How PCR Systems Work & How We Save You Time & Money

Real-time PCR is made possible through the use of a specialized thermal cycler equipped with fluorescence detection modules which monitors the fluorescence as amplification occurs. This method is referred to as real-time PCR, or qPCR (quantitative PCR).

Unlike conventional PCR, the most important distinction of this method is that the PCR product (the amplified DNA) can be detected in real time (during amplification), rather than during the post-PCR step which requires using agarose gel to detect and visualize. Another critical difference is that qPCR can be quantitative and specific. For example, starting quantities of a sequence of interest can be determined by comparison of samples to a standard curve of known quantities of DNA

To increase its specificity, TaqMan probes have been introduced to qPCR. In principle, these probes rely on the 5’–3′ exonuclease activity of Taq polymerase to cleave a dual-labeled probe during hybridization to the complementary target sequence, detecting and quantifying specific PCR products as the reaction proceeds using fluorophore-based detection.

Quantitative PCR assays have been designed to support a number of applications across a wide range of fields, including gene expression analysis, genotyping, and microRNA analysis (miRNA).

When shopping for real-time PCR thermocyclers, considerations should include the footprint, multiplexing capabilities, compatibility with standard liquid handling workstations and multichannel pipettes, reagent and consumables cost, ease-of-use, software capabilities, and throughput potential.

From an economic standpoint, we have seen a decrease in qPCR machine cost as technology has rapidly improved over the years, making these apparatuses much more accessible to the lab community.

DNA polymerases used for basic PCR require a DNA template. There are, however, numerous instances in which amplification of RNA would be preferred. To study RNA, the sample must first be reverse transcribed into complimentary (c)DNA using a reverse transcriptase enzyme.

The resulting cDNA can then be used as a template for subsequent polymerase chain reaction amplification. The specific technique PCR technique performed using the cDNA template is referred to as Reverse Transcription PCR, or RT-PCR. This method is not to be confused with real-time PCR, also referred to as quantitative PCR (qPCR).

The first step of RT-PCR includes the synthesis of a DNA/RNA hybrid. Reverse transcriptase also has an RNase (ribonuclease H) function, which degrades the RNA portion of the hybrid. The single stranded DNA molecule is then completed by the DNA-dependent DNA polymerase activity of the reverse transcriptase into cDNA (complementary DNA). It’s important to note that the efficiency of the first-strand reaction can affect the amplification process.

Afterwards, the standard PCR procedure is used to amplify the cDNA. The possibility to revert RNA into cDNA by RT-PCR has many advantages. However, RNA is single-stranded and very unstable, which makes it difficult to work with. It serves as a first step in qPCR, which quantifies RNA transcripts in a biological sample. RT-PCR is commonly used for gene expression profiling, inserting eukaryotic genes into prokaryotes, and disease diagnosis.

In comparison to qPCR, digital PCR (dPCR) devices provide precise, sensitive, and reproducible quantitative results concerning absolute counts of target nucleic acids. For that reason, dPCR has the potential to have a major impact on molecular analyses in both clinical and research settings and applications.

This includes biomarker analysis, viral detection, prognostic monitoring, and fetal screening, as well as phage-host interactions and intracellular profiling. dPCR can also be applied to assist with the library preparation needed for massively parallel next-generation sequencing methods.

Unlike conventional PCR, which provides end-point analysis and requires a secondary step post-PCR amplification, making it semi-quantitative, dPCR (similar to qPCR) provides absolute quantification. However, unlike qPCR, dPCR does not require references or standards.

Furthermore, it provides a unique sample partitioning step, where partitioning of the PCR reaction into thousands of individual reaction vessels prior to amplification occurs. This makes the technique highly sensitive and precise, building on the capabilities of qPCR.

These features make digital PCR well suited for applications that require the detection of small amounts of nucleic acid or finer resolution of target amounts among samples, which includes rare sequence detection, copy number variation (CNV) analysis, and gene expression analysis of the rare targets.

Another method for performing digital PCR, droplet digital PCR (ddPCR) utilizes Taq polymerase to amplify the targeted DNA in a complex sample. Additionally, it uses microfluidics to emulsify samples in oil, creating reproducible droplets to be processed and analyzed by fluorescence.

Before being analyzed in the ddPCR process, a sample is divided into 20,000 droplets using water-emulsion droplet technology, undergoes thermal cycling, and is then ran through a 96-well PCR plate. Following this process, each droplet is analyzed to determine the using Poisson statistics.

This analysis determines the amount of PCR-positive droplets in the original sample. These data are then further analyzed using Poisson statistics which determines the concentration of target DNA (absolute count) of the original sample.

As mentioned, this partitioning of the sample into thousands of reaction vessels is a major difference between digital PCR methods and and qPCR. However, ddPCR is seen as more advanced compared to earlier digital PCR methods due to its practicality and scalability.

ddPCR’s approaches are at the forefront of detection technology, and accordingly, it is also the most expensive purchase one can opt for.

Following the same principles as conventional PCR, Hot start PCR uses DNA polymerase to synthesize DNA from a single stranded template. However, the major difference here is that hot start includes additional heating and separation steps.

This includes blocking Taq polymerase activity at low temperatures with specific anti-Taq antibodies and other various mechanisms, such as chemical modifiation and aptamer technology.

By including these methods, hot start PCR reduces nonspecific priming/amplification and the creation of primer dimers. In addition, it improves PCR performance by increasing product yields, specificity, and sensitivity.

Unlike the conventional method, which amplifies a single target in a reaction well, mutliplex PCR allows for the simultaneous detection of multiple targets in a single reaction well. This is achieved using multiple primers that can be distinguished from one another.

By amplifying several segments at the same time, rather than individually, laboratories can greatly benefit from multiplex PCR, saving on space, time, and reagents. Furthermore, it requires a small amount of DNA as the starting template and can be performed on specimens with poor DNA quality.

Despite its benefits, multiplex PCR optimization is challenging. When using multiple primer pairs, primers from one pair can interact with primers from another. Because each primer pair can have different requirements, there isn’t an optimal melting temperature and delta G, a change in free energy.

Multiplex PCR is commonly used for variety of research applications. This includes gene panel expression, SNP genotyping, pathogen detection, template quantification, and more.

Similar to digital PCR, nested methods reduce the chances of nonspecific amplification of the target sequence. This is due to nested PCR’s use of multiple primer sets directed at the same target segment and two successive PCR reactions.

The first set will anneal to sequences ahead of the second, nested, primer set, and in doing so will amplify a large fragment of the gene that is then used as a template during the second PCR reaction. The second round targets a much smaller region of the amplicon—the segment of chromosomal DNA that has undergone amplification and contains replicated DNA—using the nested primer set.

While the primers were traditionally added one after the other, single-tube nested PCR reactions were developed to deal with the challenges of contamination and decrease in specificity.

Nested PCR, while extremely sensitive, poses the risk of contamination and is costly to run. Nonetheless, this technique has proven valuable to laboratories amplifying and detecting genes that are low in abundance.