Digital polymerase chain reaction

Digital polymerase chain reaction (digital PCR, DigitalPCR, dPCR, or dePCR) is a biotechnological refinement of conventional polymerase chain reaction methods that can be used to directly quantify and clonally amplify nucleic acids strands including DNA, cDNA or RNA. The key difference between dPCR and traditional PCR lies in the method of measuring nucleic acids amounts, with the former being a more precise method than PCR, though also more prone to error in the hands of inexperienced users. A 'digital' measurement quantitatively and discretely measures a certain variable, whereas an “analog” measurement extrapolates certain measurements based on measured patterns. PCR carries out one reaction per single sample. dPCR also carries out a single reaction within a sample, however the sample is separated into a large number of partitions and the reaction is carried out in each partition individually. This separation allows a more reliable collection and sensitive measurement of nucleic acid amounts. The method has been demonstrated as useful for studying variations in gene sequences — such as copy number variants and point mutations — and it is routinely used for clonal amplification of samples for next-generation sequencing. Digital polymerase chain reaction (digital PCR, DigitalPCR, dPCR, or dePCR) is a biotechnological refinement of conventional polymerase chain reaction methods that can be used to directly quantify and clonally amplify nucleic acids strands including DNA, cDNA or RNA. The key difference between dPCR and traditional PCR lies in the method of measuring nucleic acids amounts, with the former being a more precise method than PCR, though also more prone to error in the hands of inexperienced users. A 'digital' measurement quantitatively and discretely measures a certain variable, whereas an “analog” measurement extrapolates certain measurements based on measured patterns. PCR carries out one reaction per single sample. dPCR also carries out a single reaction within a sample, however the sample is separated into a large number of partitions and the reaction is carried out in each partition individually. This separation allows a more reliable collection and sensitive measurement of nucleic acid amounts. The method has been demonstrated as useful for studying variations in gene sequences — such as copy number variants and point mutations — and it is routinely used for clonal amplification of samples for next-generation sequencing. The polymerase chain reaction method is used to quantify nucleic acids by amplifying a nucleic acid molecule with the enzyme DNA polymerase. Conventional PCR is based on the theory that amplification is exponential. Therefore, nucleic acids may be quantified by comparing the number of amplification cycles and amount of PCR end-product to those of a reference sample. However, many factors complicate this calculation, creating uncertainties and inaccuracies. These factors include the following: initial amplification cycles may not be exponential; PCR amplification eventually plateaus after an uncertain number of cycles; and low initial concentrations of target nucleic acid molecules may not amplify to detectable levels. However, the most significant limitation of PCR is that PCR amplification efficiency in a sample of interest may be different from that of reference samples. Since PCR is an exponential process, only twofold differences in amplification can be observed, greatly impacting the validity and precision of the results. Instead of performing one reaction per well, dPCR involves partitioning the PCR solution into tens of thousands of nano-liter sized droplets, where a separate PCR reaction takes place in each one. A PCR solution is made similarly to a TaqMan assay, which consists of template DNA (or RNA), fluorescence-quencher probes, primers, and a PCR master mix, which contains DNA polymerase, dNTPs, MgCl2, and reaction buffers at optimal concentrations. Several different methods can be used to partition samples, including microwell plates, capillaries, oil emulsion, and arrays of miniaturized chambers with nucleic acid binding surfaces. The PCR solution is divided into smaller reactions and are then made to run PCR individually. After multiple PCR amplification cycles, the samples are checked for fluorescence with a binary readout of “0” or “1”. The fraction of fluorescing droplets is recorded. The partitioning of the sample allows one to estimate the number of different molecules by assuming that the molecule population follows the Poisson distribution, thus accounting for the possibility of multiple target molecules inhabiting a single droplet. Using Poisson's law of small numbers, the distribution of target molecule within the sample can be accurately approximated allowing for a quantification of the target strand in the PCR product. This model simply predicts that as the number of samples containing at least one target molecule increases, the probability of the samples containing more than one target molecule increases. In conventional PCR, the number of PCR amplification cycles is proportional to the starting copy number. Different from many peoples's belief that dPCR provides absolute quantification, digital PCR uses statistical power to provide relative quantification. For example, if Sample A, when assayed in 1 million partitions, gives one positive reaction, it does not mean that the Sample A has one starting molecule. The benefits of dPCR include increased precision through massive sample partitioning, which ensures reliable measurements in the desired DNA sequence due to reproducibility. With basic PCR, error rates are larger when detecting small fold-change differences, however, with dPCR the error rate decreases because smaller fold-change differences can be detected in DNA sequence. The technique itself reduces the use of a larger volume of reagent needed, which inevitably will lower experiment cost. Also, dPCR is highly quantitative as it does not rely on relative fluorescence of the solution to determine the amount of amplified target DNA. dPCR measures the actual number of molecules (target DNA) as each molecule is in one droplet, thus making it a discrete “digital” measurement. It provides absolute quantification because dPCR measures the positive fraction of samples, which is the number of droplets that are fluorescing due to proper amplification. This positive fraction accurately indicates the initial amount of template nucleic acid. Similarly, qPCR utilizes fluorescence; however, it measures the intensity of fluorescence at specific times (generally after every amplification cycle) to determine the relative amount of target molecule (DNA), but cannot specify the exact amount without constructing a standard curve using different amounts of a defined standard. It gives the threshold per cycle (CT) and the difference in CT is used to calculate the amount of initial nucleic acid. As such, qPCR is an analog measurement, which may not be as precise due to the extrapolation required to attain a measurement. dPCR measures the amount of DNA after amplification is complete and then determines the fraction of replicates. This is representative of an endpoint measurement as it requires the observation of the data after the experiment is completed. In contrast, qPCR records the relative fluorescence of the DNA at specific points during the amplification process, which requires stops in the experimental process. This “real-time” aspect of qPCR may theoretically affect results due to the stopping of the experiment. In practice, however, most qPCR thermal cyclers read each sample's fluorescence very quickly at the end of the annealing/extension step before proceeding to the next melting step, meaning this hypothetical concern is not actually relevant or applicable for the vast majority of researchers. qPCR is unable to distinguish differences in gene expression or copy number variations that are smaller than twofold. It is difficult to identify alleles with frequencies of less than 1% because highly abundant, common alleles would be matched with similar sequences. On the other hand, dPCR has been shown to detect a differences of less than 30% in gene expression, distinguish between copy number variations of that differ by only 1 copy, and identify alleles that occur at frequencies less than 0.1%. Digital PCR has many applications in basic research, clinical diagnostics and environmental testing. Its uses include pathogen detection and digestive health analysis; liquid biopsy for cancer monitoring, organ transplant rejection monitoring and non-invasive prenatal testing for serious genetic abnormalities; copy number variation analysis, rare sequence detection, gene expression profiling and single-cell analysis; the detection of DNA contaminants in bioprocessing, the validation of gene edits and detection of specific methylation changes in DNA as biomarkers of cancer. dPCR is also frequently used as an orthogonal method to confirm rare mutations detected through next-generation sequencing (NGS) and to validate NGS libraries. dPCR enables the absolute and reproducible quantification of target nucleic acids at single-molecule resolution. Unlike analogue quantitative PCR (qPCR), however, absolute quantification with dPCR does not require a standard curve). dPCR also has a greater tolerance for inhibitor substances and PCR assays that amplify inefficiently as compared to qPCR.