Since the discovery of the polymerase chain reaction (PCR) by the oligonucleotide chemist Kary Mullis in 1983, the method has revolutionized molecular biology and clinical diagnostics.
Before PCR, DNA amplification required multiple steps, including cloning into plasmids, insertion into bacteria, bacterial growth, isolation of plasmid DNA, and separation of inserts from plasmid vectors.
In contrast, PCR is performed in vitro as a single step, requiring only two oligonucleotide primers, a polymerase, and temperature cycling of the DNA template in the presence of deoxyribonucleotides.
Although its spread was initially limited by restrictive patent policies, the basic method is now off patent and has become a democratic cornerstone of molecular biology. Thousands of scientists have contributed to and expanded the methods and applications of PCR, including quantification of transcripts after reverse transcription and PCR followed by cycle sequencing.
Anyone can perform PCR with generic reagents and simple laboratory instruments, amplifying specific DNA segments by 106-to 109-fold for further study in genetics, oncology, and infectious disease.
Room for Improvement
Initially, PCR was slow, expensive, and required further analysis by gel electrophoresis. Even after automated thermal cyclers (PerkinElmer, Life Technologies) and thermally stable polymerases (Roche) were developed in the late 1980s, PCR protocols typically required four hours for amplification.
The feasibility of rapid cycle PCR (30 cycles in 15–30 min) was shown in the early 1990s and commercialized as the RapidCycler (Idaho Technology) with eventual adoption into the real-time capillary LightCycler (Roche) and SmartCycler (Cepheid) platforms. Amplification speed was limited by sample geometry and heat transfer, not reaction chemistry.
Nevertheless, microwell plates became the dominate format, reflecting user preference for convenience and automation over speed and temperature control. A trend toward faster amplification was recently rebranded as Fast PCR, often packaged with presumed enabling reagent modifications.
However, amplification speed remains instrument limited, not reagent or reaction limited.
Similar to the order of magnitude change in speed, the cost of PCR has diminished at least an order of magnitude. After the original Taq polymerase patents were deemed unenforceable, reagent manufacturers turned their focus to improved enzymes and hot-start methods to add value.
Physical separation of critical components with wax, inactivation of the polymerase with an antibody, heat-labile enzymes, primers, and dideoxynucleotides have all been commercialized to improve specificity. Although not necessary for the majority of applications, such improvements may compensate for poor reaction design and execution.
Twenty years ago, reaction volumes were typically 100 µL. Today, 10 µL is commonly used and microfluidic systems run a thousand times smaller yet at 1–10 nL per reaction.
Digital PCR can be performed to take advantage of limited template copies per reaction, or preamplification can expand the sensitivity limits inherent to small volumes. Even picoliter reactions are now routinely formed as emulsions or controlled droplets within oil, enabling massively parallel sequencing by expansion of individual templates or production of target libraries.
To the extent that reagent cost is proportional to reagent volume, the expense of individual reactions should be vanishingly small. However, in the case of digital PCR, many reactions are necessary for quantification, and droplets can only be analyzed individually with expensive equipment. Furthermore, if each reaction requires specific oligonucleotides, the initial synthesis costs remain the same and can be oppressive when many targets are analyzed.
As reaction volumes have decreased, the number of samples run at one time has generally increased. Microtiter trays of 96-wells are standard, and 384-well and 1,536-well instruments are available. A microfluidic matrix of 96 assays and 96 samples amplifies over 9,200 individual reactions on one chip (Fluidigm). Bridge amplification tethers PCR primers to a surface and can amplify millions of PCR clones prior to massively parallel sequencing. Of course, more reactions are not necessarily better and value does not always scale linearly with the number of samples.
More important than speed, cost, and batch size is the information (both qualitative and quantitative) that PCR enables. Historically, gel analysis was performed after PCR for size analysis of the products. Such “open” analysis provides flexibility but also the opportunity (or some would say, guarantee) that PCR products eventually contaminate the PCR reagents.
In contrast, real-time PCR is homogeneous and closed without any separation or processing steps. Quantitative information on the amount of initial template is obtained during PCR by monitoring fluorescence each cycle, correlating the log of the initial template concentration to the quantification cycle (Cq).
The term Cq replaces Ct (cycle threshold) and Cp (crossing point) as introduced by MIQE (Minimum Information for Quantitative PCR Experiments), an attempt to standardize and provide guidelines for performance and interpretation. Quantitative reverse transcriptase PCR remains the gold standard for transcript quantification and is often used to validate results obtained by other methods.
Fluorescent indicators that correlate with the amount of PCR product are necessary for real-time PCR. Indicators can either be dyes that change fluorescence in the presence of double-stranded DNA or oligonucleotide probes that change fluorescence by hybridization and/or hydrolysis.
Functional classification of real-time probes as hybridization or hydrolysis probes is more useful than trademarks. Hybridization probes reversibly change fluorescence with hybridization and typically consist of one or two probes per target.
Single hybridization probes change fluorescence after a conformational change of the probe or hybridization to the target. Dual hybridization probes change fluorescence by approximation of two fluorophores and resonance energy transfer. Hydrolysis probes generate fluorescence irreversibly by polymerase cleavage of the probes between two fluorophores.
Most fluorescent indicators are not linearly correlated with the amount of product, but this does not compromise their function in quantitative PCR. The cumulative signal from hydrolysis probes may provide a sensitivity advantage, but the ability for reversible melting analysis (characteristic of hybridization probes) is lost.
Probe-melting analysis, which is popular for clinical genotyping, partly because multiple variants can be identified under one probe, was used in the first FDA-approved genetics tests for F5 and F2 genotyping in 2002.