amplify conserved regions of the mycoplasma genome while avoiding amplification of host cell DNA or DNA from other bacteria. This is achieved by selecting target genes that are conserved across mycoplasma species but divergent from eukaryotic and other prokaryotic sequences.

Target Genes for Mycoplasma Detection

The most widely used target for mycoplasma PCR is the 16S ribosomal RNA (rRNA) gene. This gene is present in all prokaryotes, contains both highly conserved and variable regions, and exists in multiple copies per genome—typically one to two in mycoplasmas, but the high transcription rate of rRNA genes makes them excellent targets even with low genome copy numbers. Primers are designed against conserved regions flanking variable regions, allowing a single primer pair to amplify DNA from a broad range of mycoplasma species while producing amplicons of slightly different sizes or sequences that can be distinguished by restriction digestion or sequencing.

A second commonly used target is the 16S-23S rRNA intergenic spacer region (ITS). This region evolves faster than the 16S gene itself, providing better discrimination between closely related species. However, its higher variability means that a single primer pair may not amplify all species with equal efficiency, and some laboratories use multiple primer sets to ensure comprehensive coverage.

The tuf gene, encoding elongation factor Tu, is another target used in some commercial kits. It is a single-copy gene in most mycoplasmas, which reduces sensitivity compared to rRNA-based targets but offers excellent specificity. Finally, some assays target the gap gene (glyceraldehyde-3-phosphate dehydrogenase) or species-specific genes for identification of particular contaminants, though these are less common in routine screening.

Primer Design Considerations

Primer design for mycoplasma detection requires balancing broad reactivity against specificity. The primers must anneal to conserved sequences present in all relevant mycoplasma species—including Mycoplasma, Acholeplasma, Ureaplasma, and Spiroplasma genera—but must not anneal to human, mouse, or common bacterial DNA. In practice, this means selecting primers against regions of the 16S rRNA gene that are conserved among Mollicutes but contain mismatches with eukaryotic 18S rRNA and with the 16S rRNA of common environmental bacteria such as Escherichia coli and Pseudomonas species.

The annealing temperature is a critical parameter. Typical primers for mycoplasma detection have melting temperatures (Tm) between 55°C and 65°C, and the PCR annealing step is performed at 55°C to 60°C. Lower annealing temperatures increase sensitivity but also increase the risk of non-specific amplification; higher temperatures improve specificity but may reduce sensitivity for species with minor sequence mismatches. Most published protocols use an annealing temperature of 55°C with 35 to 40 cycles to maximize sensitivity. The expected amplicon size for 16S rRNA-based assays is typically 250 to 500 base pairs, which is easily resolved on a 1.5% to 2% agarose gel.

Sample Preparation for Mycoplasma PCR

The quality of the sample preparation step determines the reliability of the entire assay. Mycoplasma PCR can be performed directly on cell culture supernatant, but this approach is less sensitive than extracting DNA first, because the supernatant contains only a fraction of the total mycoplasma population—many organisms adhere tightly to the host cell membrane. For maximum sensitivity, both cells and supernatant should be collected and processed together.

DNA Extraction from Cell Culture

The first step is to collect a sample that represents the entire culture. For adherent cells, trypsinize the monolayer and collect the cells along with the culture medium. For suspension cultures, simply collect an aliquot of the cell suspension. Centrifuge 1 to 2 mL of the sample at 200 × g for 5 minutes to pellet the host cells, then transfer the supernatant to a fresh tube and centrifuge again at 16,000 × g for 10 minutes to pellet any free mycoplasmas. Combine the host cell pellet and the high-speed pellet for DNA extraction.

Several DNA extraction methods are compatible with mycoplasma PCR. Boiling lysis is the simplest: resuspend the combined pellet in 50 to 100 µL of lysis buffer containing 10 mM Tris-HCl (pH 8.0), 1 mM EDTA, and 0.5% (v/v) Nonidet P-40 or Triton X-100, then heat at 95°C for 10 minutes. This releases DNA from both host cells and mycoplasmas, and 2 to 5 µL of the lysate can be used directly as template. Boiling lysis is fast and inexpensive, but the crude lysate may contain PCR inhibitors.

Column-based kits, such as those using silica membrane binding, provide cleaner DNA and remove most inhibitors. These kits typically involve proteinase K digestion at 56°C for 10 to 30 minutes, binding of DNA to the column in the presence of chaotropic salts, washing with ethanol-based buffers, and elution in 30 to 50 µL of low-salt buffer or water. The purified DNA can be stored at −20°C for months and used repeatedly. For maximum sensitivity, elute in the smallest volume recommended by the manufacturer to concentrate the DNA.

Positive and Negative Controls

Every mycoplasma PCR run must include proper controls to validate the results. A negative control consisting of water or lysis buffer instead of template DNA confirms that the PCR reagents are not contaminated with mycoplasma DNA. A positive control, using DNA from a known mycoplasma species such as M. orale or a plasmid containing the target sequence, confirms that the PCR reaction worked and that the detection limit is adequate.

A crucial additional control is a spike-in control to detect PCR inhibition. Add a known amount of mycoplasma DNA (or an internal amplification control) to a parallel reaction containing the test sample. If the spike-in control fails to amplify, the sample contains inhibitors that must be removed by further purification or dilution. Some commercial kits include an internal control that is co-amplified with the mycoplasma target using a different fluorophore in real-time PCR, allowing simultaneous detection of inhibition and mycoplasma DNA in a single reaction.

PCR Assay Formats

Three PCR formats are commonly used for mycoplasma detection: conventional PCR with gel electrophoresis, nested PCR, and real-time PCR (quantitative PCR, qPCR). Each has distinct advantages and limitations in terms of sensitivity, specificity, cost, and turnaround time.

Conventional PCR

Conventional PCR is the simplest and most accessible format. A single round of amplification is performed with one primer pair, and the products are analyzed by agarose gel electrophoresis. The presence of a band of the expected size indicates a positive result. This method is inexpensive, requires only a thermal cycler and gel electrophoresis equipment, and is suitable for laboratories with modest throughput.

The sensitivity of conventional PCR for mycoplasma detection is typically 10 to 100 genome copies per reaction, which corresponds to roughly 10³ to 10⁴ colony-forming units (CFU) per milliliter of culture. This is adequate for detecting established contamination, which usually reaches titers of 10⁶ to 10⁸ CFU/mL, but may miss low-level or early-stage infections. The specificity depends on the primer pair and annealing conditions; non-specific bands can sometimes appear, requiring careful interpretation.

Nested PCR

Nested PCR uses two sequential rounds of amplification with two different primer pairs. The first round uses outer primers that flank a larger region; the second round uses inner primers that anneal within the first amplicon, using a small aliquot of the first reaction as template. This approach dramatically increases sensitivity because any DNA amplified in the first round is re-amplified in the second, and it also improves specificity because the inner primers must recognize sequences that were already amplified by the outer primers.

Nested PCR can detect as few as 1 to 10 genome copies, making it the most sensitive PCR format for mycoplasma detection. However, it has significant drawbacks. The two-step procedure doubles the hands-on time and increases the risk of carryover contamination, because opening the first reaction tube to transfer product to the second reaction creates an opportunity for aerosolized amplicons to contaminate the laboratory. For this reason, nested PCR is being replaced by real-time PCR in most modern laboratories.

Real-Time PCR

Real-time PCR (qPCR) monitors amplification in real time using fluorescent probes or DNA-binding dyes. The most common chemistry for mycoplasma detection is the hydrolysis probe (TaqMan) method, in which a probe labeled with a reporter fluorophore and a quencher anneals to the target sequence between the forward and reverse primers. During extension, the 5'→3' exonuclease activity of the DNA polymerase cleaves the probe, separating the reporter from the quencher and generating a fluorescence signal proportional to the amount of amplicon produced.

SYBR Green-based qPCR is an alternative that uses a dye binding to double-stranded DNA. It is less expensive than probe-based assays but less specific, because SYBR Green binds to any double-stranded product, including primer-dimers and non-specific amplicons. Melting curve analysis can distinguish specific from non-specific products, as discussed below.

Real-time PCR offers several advantages over conventional and nested PCR. It is quantitative, allowing estimation of the mycoplasma load. It is performed in a closed-tube system, eliminating the risk of amplicon carryover contamination. It has sensitivity comparable to nested PCR (1 to 10 genome copies) when optimized. And it provides results in 1 to 2 hours, including sample preparation. The main disadvantage is the higher cost of reagents and the need for a real-time thermal cycler.

Feature	Conventional PCR	Nested PCR	Real-Time PCR
Sensitivity (genome copies)	10–100	1–10	1–10
Time to result	3–4 hours	5–6 hours	1–2 hours
Risk of carryover contamination	Moderate	High	Low
Quantitative	No	No	Yes
Cost per reaction	Low	Moderate	High
Equipment required	Thermal cycler, gel rig	Thermal cycler, gel rig	Real-time thermal cycler
Closed-tube format	No	No	Yes

Interpreting PCR Results

Correct interpretation of PCR results is essential to avoid both false positives and false negatives. The interpretation method depends on the PCR format used.

Gel Electrophoresis Analysis

For conventional and nested PCR, the products are separated on a 1.5% to 2% agarose gel containing a DNA-binding dye such as ethidium bromide or a safer alternative like GelRed. A positive result is the presence of a discrete band at the expected molecular weight, typically 250 to 500 base pairs for 16S rRNA-based assays. The positive control lane must show a band of the expected size; the negative control lane must show no band. If the negative control shows a band, the entire run is invalid because reagent contamination is suspected.

Faint bands near the expected size require careful judgment. A faint band in a sample that was expected to be negative may represent low-level contamination, but it could also be a non-specific amplification product. Repeating the assay with a higher annealing temperature or using a nested or probe-based format can resolve this ambiguity. Bands of unexpected sizes may indicate primer-dimer artifacts (usually less than 100 base pairs) or non-specific annealing to host DNA, and they should not be interpreted as positive results unless confirmed by sequencing or a second assay.

qPCR Melting Curves

In SYBR Green-based qPCR, specificity is assessed by melting curve analysis. After the amplification cycles, the temperature is gradually increased while fluorescence is monitored. As the double-stranded DNA denatures, the SYBR Green dye is released, causing a drop in fluorescence. The melting temperature (Tm) of a specific amplicon is characteristic of its length and GC content. A single, sharp melting peak at the expected Tm indicates a specific product. Multiple peaks or a broad peak suggest non-specific amplification or primer-dimers, and such results should be considered suspect.

For probe-based qPCR, specificity is conferred by the probe itself, which must anneal perfectly to the target sequence to produce fluorescence. Non-specific amplification does not generate a signal because the probe does not bind to non-target sequences. The amplification curve should show a characteristic exponential increase in fluorescence, with the cycle threshold (Ct) value inversely proportional to the initial amount of mycoplasma DNA. A sample with a Ct value below 40 is generally considered positive, while samples with no amplification after 40 cycles are considered negative. Some laboratories use a cutoff of 38 cycles to reduce the risk of false positives from low-level contamination.

Common Pitfalls in Mycoplasma PCR

Even well-designed mycoplasma PCR assays can fail if common pitfalls are not addressed. The most frequent problems are contamination of reagents, PCR inhibition, and false negatives due to inadequate sample collection or primer mismatches.

Contamination of PCR Reagents

The most common cause of false positives in mycoplasma PCR is contamination of reagents with mycoplasma DNA or with amplicons from previous reactions. Mycoplasma DNA can be introduced through contaminated reagents, particularly water, buffers, or enzymes used in the PCR. Amplicon contamination occurs when PCR products from a previous run are aerosolized and settle on lab surfaces, pipettes, or reagents. This is a particular risk with nested PCR, where the first-round product is transferred to a second tube. The Aerosol Contamination in PCR article discusses this risk in detail.

To minimize contamination, physically separate the pre-amplification area (where master mixes are prepared) from the post-amplification area (where PCR products are handled). Use dedicated pipettes with aerosol-resistant filter tips, aliquot all reagents into single-use volumes, and include a negative control in every run. If contamination is suspected, replace all reagents, clean all surfaces with 10% bleach followed by 70% ethanol, and use UV irradiation in the PCR hood before preparing new reactions. Note that PCR Specimen Contamination Is Rare when proper technique is followed, but the consequences of even a single contamination event are severe enough to justify rigorous precautions.

PCR Inhibitors

PCR inhibitors are substances that interfere with the activity of the DNA polymerase or with the annealing of primers to the template. Common inhibitors in cell culture samples include heparin (used in some culture media), phenol (from incomplete DNA extraction), ethanol (from column wash buffers), and high concentrations of proteins or polysaccharides. Hemoglobin from red blood cells is a potent inhibitor if the sample contains blood.

The spike-in control described earlier is the most reliable way to detect inhibition. If the spike-in control fails to amplify in the presence of the test sample, the sample contains inhibitors. Solutions include diluting the sample 1:10 or 1:100 in water, re-extracting the DNA using a different method, or adding bovine serum albumin (BSA) to the PCR at a final concentration of 0.1 to 0.5 µg/µL, which binds to inhibitory substances and protects the polymerase.

False Negatives

False negatives can occur for several reasons. The most common is sampling error: if the sample is taken from a culture that has been passaged many times without medium change, the mycoplasma titer may be low because the organisms have been diluted out. Always sample from a culture that has been growing for at least 48 to 72 hours without a medium change, as this allows mycoplasma titers to reach detectable levels.

Another cause of false negatives is primer mismatch. Some mycoplasma species have sequence variations in the 16S rRNA gene that prevent primer annealing. If a particular species is suspected, verify that the primers are compatible with that species' sequence. Finally, some mycoplasmas, particularly M. hyorhinis, are tightly associated with host cell membranes and may be lost if only the supernatant is tested. Always include the cell pellet in the sample, as described in the sample preparation section.

Prevention and Eradication Strategies

Prevention is far more effective than treatment when it comes to mycoplasma contamination. Once a culture is infected, eradication is difficult and often unsuccessful, and the safest course is usually to discard the contaminated culture and replace it with a new one from a verified mycoplasma-free source.

Good Cell Culture Practices

The most important preventive measure is to maintain a strict barrier between contaminated and uncontaminated cultures. Use only mycoplasma-tested reagents, including serum, trypsin, and media. Quarantine all new cell lines for at least two weeks and test them for mycoplasma before introducing them into the main laboratory. Never share media or reagents between cell lines, and use separate bottles for each cell line.

Aseptic technique is critical. Mycoplasmas are transmitted primarily through aerosols and droplets, so work in a laminar flow hood and avoid splashing or vigorous pipetting. Wipe all surfaces with 70% ethanol before and after use, and disinfect the hood with UV light when not in use. Regularly test the incubator water pan and the water bath, as these are common reservoirs of mycoplasma contamination. The Environmental Contamination PCR article provides additional context on monitoring laboratory environments.

Routine testing is the cornerstone of prevention. Test all cell lines monthly, and test any new line before it enters the laboratory. Test any line that has been in contact with a contaminated line, even if the contact was indirect. Many laboratories also test cryopreserved stocks before freezing and after thawing, to ensure that the banked cells are clean.

Antibiotic Treatment

Treatment of contaminated cultures with antibiotics is possible but not recommended for routine use. The most effective regimen uses a combination of a macrolide (such as clarithromycin or azithromycin) and a quinolone (such as ciprofloxacin or enrofloxacin), sometimes with the addition of a tetracycline. These antibiotics inhibit protein synthesis or DNA gyrase in mycoplasmas, which lack the cell wall that would otherwise protect them from many drugs.

A typical treatment protocol involves culturing the cells in medium containing 10 µg/mL ciprofloxacin for 14 days, with a medium change every 3 to 4 days. After treatment, the cells are cultured in antibiotic-free medium for at least two weeks and then tested again by PCR. If the test is negative, the culture is considered cured. However, antibiotic treatment often selects for resistant mycoplasma strains, and the treatment itself can alter cell physiology. The Test of Cure Mycoplasma article discusses the importance of confirming eradication after treatment.

Given the high failure rate and the risk of selecting resistant strains, most experts recommend discarding contaminated cultures and replacing them from a verified clean source. This is particularly true for valuable or irreplaceable cell lines, where the risk of losing the line to antibiotic toxicity or resistance outweighs the benefit of attempted salvage.

Regulatory and Quality Control Considerations

Mycoplasma testing is not just a matter of good laboratory practice; it is a regulatory requirement in many contexts, particularly in the production of biopharmaceuticals and cell-based therapies.

Regulatory Guidelines

The major pharmacopoeias—including the United States Pharmacopeia (USP), the European Pharmacopoeia (Ph. Eur.), and the Japanese Pharmacopoeia (JP)—require mycoplasma testing of cell banks, viral seeds, and final products derived from cell cultures. The Ph. Eur. Chapter 2.6.7 and the USP Chapter 63 describe the approved methods, which traditionally include the culture method (broth and agar) and the indicator cell culture method (using Vero cells and Hoechst staining). These methods are sensitive but slow, requiring 28 days for a definitive result.

PCR is increasingly accepted as an alternative or complementary method. The Ph. Eur. 2.6.7 includes a section on nucleic acid amplification techniques (NAT) for mycoplasma detection, and the USP has proposed similar language. However, PCR methods must be validated to demonstrate that they are at least as sensitive and specific as the compendial methods. Validation typically involves testing a panel of mycoplasma species, determining the limit of detection, and demonstrating that the method is not affected by the sample matrix.

Validation of PCR Methods

Validation of a mycoplasma PCR method for regulatory use requires several parameters to be established. The limit of detection (LOD) must be determined for each relevant species, typically using serial dilutions of a reference strain. The LOD is usually expressed in CFU/mL or genome copies/mL, and for regulatory acceptance, the PCR method should have a sensitivity of at least 10 CFU/mL or 100 genome copies/mL, depending on the matrix.

Specificity must be demonstrated by testing the method against a panel of non-mycoplasma organisms, including common cell culture contaminants such as E. coli, Staphylococcus aureus, and Pseudomonas aeruginosa, as well as host cell DNA. The method must not produce false positives with these samples. Robustness is assessed by varying parameters such as annealing temperature, primer concentration, and MgCl₂ concentration, and confirming that the method still performs within acceptable limits.

Finally, the method must be shown to be compatible with the sample matrix. For biopharmaceutical production, this means testing the method with the specific cell line, culture medium, and downstream processing buffers used in the facility. Matrix effects can cause inhibition or reduced sensitivity, and the validation must demonstrate that the method is reliable under the actual conditions of use.

Practical Summary and Best Practices

Successful mycoplasma PCR testing requires attention to every step of the process, from sample collection to result interpretation. The following best practices will maximize the reliability of your results.

First, collect samples from cultures that have been growing for at least 48 to 72 hours without a medium change, and include both cells and supernatant in the sample. Second, use a validated DNA extraction method that removes inhibitors, and always include a spike-in control to detect inhibition. Third, run positive and negative controls in every PCR batch, and repeat the assay if either control fails. Fourth, use a real-time PCR format with a probe-based chemistry for maximum sensitivity and minimal contamination risk. Fifth, interpret results conservatively: a positive result should be confirmed by a second assay or by sequencing before taking drastic action. Sixth, test all cell lines monthly, and test any new line before it enters the laboratory. Finally, if a culture is positive, discard it and replace it from a verified clean source rather than attempting antibiotic treatment.

Routine monitoring is the single most effective strategy for managing mycoplasma contamination. The cost of regular PCR testing is trivial compared to the cost of lost experiments, invalidated data, and the potential regulatory consequences of releasing contaminated products. By integrating PCR-based mycoplasma testing into your laboratory's standard operating procedures, you protect both the integrity of your research and the safety of any products derived from your cells.

Frequently Asked Questions

What is mycoplasma contamination PCR?

Mycoplasma contamination PCR is a molecular biology technique that detects the presence of mycoplasma DNA in cell cultures or biological products. It uses the polymerase chain reaction to amplify conserved regions of the mycoplasma genome, such as the 16S rRNA gene, and the amplified product is detected by gel electrophoresis or real-time fluorescence. It is the most rapid and sensitive method for routine mycoplasma screening.

How does PCR detect mycoplasma?

PCR detects mycoplasma by amplifying specific DNA sequences that are conserved across mycoplasma species. The reaction uses a heat-stable DNA polymerase, typically Taq polymerase, along with forward and reverse primers that flank the target region. Each cycle of denaturation (94–95°C), annealing (55–60°C), and extension (72°C) doubles the amount of target DNA, and after 35–40 cycles, even a single genome copy can be amplified to detectable levels.

What are the common targets for mycoplasma PCR?

The most common target is the 16S ribosomal RNA gene, which is present in all mycoplasmas and contains conserved regions suitable for primer design. Other targets include the 16S-23S rRNA intergenic spacer region, the tuf gene encoding elongation factor Tu, and the gap gene. The 16S rRNA gene is preferred for broad-spectrum detection because it is conserved across species while still allowing species discrimination through sequencing or restriction analysis.

What is the sensitivity of PCR for mycoplasma detection?

The sensitivity depends on the PCR format. Conventional PCR can detect approximately 10 to 100 genome copies per reaction, corresponding to 10³ to 10⁴ CFU/mL of culture. Nested PCR and real-time PCR are more sensitive, detecting 1 to 10 genome copies per reaction, which corresponds to roughly 10 to 100 CFU/mL. This is sufficient to detect established contamination, which typically reaches titers of 10⁶ to 10⁸ CFU/mL.

Can PCR detect all mycoplasma species?

No single primer pair can detect every mycoplasma species with equal efficiency. Most PCR assays are designed to detect the common cell culture contaminants, including Mycoplasma arginini, M. hyorhinis, M. orale, M. fermentans, and Acholeplasma laidlawii. Some assays also detect Ureaplasma and Spiroplasma species. However, rare or novel species with sequence mismatches in the primer binding sites may escape detection. For comprehensive coverage, some laboratories use multiple primer sets or a degenerate primer approach.

How do I avoid false positives in mycoplasma PCR?

False positives are most commonly caused by contamination of reagents with mycoplasma DNA or with amplicons from previous reactions. To avoid this, use dedicated pipettes with filter tips, physically separate pre- and post-amplification areas, aliquot reagents into single-use volumes, and include a negative control in every run. If contamination is detected, replace all reagents and thoroughly clean the workspace with bleach and UV irradiation.

What should I do if my cell culture is mycoplasma positive?

The safest course is to discard the contaminated culture and replace it from a verified mycoplasma-free source. If the culture is irreplaceable, antibiotic treatment with a combination of a macrolide and a quinolone may be attempted, but this is often unsuccessful and can select for resistant strains. After any treatment, culture the cells in antibiotic-free medium for at least two weeks and confirm eradication by PCR before resuming normal use.

How often should I test for mycoplasma?

Test all cell lines at least monthly, and test any new line before it enters the laboratory. Test any line that has been in contact with a contaminated line, even if the contact was indirect. Test cryopreserved stocks before freezing and after thawing. Laboratories that handle many cell lines or that produce cells for clinical use should test more frequently, and some facilities test every batch of cells before use.

Key Takeaways

Mycoplasma contamination is a silent and widespread problem in cell culture, affecting 5–30% of continuous cell lines, and it can invalidate experimental results without producing visible signs.
PCR detection targets conserved genes such as the 16S rRNA gene, using primers that amplify mycoplasma DNA but not host or common bacterial DNA.
Real-time PCR with probe-based chemistry offers the best combination of sensitivity, specificity, and low contamination risk, detecting as few as 1–10 genome copies.
Proper sample preparation, including collection of both cells and supernatant and removal of PCR inhibitors, is essential for reliable results.
Every PCR run must include positive and negative controls, and a spike-in control to detect inhibition.
False positives are usually caused by reagent contamination; false negatives are usually caused by sampling error, PCR inhibition, or primer mismatch.
Prevention through good cell culture practice and routine monthly testing is far more effective than antibiotic treatment, which is unreliable and can select for resistant strains.
Regulatory guidelines from the Ph. Eur. and USP require mycoplasma testing for biopharmaceutical production, and PCR methods must be validated to demonstrate sensitivity, specificity, and matrix compatibility.