Preventing False Positive and False Negative PCR Results

False negative and false positive results are unfavorable outcomes of the remarkably high sensitivity and specificity of PCR tests, and they can have serious consequences in clinical testing. False negatives can lead to a missed or late diagnosis, putting a patient’s health and survival at risk, while false positives can result in unnecessary additional tests and treatment, including unnecessary costs.

In the case of a contagion, such as COVID-19, a false negative can result in an infected person freely contacting uninfected individuals, rather than being quarantined and treated, accelerating the spread of the pathogen. This is a particular problem when a lab is using a group testing or pooled testing method to save time and resources. When using this method, samples from a number of patients are mixed into one test pool, and the individual samples are only tested when a positive result for the pool is discovered. Therefore, a false negative result from group testing could mean several patients remain undiagnosed.

A false positive result more often results in wasted time and medical resources and may cause unnecessary psychological distress to the person being diagnosed. However, the life of the patient can also be put at risk in some cases. A false diagnosis of Lyme’s disease, probably resulting from sample contamination, reportedly led to the death of a 30-year-old patient who received extensive antibiotic therapy and subsequently developed a Candida complication related to her prolonged catheterization. Fortunately, compared to other methods, false positives and negatives in PCR tests are relatively uncommon, and various steps and safeguards can be taken to prevent, detect, and mitigate these errors.

Preventing false positives

Ensuring good laboratory practice and strict adherence to protocols are fundamental to minimizing erroneous PCR results. The vast majority of false positive results can be avoided using strict laboratory hygiene practices, good pipetting techniques, and sterile labware, and the allocation of dedicated PCR stations or rooms.

Sources of contamination include laboratory personnel and cross-contamination from positive samples or positive controls. Possible routes of contamination are:

Aerosols—microscopic droplets of fluid introduced into the air through spillages, overly energetic pipetting, careless vortexing, high ambient temperatures, etc.
Contaminants on lab coats, watches, hair, skin, etc.
Contaminants in reagents used in the PCR reaction.
Contaminated pipettes, overuse of disposable tips, use of non-sterile tubes, glass wear, etc.

Techniques for preventing contamination

If possible, separate areas or rooms should be allocated to preprocessing, PCR amplification, and post-PCR steps to avoid cross-contamination. Maintaining a unidirectional flow of traffic between these areas is recommended. Further strategies to avoid contaminants moving between the areas include:

The application of fresh gloves before moving from area to area.
The availability of dedicated lab coats for use in each area.
The use of swinging doors that can be opened without using your hands and that also ensure the unidirectional flow of personnel.

Good laboratory practice and aseptic techniques can be used to minimize contaminants affecting PCR.

Although the stringent aseptic techniques used in microbiology are usually unnecessary for PCR, adopting some of the practices can help eliminate contaminant transfer. For example, being able to open and use tubes with one hand and without setting the cap on the bench and preventing pipette tips from touching solid surfaces. If contamination is continuously affecting the results of certain PCRs, despite appropriate measures, more stringent techniques can be applied. Other examples include:

Keeping all sample and reagent vessels sealed for as long as possible and using aseptic techniques when opening bottles and tubes.
Removing watches, wrist bands, and jewelry.
Preparing single-use aliquots of reagents.
Using a laminar flow cabinet.
Using gentle, precise, and controlled pipetting and avoiding spills.
Maintaining dedicated equipment within each area, such as tubes, pipette tips, pipette guns, vortexers, and microcentrifuges.
Autoclaving all tubes, pipettes, etc.
Sterilizing benches and equipment with 70 percent ethanol, 10 percent sodium hypochlorite (minimum 10 min contact time) followed by sterile water, 1 M hydrogen chloride, and/or UV light.
1. In the case of recalcitrant contamination (poorly biodegradable or non-biodegradable chemicals that can include hydrocarbons, pesticides, personal care products, nanomaterials, and some toxins), cleaning with psoralene compound should be considered.
Opening and cleaning pipette guns and micropipettes. The interior of micropipettes and guns can become dirty from improper pipetting techniques and being placed horizontally with a tip full of liquid. This could present a hidden source of contamination. These can be sent to specialized companies for cleaning, servicing, and recalibration. Servicing and recalibration will also ensure the pipettes are accurate and do not contribute to erroneous results.
Wearing masks and regularly changing gloves.

PCR techniques for improving specificity

Although designing longer primers with non-specific, homopolymer, or loop sequences at the 5′ end improves specificity and annealing stability, the universal 5′ end sequence also leads to nonspecific amplification. Annealing-control primers are designed with a polydeoxyinosine linker, which forms a bubble structure during annealing that prevents nonspecific binding, improving specificity; this type of primer is commercially available from Seegene.

Primers should be designed to target genes with low levels of conservation and that are highly specific to the species of interest. The sequences of primers should be regularly verified via BLAST search against the NCBI database to ensure they are specific and unlikely to cross-react with nontarget sequences.

Contamination by previously amplified PCR products, i.e., “carry-over” contamination, may be a particular issue in areas of high-throughput PCR laboratories. To reduce contamination by amplicons, uracil-DNA-glycosylase (UNG) can be added to the PCR reaction. In fact, UNG is often an ingredient in commercially available PCR master mixes.

Targeting longer amplification sequences decreases the likelihood of amplifying an unspecific sequence. The U.S. FDA recommends a minimum target length of 100 base pairs for tests aimed at detecting pathogenic viral genes. The use of primers designed to target sequences that are only 25 bases long has been postulated to be one of the reasons for false positives arising from the U.S. CDC’s rRT-PCR kits for SARS-CoV-2.

Hot start PCR

One of the principles of PCR is that an annealing temperature specific to the primer is chosen to promote specific annealing; however, unspecific binding and amplification can occur during PCR preparations at room temperature and during heating. Therefore, to increase specificity, a modified technique known as hot start PCR uses polymerase enzymes whose function is inhibited until the ideal reaction temperature is reached.

Touchdown PCR

In touchdown PCR, an initial annealing temperature 5°C–10°C above the estimated melting temperature (T_m) of the primer is applied. The temperature is then slowly reduced by 1°C–2°C decrements over several cycles until the optimal temperature is reached. The stringent conditions favor specific primer-template adhesion over nonspecific binding.

Detecting contamination

A no-template control prepared and run simultaneously with the sample reactions can be used to identify sources of contamination. A negative result in this control means that the reagents, pipette tips, water, and the preparation environment are unlikely to be sources of contamination. These controls should be kept as far as possible from positive samples and controls during preparation to avoid cross-contamination.

Note: the presence of amplification products in the negative control may not necessarily indicate contamination when “late amplification” (more than 34 cycles, e.g., in SYBR Green-dye-based assays) is used, as it could also be a result of dimer amplification. Performing melting curve analysis after running the PCR can be used to confirm the presence of primer-dimers. In addition, if a colored or fluorescent probe is being used, a positive result in a negative control sample can be due to the degradation of the probe, which releases signal molecules, leading to high background noise. Signal-to-noise assessment, mass spectrometry, or a fluorometric scan can be used to check for probe degradation.

In the event of contamination

If PCR contamination is confirmed, all reagents, enzymes, etc., should be replaced; all equipment and surfaces carefully sterilized; bottles, tips, glassware, etc., should be autoclaved; and the above-mentioned contamination control measures implemented.

Preventing false negatives

The causes of false negatives are wide-ranging and include degraded or insufficient nucleic acid, such as from excessive freeze-thaw cycles; contamination with inhibitors from the environment or DNases and RNases; poorly designed PCR; low-quality reagents; poor or inconsistent techniques during sample processing; and low-quality, faulty, or poorly calibrated equipment.

Therefore, in addition to steps taken to reduce contamination using aseptic techniques, various other methods can be used to prevent false negatives:

Certain substances known to inhibit PCR, such as powdered gloves and wooden cotton or calcium alginate swabs should be replaced with alternatives.
Samples should be carefully stored at low temperatures, in small aliquots to avoid excessive freeze-thawing, and in suitable nuclease-free buffers.
Inhibition may also be prevented by incorporating 200 to 400 ng/pl of bovine serum albumin into the reaction mix, although this only works for some types of inhibitors, such as phenolic compounds.
Reduce exposure of nucleic acids to RNases and DNases by using nuclease-free water, reagents, and equipment (e.g., pestles, tubes and pipette tips, gloves), and maintaining good laboratory practice.
1. Note that DEPC-treated water is often contaminated with reverse transcription and/or PCR inhibitors and should be avoided for PCR.
Stringent adherence to standard operating procedures, using good quality reagents and equipment, and regular equipment servicing and calibration will aid with reproducibility and sensitivity.

Using controls to identify false negatives

Controls can be designed not only to act as indicators of a (potential) false positive result but also to provide evidence of the causes of a false negative.

To confirm any problems with the enzyme, reagents, and thermal cycling conditions, it is recommended to include an “internal control.” This is usually the amplification of a mammalian housekeeping gene, such as glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which is expressed in patient cells.

An “external control,” involving the amplification of a known sequence of a plasmid vector, can be used to determine the detection limit of the PCR assay and to indicate if nucleic acid has been lost or degraded during purification steps. Blood samples spiked with DNA from a target pathogen (genomic DNA or reverse-transcribed RNA) at concentrations ranging from slightly above the detection limit of the PCR to higher concentrations is an example of a good positive control.