PCR consists of three main thermal cycling steps with several essential reaction components as described in the previous sections. Depending on the application, PCR setup may be modified to achieve specific experimental outcomes such as increased yield, improved specificity, or reduced assay time, as explained in the following section (Table 1).
Table 1. Common PCR methods and their core benefits.
Hot-start PCR is commonly used to enhance specificity in PCR amplification. Methods of hot-start PCR employ an enzyme modifier such as an antibody, affibody, aptamer, or chemical modification to inhibit DNA polymerase activity at room temperature. This modification prevents nonspecific amplification due to primers binding to template sequences with low homology (mispriming), and primers binding to each other (primer-dimers), during reaction setup. Since the activity of the DNA polymerase is blocked at room temperature, the hot-start strategy enables the convenience of setting up multiple reactions at ambient temperature (as in high-throughput experiments) without significantly compromising specificity and amplification (learn more in DNA polymerase characteristics).
After the reactions are set up, the DNA polymerase is activated in an initial heating step or “hot start” in which the enzyme modifier is released at a high temperature (usually above 90°C) (Figure 1). The activation time and temperature vary depending on the DNA polymerase and the nature of the hot-start modifier. For some DNA polymerases, activation and initial denaturation steps may be combined into a single step.
Figure 1. DNA polymerase with antibody-based hot-start technology.
Another approach to promoting specificity is to modify the PCR cycling parameters. In touchdown PCR, the annealing temperature of the first few cycles is set to be a few degrees higher than the highest melting temperature (Tm) of the primers [1,2]. Higher temperatures help destabilize the formation of primer-dimers and nonspecific primer-template complexes, thus minimizing undesirable amplification. As such, higher annealing temperatures reduce nonspecific PCR products and promote specific amplification at the start of PCR (learn more about PCR annealing step).
While preventing primer-dimers and nonspecific primer binding, the higher annealing temperatures may result in lower PCR yield due to increased dissociation of primers from their intended target. To overcome this challenge, the annealing temperature is often decreased 1°C at every cycle of the initial few cycles to produce a sufficient yield of the desired amplicon. Once the annealing temperature reaches, or “touches down”, at the optimal temperature (usually 3–5°C lower than the lowest primer Tm), it is maintained throughout the remaining cycles for primer annealing. In this manner, desired PCR products are selectively increased with little or no amplification of nonspecific targets over the course of PCR (Figure 2).
Figure 2. Touchdown PCR. The method promotes specificity (yellow curve) by starting with a higher-than-optimal annealing temperature, which is then gradually lowered (black line) as cycling continues until the optimal annealing temperature is reached. The yield of the intended amplicon (green curve) accumulates considerably with the optimized annealing temperature.
Nested PCR is a variation of standard PCR that enhances the specificity and yield of the desired amplicons . In this method, two pairs of PCR primers are designed: one set (outer primers) flanks a region of DNA containing the amplicon of interest, while a second set (nested primers) corresponds to the precise region of DNA to be amplified. The outer primers are used in a first round of PCR to amplify the target with extended flanking regions. The product of this first round then serves as a template in a second round of PCR with the nested primers (Figure 3).
Figure 3. Nested PCR.
In the event that nonspecific products are amplified because of mispriming by the first set of primers (outer primers), it is very unlikely that the same nonspecific region would be recognized and sequentially amplified by the second primer set, so specificity is still promoted by the second set of primers if they recognize the intended target. Another benefit of two PCR rounds is that this approach helps to obtain a sufficient yield of the desired target from a limited amount of input DNA.
In fast PCR, durations of PCR steps are shortened to complete amplification faster without impacting yield or efficiency. Fast cycling conditions are especially suitable with highly processive DNA polymerases, as these enzymes are capable of incorporating a greater number of nucleotides during each binding event (learn more about processivity). Highly processive DNA polymerases can maintain high amplification efficiency with PCR extension times that are 1/2 to 1/3 the duration of the extension times needed for Taq polymerase, which has low processivity (Figure 4). PCR times can be further shortened by combining primer annealing and extension into a single step, if their temperatures are within a few degrees of each other. This procedure is also known as a two-step PCR protocol.
Figure 4. Comparison of DNA polymerases with low vs. high processivity in the amplification of a 3.8 kb fragment from human gDNA. A two-step PCR protocol (with annealing and extension steps combined) was performed.
When a DNA polymerase with low processivity, such as Taq polymerase, is used, fast cycling conditions may be possible for short targets of <500 bp. Amplicons of this size generally do not require extended polymerization times, so it may be possible to shorten the extension step in the PCR protocol. To find the shortest elongation time that does not compromise product yield, PCR may be performed with a series of extension time decrements (in seconds). Each target and primer set will likely produce variable results and thus will need optimization for fast PCR under specified conditions.
Another modification of fast PCR is to shorten the denaturation time while increasing the temperature to 98°C to compensate. The caveat to this strategy is that enzymes that are not highly thermostable will easily become denatured at such high temperatures (learn more about DNA polymerase thermostability).
Table 2. Reaction parameters for achieving fast PCR using DNA polymerases with low processivity.
|Parameter||Optimization for fast PCR|
|Amplicon length||<500 bp|
Direct PCR refers to amplification of target DNA directly from samples without nucleic acid isolation. In direct PCR, samples such as cells or tissue, are lysed in a specially formulated buffer to release DNA during high-temperature denaturation steps. Therefore, this method simplifies workflows, saves hands-on time, and prevents DNA loss caused by purification steps (Figure 5).
Figure 5. Conventional vs. direct PCR.
DNA polymerases with high processivity are often recommended for direct PCR. Cellular debris, proteins, lipids, and polysaccharides are released into the lysates along with the DNA, and they can inhibit PCR. DNA polymerases with high processivity tolerate such inhibitors and make direct PCR possible. Enzymes with high processivity often display higher sensitivity and therefore can be used to successfully amplify low amounts of DNA from unpurified samples.
DNA templates containing high GC content (>65%) can be difficult to amplify because of the stronger hydrogen bonds between G and C bases. GC-rich sequences can also be involved in secondary structures. Thus, GC-rich sequences can cause DNA polymerases to “stutter” along templates and interrupt DNA synthesis.
To amplify GC-rich targets, the double-stranded template must be separated for the primers to bind and DNA polymerase to read through the sequence. To overcome strong GC interactions, the most common approach relies on PCR additives or co-solvents such as DMSO to help DNA denature (Figure 6A). These reagents often lower the primer Tm, so the annealing temperature should be adjusted accordingly.
Highly processive DNA polymerases are beneficial for GC-rich PCR, because of their strong binding to the templates during primer extension (Figure 6B). Hyperthermostable DNA polymerases are also advantageous for GC-rich PCR, since a higher denaturation temperature (e.g., 98°C instead of 95°C) may facilitate strand separation and PCR amplification (learn more about PCR cycling).
Figure 6. Amplification of regions of human gDNA with differences in GC content. (A) A ~0.8 kb target with 76% GC was amplified using a DNA polymerase with low processivity. Increasing amounts of DMSO as an additive promote specificity. (B) Seven fragments of various GC content were amplified using a highly processive DNA polymerase. A GC enhancer was used only for the fragments with 70% and 76% GC.
Multiplex PCR allows concurrent amplification of different targets in a single PCR tube. Multiplexing not only saves time, reagents, and samples but also makes simultaneous comparison of multiple amplicons possible (Figure 7).
Figure 7. Singleplex PCR vs. multiplex PCR. In singleplex PCR, each reaction contains one primer pair to amplify one target. In multiplex PCR, multiple primer pairs are used to amplify multiple targets in one reaction.
When there are multiple primer pairs in a single tube as in multiplex PCR, nonspecific amplification and reduced efficiency are concerns because the reaction cannot be optimized for a single primer pair and target but rather for all of the primers and targets. Therefore, primer design is critical to minimize mispriming that would lead to nonspecific amplification. Primer sequences should be as unique to their target as possible, and the Tms of all primers should be within 5°C of each other. Prior to multiplexing, each primer set should be validated in a singleplex reaction for specificity and efficiency. Furthermore, the amplicons should be of distinct sizes that can be resolved by gel electrophoresis for identification. In addition to primer design and amplicon size, a hot-start DNA polymerase and a PCR buffer specially formulated for multiplexing can aid PCR success and specificity (Figure 8).
Figure 8. Singleplex and multiplex PCR results displayed by gel electrophoresis. Invitrogen™ Platinum™ Multiplex PCR Master Mix was used in this experiment.
Although multiplex PCR is routinely performed as endpoint reactions, the approach is more popular with real-time PCR because of capabilities in multi-labeling and detection of target amplification. Multiplex real-time PCR is also frequently employed in detection of genetic markers for human identification.
Long PCR generally refers to amplification of DNA targets longer than 5 kb. Long PCR traditionally has been performed with a blend of Taq DNA polymerase (for fast elongation) and a high-fidelity enzyme (for accuracy).
With the inventions of highly processive, high-fidelity DNA polymerases, long PCR can now be performed with greatly improved accuracy in a shorter time. High processivity is achieved by designing a strong DNA-binding domain, which enables amplification of long fragments (e.g., >20 kb from gDNA) in a few hours (Figure 9, Table 3). In addition, extremely high fidelity (e.g., >100x fidelity relative to Taq polymerase) helps to ensure a low error rate in replication of long fragments (Table 3). (Learn more about DNA polymerase characteristics.)
Figure 9. Amplification of long fragments using a highly processive DNA polymerase. Specific amplification of 15 kb and 30 kb fragments was obtained from human gDNA samples.
Table 3. Advantages of highly processive, high-fidelity engineered DNA polymerase in long PCR and cloning.
High processivity of a DNA polymerase shortens the reaction times of long PCR significantly (by half in this example), while high fidelity reduces the effort required to screen for colonies with error-free inserts.
|Blend of Taq and proofreading DNA polymerases||Highly processive, high-fidelity engineered DNA polymerase|
|Extension rate||60 sec/kb||30 sec/kb|
(20 kb target, 30 cycles)
|~10.5 hr||~5.2 hr|
(relative to Taq DNA polymerase)
|Cloning error rate
(20 kb, 30-cycle PCR)
|Average of 2.5 errors per clone||1 in 4 clones may contain an error|
When amplifying targets of >10 kb, the PCR protocol may be optimized in the following five key areas:
- Ensure that DNA samples are of good quality and purity.
- Use higher amounts of a DNA polymerase if it is of low thermostability, to compensate for loss of activity over extended cycling times.
- Decrease the temperatures of the annealing and extension steps to help with primer binding.
- Increase the durations of the PCR steps to promote complete separation of the template DNA and binding of the primers.
- Extend PCR elongation times accordingly to ensure full-length replication of the target regions.
Inverse PCR was originally designed to determine sequences of adjacent unknown regions. Inverse PCR is helpful for investigating the promoter sequence of a gene; oncogenic chromosomal rearrangements such as gene fusion, translocation, and transposition; and viral gene integration. The method is known as inverse PCR because the primers are designed to extend away from each other rather than toward each other as in regular PCR [4,5]. Today, inverse PCR is routinely employed in site-directed mutagenesis to replicate a target plasmid while introducing desired mutations.
In a conventional workflow for studying unknown sequences of genomic DNA, restriction digestion and ligation precede inverse PCR, which is then followed by sequencing of the PCR amplicon. For the gDNA digestion, a restriction enzyme is chosen to generate fragments of suitable lengths that can self-ligate. Also, the selected restriction enzyme should not cleave the known sequence, so ligation occurs between the flanking unknown sequences. Ligation is optimized by using low concentrations of digested DNA fragments to favor self-ligation over multi-fragment ligation (i.e., concatemer formation).
After self-ligation, inverse PCR is performed by priming from the known region of DNA. The resulting amplicons contain a portion of the known DNA sequence at each end. These amplicons can then be sequenced from the end(s) to examine the regions adjacent to the previously known sequence (Figure 10).
Figure 10. Inverse PCR for amplification and characterization of adjacent unknown sequences.
As the extent (yield) of amplification of a sequence depends on the amount of template input, PCR is commonly employed to quantify the amount of DNA present in a sample, of which the most common application is the quantitation of gene expression. Endpoint PCR is one possible method, but it has major disadvantages since the yield is determined by gel electrophoresis, where detection sensitivity is limited. More importantly, quantitation would be at the end of PCR when amplification has reached the plateau phase (Figure 11), where intensities of DNA gel staining do not correlate linearly with amounts of DNA input. Nevertheless, for semi-quantitative analysis of gene expression by endpoint PCR before reaching the plateau phase, serially diluted DNA samples can be used as inputs, or amplicons at specific PCR cycles can be collected for estimation of expressed genes via gel intensity [6,7].
Figure 11. Amplification curve or reaction dynamic of PCR. In endpoint PCR, amplicons are detected when amplification has reached the plateau phase after PCR. In real-time PCR, quantitation of amplicons occurs during the exponential phase.
The limitations of endpoint PCR–based quantitation were overcome when Higuchi et al. reported real-time monitoring of PCR amplification using fluorescent signals, in 1993 . The technique became the basis of quantitative PCR (qPCR) as we know it today. In 1997, the first qPCR machines were introduced to the market [9,10], making accessible the power of PCR to accurately quantify gene expression and copy numbers. qPCR relies upon real-time monitoring of fluorescence intensity associated with target amplification during the exponential phase (Figure 11), avoiding the pitfalls of endpoint PCR quantitation. While qPCR is quantitative in measuring relative and absolute gene expression, its quantitation is still limited by detection capabilities.
True absolute quantitation of DNA samples became possible with digital PCR (also called limiting dilution PCR), a method developed in parallel with real-time PCR in the 1990s [11-13]. In digital PCR, a highly diluted DNA sample is partitioned in a multi-compartment chip such that each compartment contains no more than one copy of the target of interest. Amplification within each compartment is determined to obtain positive or negative results (from 1 or 0 template copies, respectively; i.e., "digital" results). The copy number of the sample is determined from the fraction of negative reactions using a statistical model (Poisson distribution) without a need for known samples (standards) for quantitation (Figure 12). In addition to gene expression and copy number determination, digital PCR is suitable for applications such as discrimination of low-frequency alleles, viral titering, and absolute quantitation of next-generation sequencing libraries.
Figure 12. Common workflow of digital PCR for absolute quantification.
In summary, modified PCR protocols and DNA polymerases are routinely employed to improve amplification results. Although the fundamental concept of PCR has remained unchanged, novel methods in PCR have continued to advance and streamline molecular biology research.
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For Research Use Only. Not for use in diagnostic procedures.