INTRODUCTION — The polymerase chain reaction (PCR) is the basis of many modern molecular biology and molecular genetics techniques. In just a few hours, PCR can amplify a single DNA molecule a million-fold [1]. The greatly amplified target DNA is subsequently analyzed via other techniques.
This topic presents a brief overview of PCR as well as a discussion of some applications of PCR and quantitative PCR. Other molecular genetic tools are discussed separately.
●(See "Tools for genetics and genomics: Cytogenetics and molecular genetics".)
●(See "Tools for genetics and genomics: Gene expression profiling".)
●(See "Tools for genetics and genomics: Specially bred and genetically engineered mice".)
●(See "Tools for genetics and genomics: Model systems".)
THE PCR PROCESS — Since its first publication in 1985 [2], the impact of PCR on biomedical research has been immense. This technology allows large quantities of rare sequences to be synthesized, cloned, and analyzed with high reliability and minimum effort. The award of the 1993 Nobel Prize in Chemistry to Kary B Mullis for inventing the technique recognized the importance of PCR-based methods [3].
PCR allows for rapid and highly specific amplification of DNA fragments. The method is relatively inexpensive and commonly performed in most molecular laboratories. Pieces of DNA from about 50 base pairs (bp) to over 10 kilobases (kb) can be amplified, even from vanishingly small amounts of starting genomic DNA. The two most important principles underlying PCR are:
●Complementarity-driven binding of single DNA strands to form a duplex
●Template-driven, semi-conservative synthesis of DNA, by DNA polymerases
These two principles are discussed separately in detail. (See "Basic genetics concepts: DNA regulation and gene expression".)
PCR is typically a single tube reaction, with each reaction customized to amplify one specific genomic region (a specific sequence). This specificity is dictated by the pair of oligonucleotide primers (short DNA sequences) designed to hybridize to flanking regions of the target sequence.
The standard PCR reaction mixture consists of a large excess of oligonucleotide primer pairs, a template DNA (typically genomic DNA), free deoxynucleotide triphosphates (DNA bases), reaction buffer, and thermostable DNA polymerase (the enzyme that drives the PCR reaction).
Thermostable DNA polymerases can withstand heating to 95°C with minimal loss of activity and function optimally near 70°C [1]. The standard enzyme currently used in most PCR is derived from the bacterium Thermus (also Thermophilus) aquaticus (Taq). The error rate (per nucleotide per cycle) for standard Taq is about 1 in 105 nucleotides. High-fidelity polymerases with in vitro proofreading ability, such as those derived from the archaebacterium Pyrococcus furiosus (Pfu), have error rates that are approximately 10 times lower than Taq and are used for applications that mandate more stringent DNA synthesis [4].
The PCR consists of three stages that are repeated 30 to 40 times (figure 1):
●Denaturing – The mixture is heated to 95°C to allow the double-stranded template DNA to denature into single strands.
●Annealing – The mixture is cooled to a temperature just below the predicted primer pair melting temperatures, resulting in primer binding to the single-stranded DNA templates, followed by binding of DNA polymerase to the 3’ end of the primers.
●Elongation – The temperature is then raised to a polymerase activation temperature (~70 to 72°C) to initiate chain elongation. The polymerase catalyzes the addition of free nucleotides to the 3’ ends of the primers. Bases are added to complement the template sequence (A with T, C with G). Elongation continues for about one minute and is terminated by cycling back to step one of the reaction: heating the temperature to 95°C, resulting in strand separation.
Amplification is exponential. The number of potential targets for primer annealing doubles with each PCR cycle, and the amplified sequences (amplicons) produced in one cycle serve as templates for subsequent cycles. It is possible to start with as little as one template DNA molecule and generate tens of millions of copies. If the reaction starts with two copies, then N cycles will theoretically yield 2N copies.
To amplify the sequence of interest, single-stranded oligonucleotide primers (which are typically 20 nucleotides in length) must be carefully designed and synthesized. Highly specific amplification can be achieved if the sequence combination of the primer pairs is present only once in the genome. Software that facilitates primer design and calculates annealing temperatures are freely distributed online [5].
After the reaction is completed, the PCR product can be visualized by gel electrophoresis: negatively charged DNA fragments migrate through the porous agarose gel toward a positive electrode, with the distance traveled determined by their molecular weight. After electrophoresis and addition of a fluorescent molecule that binds to DNA and forms a complex with nucleic acids, exposure of the gel to ultraviolet light demonstrates the amplified fragment as a single fluorescing band. The gel can be photographed for a permanent record. For high-throughput applications in commercial or genomics laboratories, this sorting is accomplished using capillary-based electrophoretic methods.
Compared to alternative DNA amplification methods, PCR offers the following advantages:
●Rapidity – The amplification reaction takes approximately three hours. Additional time (several hours) is required to first extract DNA from cells and then perform gel electrophoresis after completion of PCR. Single tube reactions enable more rapid sample processing and point-of-care testing.
●Sensitivity – PCR can amplify the DNA from a single cell and thus is extremely sensitive, sufficiently so that it can be used in preimplantation genetic diagnosis in embryos.
●Robustness – Degraded DNA can frequently be successfully amplified, including from archived tissue blocks.
●Specificity – The technique also permits the detection of small nucleotide mutations (see below), as well as trinucleotide repeat expansions.
●Low cost and ease of use – Low cost and ease of use, including automation, allow PCR to be used in numerous clinical applications.
There are several disadvantages to the PCR technique:
●Sequence-dependency – To design appropriate PCR primers, the nucleotide sequence information of the region of interest must be known. This limits its use when attempting to isolate and amplify novel gene sequences.
Error rates – In contrast to intracellular amplification methods (bacterial cloning), in vitro PCR lacks DNA proofreading machinery. Though adequately low for most experimental applications, the resultant error rates of PCR (on the order of 10-5) are too high for therapeutic applications.
●Size-limitation – The ideal length of a sequence to be amplified (PCR amplicon) is several hundred nucleotides. Although long-range PCR methods are available to amplify 5 to 10 kb, they are susceptible to higher error rates and are less robust. Similarly, large mutations such as duplications or deletions that encompass the amplified region cannot be detected because only the unaffected allele will be amplified.
●Contamination risk – Because of the extreme sensitivity of PCR to detect DNA sequence, contamination of the sample with small amounts of extraneous DNA may result in a spurious or false-positive finding. To help decrease this problem, diagnostic PCR laboratories are often subdivided into separate, designated pre- and post-amplification areas in different rooms.
UNIQUE GENOMIC SEQUENCES — Although the human genome sequence is very long (3 billion bases), relatively short DNA sequences suffice to specify a unique sequence in the genome. The expected count of a specified sequence of length N bases can be expressed by the following formula, based on probability theory:
(3 X 109) x 1/4N
1/4N is the probability of a given sequence of length N bases, and the constant 3 X 109 is the number of bases in the human genome.
Thus, PCR primers of 20 and 25 nucleotides in length would be estimated to occur by chance at a frequency of 0.003 and 0.000003 per genome, respectively. Given that PCR requires two primers to colocalize to a relatively small genomic region with a specific orientation to each other (one on each DNA strand, facing each other), the probability of amplifying more than one DNA region is exceedingly small.
Sequence alignment algorithms can be useful to align designed primer pairs to the known genome sequence, to verify if a primer pair is unique in the genome, to check if any polymorphisms underlie the primers, and to calculate the exact length of the expected amplicon [6].
PCR COMPARED TO OTHER AMPLIFICATION METHODS — The exponential amplification of PCR can be contrasted with other types of nucleic acid amplification that are widely used in molecular biology. Two examples of other amplification methods are whole genome amplification and RNA amplification.
Whole genome amplification — Whole genome amplification (WGA) non-specifically amplifies the entire genome and is used when sample quantities are too small for multiple PCR reactions or to meet the desired application's requirements. Random, very short primers are used, along with Phi29 polymerase, a highly active polymerase capable of incorporating over 70,000 nucleotides per priming event [7]. In order to limit amplification bias (uneven amplification of different genomic sequences or alleles), it is best to amplify the template DNA only a few 1000-fold.
WGA is distinct from standard PCR in that it is non-specific (all genomic sequences are targeted, not only one site), and it generates fragments that are tens of thousands of bases in length, compared to 250 to 1000 bases that are typical for PCR. Often the fragments generated by WGA are short, thus precluding long-range PCR or applications that require long template strands.
RNA amplification — Linear amplification of RNA is the precursor to most gene expression microarray experiments. Linear amplification is preferred over exponential amplification (ie, classical PCR) in this setting because occasional imbalances in sequence amplification rates (that result from preferential reaction kinetics and favor amplification of some regions over others) can bias subsequent quantification [8]. The general method for global amplification of mRNA is based upon the Eberwine protocol [9], whereby single-stranded RNA is the template, the reaction is primed by a poly-T oligonucleotide primer that binds the 3’ poly-A tail common to most mRNA sequences, and the reaction is catalyzed by a reverse transcriptase which generates complementary DNA strands. Following second strand synthesis with DNA polymerase, and subsequent sample purification, T7 polymerase is used to drive in vitro transcription (IVT), producing multiple copies of amplified RNA (aRNA) from each dsDNA. By incorporating labeled nucleotides in the in vitro transcription step, amplification products to be used as probes in expression analysis can also be prepared. Amplification achieved is typically in the range of 10- to 100-fold, but can vary depending on the amount of input RNA.
CLINICAL APPLICATIONS OF PCR — Though the applications of PCR are innumerable, its clinical utility is greatest in the context of genotyping and sequencing for diagnostic and predictive testing.
Genotyping and sequencing — Genotyping is the process of characterizing an individual’s genotype (the combination of alleles) at a particular genomic locus (location). Most types of genetic variation can be characterized using PCR techniques, including microsatellite and single nucleotide polymorphism (SNP) markers, insertion/deletion variants (indels), and some structural variants, such as copy-number variants.
SNP genotyping — PCR is the basis of the most commonly used single-nucleotide polymorphism (SNP) genotyping methods, including probe-hybridization based assays (such as the Taqman SNP assay) [10], allele-specific PCR [11,12], and minisequencing assays, such as those implemented in genome-wide SNP microarrays [13]. These and other methods have been adopted for clinical use in CLIA-certified laboratories.
DNA sequencing is the process of characterizing the nucleotide (base-pair) sequence of a specific DNA sequence. Clinically, sequencing is used to identify the pathogenic mutations in individuals with genetic disorders caused by rare variants at a specific locus. An example is resequencing of the BRCA1 locus in individuals at risk for familial breast cancer. Single gene clinical sequencing is PCR-based, with each assay focused on a specific exon or gene region. While some next-generation sequencing platforms are often PCR-free, PCR-based amplification of DNA libraries are frequently used prior to sequencing in whole genome sequencing applications. (See "Next-generation DNA sequencing (NGS): Principles and clinical applications".)
Detection of rare sequences — PCR technology also allows detection of rare sequences in a population of DNA or RNA molecules. This application is particularly useful in searching for DNA rearrangements in the setting of neoplasia. An example is the discovery that a Herpes simplex–related virus is involved in the pathogenesis of Kaposi's sarcoma [14-16].
Rare sequence detection by PCR is used routinely in obstetrics for preimplantation testing and non-invasive prenatal testing (NIPT). Preimplantation testing (whereby established embryos are screened for suspected genetic mutations prior to uterine implantation) is demanding because PCR must be performed from a single cell, removed from the very early embryo. The major concerns in single-cell PCR are preventing allele dropout and contamination with maternal or other outside DNA [17]. NIPT relies on detection of free fetal DNA (ffDNA) in maternal blood; since ffDNA makes up only a small percentage of the circulating DNA, pains must be taken to differentiate fetal from maternal sequences [18]. (See "Prenatal screening for common aneuploidies using cell-free DNA".)
Detection of infectious organisms — Clinical assays are being developed that would allow PCR-based identification of common infectious pathogens including bacteria and fungi. An example is a PCR-based panel that detects Enterococcus faecium, Staphylococcus aureus, Klebsiella pneumoniae, Pseudomonas aeruginosa, and Escherichia coli (so-called ESKAPE organisms) [19]. Magnetic beads that bind to the PCR products are detected using magnetic resonance. In a series of hospitalized patients with suspected bacterial sepsis, this test was able to positively identify an infectious organism within 4 to 8 hours, versus 2 to 3 days for most culture results [20]. Compared with culture, the sensitivity and specificity of the PCR-based test were both 90 percent. The PCR test was also able to identify infectious organisms that were not detected by blood cultures in a small number of patients. As noted by an editorialist, this type of testing could be a useful adjunct to culture in obtaining more rapid results but would not reduce the need for culture because the PCR assay does not identify organisms other than the five tested and does not provide antibiotic sensitivities [21].
Next-generation DNA sequencing (NGS) as a means of identifying infectious pathogens may replace PCR-based tests as NGS can survey a much broader range of organisms simultaneously. Higher cost and longer turnaround time for NGS have slowed its widespread implementation in the acute setting, although its availability is steadily increasing. (See "Next-generation DNA sequencing (NGS): Principles and clinical applications", section on 'Diagnosis of infections'.)
Further information about the clinical uses of gene-based testing in individuals with suspected infections is discussed separately. (See "Detection of bacteremia: Blood cultures and other diagnostic tests".)
Quantifying nucleic acid sequence abundance
RT-PCR — Real-time quantitative PCR (RT-PCR; qPCR) is a highly sensitive method for quantifying the absolute or relative amount of a specific nucleic acid sequence in which the accumulation of PCR products over time is measured directly, without post-PCR modifications. Quantitation is estimated from the cycle number required for the reaction to generate sufficient fluorescence to cross a set threshold. This method is thus reliant on performing multiple rounds of amplification [22]. Common applications of qPCR include gene expression profiling, quantification of viral load, and determination of copy number.
qPCR is performed by quantifying the amount of amplified product with each round of PCR cycling. In addition to the two primers that are necessary for successful PCR amplification, this method frequently applies a fluorescent, non-extendible probe that hybridizes to the target sequence between the primer pair.
This probe contains a fluorogenic reporter dye at its 5' end and a quencher at the 3' end. The quencher blocks fluorescence emission as long as it is in close proximity to the reporter. As the primer extends downstream during amplification, however, the exonuclease activity of the DNA polymerase cleaves away the hybridized probe and removes the quencher. An increase in the fluorescence signal ensues (figure 2).
Due to the exponential nature of the PCR, the fluorescence signal increases proportionally to the amount of generated PCR product until a plateau is reached. Quantification is accomplished by comparing the cycle number at which the patient sample reaches a predetermined level of fluorescence to a standardized curve of a control sample, thus deriving copy number at the start of the reaction (figure 3). This method is rapidly becoming the method of choice for monitoring residual disease in patients receiving chemotherapy and/or hematopoietic cell transplantation for hematologic malignancies [23,24]. (See "Genetic abnormalities in hematologic and lymphoid malignancies".)
Advantages of this method include the following:
●Extreme sensitivity and a wide dynamic range
●Time-efficient, as the method requires no post-PCR processing of samples
●Automated high-throughput instruments (such as the ABI PRISM 7700 Sequence Detection System) are available with interpretation-supporting software
Digital PCR (dPCR) — Digital PCR (dPCR), also known as digital drop PCR, is an adaptation of traditional PCR for the purposes of sample quantification without the need for multiple PCR cycles. The basic principle of dPCR is that a sample is separated (using complex microfluidics) into discrete aliquots (packets), each containing a very small number of nucleic acid sequences from the patient (on average, 1). PCR is then performed in each packet, resulting in a binary result in which the target nucleic acid of interest is either present or absent (in binary terms, either positive [1] or negative [0]). Subsequent modeling (assuming a Poisson distribution) enables estimation of the starting amount of target nucleic acid without the reliance on multiple amplification cycles as required in qPCR. The measured quantity in dPCR is thus considered an absolute measure, rather than the relative measure of traditional PCR (compared to the threshold value of a standard). dPCR is being used in several clinical settings, including mutation detection and the detection of infectious agents [25,26].
Gene expression profiles — Specific gene transcripts can be measured by qPCR from RNA samples that have been converted to cDNA by reverse transcription. Typically, relative quantification is used to calculate fold-change among a set of samples. For example, inflammation-related genes can be assayed by Taqman qPCR, comparing normal tissue with colon cancer samples [27]. qPCR expression profiling is used to validate measurements of specific candidate genes identified through genome-wide expression microarray studies.
Clinical measurement of viral RNA or DNA — PCR of RNA isolated from blood is a standard tool in monitoring the viral load in HIV-infected patients [28,29]. Several available commercial real-time PCR assays provide absolute quantitation of HIV-1 RNA copies per mL of plasma [30]. Key considerations for HIV RNA testing are throughput, automation, accuracy, and dynamic range. PCR quantification has also been applied to other RNA viruses causing chronic infection, such as hepatitis C virus [31,32]. (See "Techniques and interpretation of HIV-1 RNA quantitation" and "Screening and diagnosis of chronic hepatitis C virus infection".)
The 2009 outbreak of swine-origin influenza A (H1N1) prompted rapid development of diagnostic real-time, RT-PCR assays [33]. The assays developed by the Centers for Disease Control consist of four sets of primers and Taqman probes for universal detection of type A influenza, universal detection of swine influenza A, specific detection of swine H1 influenza, and the positive reference control gene, RnaseP [34].
Another area in which qPCR is becoming useful is in the quantitation of Epstein-Barr viral load in the diagnosis and monitoring of patients with post-transplant lymphoproliferative disease [35]. (See "Epidemiology, clinical manifestations, and diagnosis of post-transplant lymphoproliferative disorders", section on 'Measurement of EBV viral load'.)
Monitoring of hepatitis B treatment response by quantifying viral DNA in serum, with real-time PCR, is another important clinical application [36]. (See "Hepatitis B virus: Screening and diagnosis", section on 'Serum HBV DNA assays'.)
CRITICAL EVALUATION OF DATA — The ease of performing PCR has led to wide dissemination of the methodology. However, reports of PCR analyses should be reviewed for validity of the results.
Every PCR experiment requires a minimum of two technical controls (positive and negative controls), in addition to biological controls specific to the research question being addressed. The technical controls should demonstrate that amplification occurs when it should (positive control) and that it does not occur when it should not (negative control). The positive control template should be prepared in the same manner as the test sample, with the positive control DNA typically extracted using the same extraction protocol and starting material as the sample. The negative control (also referred to as a no-template control) involves using all components of the PCR mixture except that water is added in lieu of the DNA template.
The quality of the test sample DNA is critical, as highly fragmented DNA can result in allelic dropout (selective amplification of only one allele at a heterozygous locus) or false-negative results. Impurities in the template DNA sample could inhibit amplification. This is of particular concern in qPCR, since inhibitors will lower reaction efficiency.
SUMMARY
●Method – Polymerase chain reaction (PCR) allows for rapid and highly specific amplification of DNA fragments. PCR uses thermostable DNA polymerases purified or cloned from microorganisms living in hot springs and specific oligonucleotide primers of 20 to 30 residues. The reaction consists of three stages, multiply repeated: denaturing, annealing, and elongation. After the reaction is completed, the amplified product (called an amplicon) can be visualized by gel electrophoresis. Relatively short DNA primer sequences suffice to specify a unique sequence in the genome. (See 'The PCR process' above.)
●Comparison with other amplification methods – The exponential amplification of PCR can be contrasted with other types of nucleic acid amplification that are widely used in molecular biology. Whole genome amplification (WGA) non-specifically amplifies the entire genome and is used when sample quantities are too small for PCR to meet the desired application's requirements. Linear amplification of RNA is the precursor to most gene expression microarray experiments. (See 'PCR compared to other amplification methods' above.)
●Uses
•Most types of genetic variation can be characterized using PCR techniques, including microsatellite and single nucleotide polymorphism (SNP) markers, insertion/deletion variants (indels), and some structural variants such as copy-number variants. (See 'SNP genotyping' above.)
•PCR technology allows detection of rare sequences in a population of DNA or RNA molecules. This application is particularly useful in searching for DNA rearrangements in the setting of neoplasia. Rare sequence detection by PCR also is used routinely in obstetrics for preimplantation testing and non-invasive prenatal testing (NIPT). (See 'Detection of rare sequences' above.)
•Real-time quantitative PCR (qPCR) and digital PCR (dPCR) can be used to quantify the absolute or relative amount of a specific nucleic acid sequence. This method is often used for monitoring residual disease in patients receiving chemotherapy and/or hematopoietic cell transplantation for hematologic malignancies. (See 'Quantifying nucleic acid sequence abundance' above.)
•PCR of RNA isolated from blood is a standard tool in monitoring the viral load in HIV-infected patients. qPCR also has been used for assays of H1N1, Epstein-Barr viral load monitoring post-transplant patients, and monitoring response to therapy for hepatitis B. (See 'Clinical measurement of viral RNA or DNA' above.)
