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For reverse transcription polymerase chain reaction (RT-PCR), see reverse transcription polymerase chain reaction.
SYBR Green fluorescence chart produced in real-time PCR.

A real-time polymerase chain reaction is a laboratory technique of molecular biology based on the polymerase chain reaction (PCR). It monitors the amplification of a targeted DNA molecule during the PCR, i.e. in real-time, and not at its end, as in conventional PCR. Real-time PCR can be used quantitatively (Quantitative real-time PCR), semi-quantitatively, i.e. above/below a certain amount of DNA molecules (Semi quantitative real-time PCR).

Two common methods for the detection of PCR products in real-time PCR are: (1) non-specific fluorescent dyes that intercalate with any double-stranded DNA, and (2) sequence-specific DNA probes consisting of oligonucleotides that are labelled with a fluorescent reporter which permits detection only after hybridization of the probe with its complementary sequence.

The Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines propose that the abbreviation qPCR be used for quantitative real-time PCR and that RT-qPCR be used for reverse transcription–qPCR.[1] The acronym "RT-PCR" commonly denotes reverse transcription polymerase chain reaction and not real-time PCR, but not all authors adhere to this convention.[2]

Background[edit]

Real time PCR uses fluorophores in order to detect levels of gene expression.

Real-time PCR is one step of a 5-step process, with typical setup of controls.

Real-time PCR setup[edit]

Because the performance of wet lab work is rarely perfect, measures should be taken to account for the sub-optimal performance. It is done by setup of various types of controls (described later in details). Thus, a procedure involving real-time PCR has a typical setup consisting of the following 3 elements:

  1. Test sample - the material containing the target nucleic acids
  2. Controls, standards and/or calibrators of various types
  3. Data analysis and rules instructing how to process and interpret the data obtained from the test sample and controls, standards and/or calibrators.

Workflow overview[edit]

General workflow scheme of a real-time PCR involving procedure. The first two steps involve mostly wet-lab work, while the last three steps are involve data analysis only.

The 5-steps process of real-time PCR involving procedure can be described by linear workflow. The first two steps in a real-time PCR involving procedure consist of mostly wet-lab work and minor data analysis, while the last three consist of data analysis only (melt curve analysis and garbage disposal were not included in the workflow).

Pre-PCR steps[edit]

Pre-PCR steps consider all actions required before the PCR itself[1]. This often includes, but not limited to, sample collection, storage and transport, nucleic acids extraction and purification, evaluation of nucleic acids quality and quantity by non-PCR methods, nucleic acids storage and various treatments like Reverse Transcription and bisulfite conversion when necessary.

The versatility in pre-PCR steps is as large as the number of different types of sample sources. For instance, the sample collection technique is customized to the sample type and conditions. E.g., while blood is typically collected with syringe and special tubes to avoid coagulation and PCR inhibition[3], biopsies are collected by other methods, e.g. by Fine_needle_aspiration and some body liquids are collected by swabs.

Once the sample was collected, it should be stored and transported to the lab in conditions that will reduce degradation of nucleic acids[3]. This is especially important when working with RNA.

Often, the next step in the process is the extraction and sometimes the purification of nucleic acids. This step removes various PCR inhibitors that otherwise might kill the reaction or at least invalidate its result[4]. It should be noted that there are some procedures working directly on samples without nucleic acids extraction or purification[5], but this is not the case in most real-time PCR involving procedures. The purified nucleic acids may be used directly in real-time PCR or stored for later use. When working with RNA Reverse Transcription is required to convert the RNA to DNA, to enable efficient replication of the target sequence.

Basic principles of real-time PCR[edit]

Main article: Polymerase chain reaction

Real-time PCR is carried out in a thermal cycler with the capacity to illuminate each sample with a beam of light of at least one specified wavelength and detect the fluorescence emitted by the excited fluorophore. The thermal cycler is also able to rapidly heat and chill samples, thereby taking advantage of the physicochemical properties of the nucleic acids and DNA polymerase.

The PCR process generally consists of a series of temperature changes that are repeated 25 – 50 times. These cycles normally consist of three stages: the first, at around 95 °C, allows the separation of the nucleic acid’s double chain; the second, at a temperature of around 50-60 °C, allows the binding of the primers with the DNA template;[6] the third, at between 68 - 72 °C, facilitates the polymerization carried out by the DNA polymerase. Due to the small size of the fragments the last step is usually omitted in this type of PCR as the enzyme is able to increase their number during the change between the alignment stage and the denaturing stage. In addition, in four steps PCR the fluorescence is measured during short temperature phase lasting only a few seconds in each cycle, with a temperature of, for example, 80 °C, in order to reduce the signal caused by the presence of primer dimers when a non-specific dye is used.[7] The temperatures and the timings used for each cycle depend on a wide variety of parameters, such as: the enzyme used to synthesize the DNA, the concentration of divalent ions and deoxyribonucleotides (dNTPs) in the reaction and the bonding temperature of the primers.[8]

Chemical classification[edit]

Real-time PCR technique can be classified by the chemistry used to detect the PCR product, specific or non-specific fluorochromes.

Non-specific detection: Real-time PCR with double-stranded DNA-binding dyes as reporters[edit]

A DNA-binding dye binds to all double-stranded (ds) DNA in PCR, causing fluorescence of the dye. An increase in DNA product during PCR therefore leads to an increase in fluorescence intensity measured at each cycle. However, dsDNA dyes such as SYBR Green will bind to all dsDNA PCR products, including nonspecific PCR products (such as Primer dimer). This can potentially interfere with, or prevent, accurate monitoring of the intended target sequence.

In real-time PCR with dsDNA dyes the reaction is prepared as usual, with the addition of fluorescent dsDNA dye. Then the reaction is run in a real-time PCR instrument, and after each cycle, the intensity of fluorescence is measured with a detector; the dye only fluoresces when bound to the dsDNA (i.e., the PCR product). This method has the advantage of only needing a pair of primers to carry out the amplification, which keeps costs down; however, only one target sequence can be monitored in a tube.

Specific detection: fluorescent reporter probe method[edit]

(1) In intact probes, reporter fluorescence is quenched. (2) Probes and the complementary DNA strand are hybridized and reporter fluorescence is still quenched. (3) During PCR, the probe is degraded by the Taq polymerase and the fluorescent reporter released.

Fluorescent reporter probes detect only the DNA containing the sequence complementary to the probe; therefore, use of the reporter probe significantly increases specificity, and enables performing the technique even in the presence of other dsDNA. Using different-coloured labels, fluorescent probes can be used in multiplex assays for monitoring several target sequences in the same tube. The specificity of fluorescent reporter probes also prevents interference of measurements caused by primer dimers, which are undesirable potential by-products in PCR. However, fluorescent reporter probes do not prevent the inhibitory effect of the primer dimers, which may depress accumulation of the desired products in the reaction.

The method relies on a DNA-based probe with a fluorescent reporter at one end and a quencher of fluorescence at the opposite end of the probe. The close proximity of the reporter to the quencher prevents detection of its fluorescence; breakdown of the probe by the 5' to 3' exonuclease activity of the Taq polymerase breaks the reporter-quencher proximity and thus allows unquenched emission of fluorescence, which can be detected after excitation with a laser. An increase in the product targeted by the reporter probe at each PCR cycle therefore causes a proportional increase in fluorescence due to the breakdown of the probe and release of the reporter.

  1. The PCR is prepared as usual (see PCR), and the reporter probe is added.
  2. As the reaction commences, during the annealing stage of the PCR both probe and primers anneal to the DNA target.
  3. Polymerisation of a new DNA strand is initiated from the primers, and once the polymerase reaches the probe, its 5'-3'-exonuclease degrades the probe, physically separating the fluorescent reporter from the quencher, resulting in an increase in fluorescence.
  4. Fluorescence is detected and measured in a real-time PCR machine, and its geometric increase corresponding to exponential increase of the product is used to determine the quantification cycle (Cq) in each reaction.

Data analysis[edit]

Usually, data analysis of real-time PCR consists of three steps. The versatility in these steps is large. To reduce bias of results it is recommended to set the data analysis before collecting the data[9].

In specific detection chemistry, the signal of a responsive amplification curve is due to amplification of the target sequence. In non-responsive curve the signal is not due to amplification of the target sequence. Ambiguous curves cann't be classified confidently to one of the two options (responsive or non-responsive).

The first analysis of the data focuses on curve classification[10], examining whether the reaction is responsive or not, i.e., in sequence specific detection chemistries, if the signal is due to PCR amplification of the target sequence, or if the responsiveness of the curves can not be determined, and then the curve is referred to as ‘ambiguous’. In non-specific detection chemistry, the responsiveness of the curve considers any type of PCR product. Note the difference between ‘positive’ and ‘responsive’, ‘negative’ and ‘non-responsive’ and ‘equivocal’ and ‘ambiguous’. While the first in each pair is a result of a test, taking into account several more parameters like controls, standards, calibrators and biological or clinical rules and criteria (see below), the ‘responsiveness’ is a characteristics of a PCR amplification curve[11]. A curve, which is part of a test, can be responsive, and the result of the test can be negative. For instance, a responsive positive control and non-responsive target sequence may sum up to a negative result of a test.

Various factors may affect curves classification, most of them are chemical-technical factors that affect the shape of the curve, and thus, the result of classification[10]. Many of these factors are local to various degree. For instance, weak contamination combined with PCR inhibition is lab specific and sample specific correspondingly; unspecific amplification is assay specific; Crosstalk among detection channels are device specific[12]; reagents, e.g. probe, degradation are run or batch specific; PCR inhibition are sample specific[13] and biases and errors in background subtraction are reaction specific[14],[15].

But not only chemical-technical factors affect curves classification, the knowledge and experience of the interpreting person are specific. Since so many local factors affect curves classification, so far none of the various methods for automatic classification of amplification curves became a common practice[16][17]. The lack of well accepted computational method and the complexity of curves classification led many labs to develop their own classification procedures. Most commonly, curves are classified by visual inspection by the lab worker, or by manual setting of the threshold. Visual and manual analysis, not like fully automated analysis, are not mathematically well defined, and therefore are dependent on the person who analyses the data and his working conditions. This might lead to reduced reproducibility of results, contrary to the required by regulatory guidelines[18][19]. Because positive controls, which are mandatory elements in clinical real-time PCR, are often designed to be as close as possible to the limit of detection of the test they control, when small amounts of nucleic acids are detected or quantified[20], they are prone to ambiguity in curves classification. When higher amounts of nucleic acids are in question, usually the positive control will be designed to be somewhat further from the LOD to avoid this problem (see Positive control below).

Data validation[edit]

Once curves were classified the data can be used to validate the results. Here again, the technical variability results in large variability in validation methods, but three validation elements are common in many of real-time PCR involving procedures.

  1. Positive control: are analyzed to verify that the method is capable of adequately recovering and amplifying the target. The concentration of the sequence of interest in these positive controls should be 10 to 100 times higher than the defined detection limit of the PCR. It is recommended to include a positive control in each group of samples that are processed and amplified at the same time under the same conditions, using the same PCR master mix, and in the same thermocycler[21]. The origin of the nucleic acids in the positive control may vary according to the application.
  2. Negative control: are used to verify that no contaminating nucleic acid has been introduced into the master mix. These controls are prepared when template is added to the master mix. They are prepared as separate samples to which aliquots of molecular-grade water or buffer are added to the master mix in place of target nucleic acid or sample. A negative result with this control indicates that the master mix and final processing reagents are not contaminated[21].
  3. Inhibition control, also referred to as Internal Amplification control: is a non-target DNA or RNA in the test sample which is co-amplified with the target sequence. Following the amplification, the Cq of the control is compared to the Cq of the same amount of control quantified without the test sample. A difference between the Cqs indicates inhibition. This concept was adopted for detection of PCR inhibition in DNA quantification by the European Standardization Committee (CEN) and the International Organization for Standardization (ISO)[22]. In some settings, inhibition controls may serve as positive control.

Rules implementation[edit]

Once the data were validated tens of analysis methods may be applied[23][24] (see Rebrikov for a review [25]. For a comprehensive list see[26] ), and specific clinical criteria can be implemented. For instance, several methods consider Cq above a certain cut-off value as non-responsive curve, even if the curve is responsive[27].

Fusion temperature analysis[edit]

Distinct fusion curves for a number of PCR products (showing distinct colours). Amplification reactions can be seen for a specific product (pink, blue) and others with a negative result (green, orange). The fusion peak indicated with an arrow shows the peak caused by primer dimers, which is different from the expected amplification product.[28]

Real-time PCR permits the identification of specific, amplified DNA fragments using analysis of their melting temperature (also called Tm value, from melting temperature). The method used is usually PCR with double-stranded DNA-binding dyes as reporters and the dye used is usually SYBR Green. The DNA melting temperature is specific to the amplified fragment. The results of this technique are obtained by comparing the dissociation curves of the analysed DNA samples.[29]

Unlike conventional PCR, this method avoids the previous use of electrophoresis techniques to demonstrate the results of all the samples. This is because, despite being a kinetic technique, quantitative PCR is usually evaluated at a distinct end point. The technique therefore usually provides more rapid results and / or uses fewer reactants than electrophoresis. If subsequent electrophoresis is required it is only necessary to test those samples that real time PCR has shown to be doubtful and / or to ratify the results for samples that have tested positive for a specific determinant.

Modeling[edit]

Unlike end point PCR (conventional PCR) real time PCR allows monitoring of the desired product at any point in the amplification process by measuring fluorescence (in reality, measurement is made of its level over a given threshold). A commonly employed method of DNA quantification by real-time PCR relies on plotting fluorescence against the number of cycles on a logarithmic scale. A threshold for detection of DNA-based fluorescence is set 3-5 times of the standard deviation of the signal noise above background. The number of cycles at which the fluorescence exceeds the threshold is called the threshold cycle (Ct) or, according to the MIQE guidelines, quantification cycle (Cq).[30]

During the exponential amplification phase, the quantity of the target DNA template (amplicon) doubles every cycle. For example, a DNA sample whose Cq precedes that of another sample by 3 cycles contained 23 = 8 times more template. However, the efficiency of amplification is often variable among primers and templates. Therefore, the efficiency of a primer-template combination is assessed in a titration experiment with serial dilutions of DNA template to create a standard curve of the change in (Cq) with each dilution. The slope of the linear regression is then used to determine the efficiency of amplification, which is 100% if a dilution of 1:2 results in a (Cq) difference of 1. The cycle threshold method makes several assumptions of reaction mechanism and has a reliance on data from low signal-to-noise regions of the amplification profile that can introduce substantial variance during the data analysis.[31]

To quantify gene expression, the (Cq) for an RNA or DNA from the gene of interest is subtracted from the (Cq) of RNA/DNA from a housekeeping gene in the same sample to normalize for variation in the amount and quality of RNA between different samples. This normalization procedure is commonly called the ΔCt-method[32] and permits comparison of expression of a gene of interest among different samples. However, for such comparison, expression of the normalizing reference gene needs to be very similar across all the samples. Choosing a reference gene fulfilling this criterion is therefore of high importance, and often challenging, because only very few genes show equal levels of expression across a range of different conditions or tissues.[33][34] Although cycle threshold analysis is integrated with many commercial software systems, there are more accurate and reliable methods of analysing amplification profile data that should be considered in cases where reproducibility is a concern.[31]

Mechanism-based qPCR quantification methods have also been suggested, and have the advantage that they do not require a standard curve for quantification. Methods such as MAK2[35] have been shown to have equal or better quantitative performance to standard curve methods. These mechanism-based methods use knowledge about the polymerase amplification process to generate estimates of the original sample concentration. An extension of this approach includes an accurate model of the entire PCR reaction profile, which allows for the use of high signal-to-noise data and the ability to validate data quality prior to analysis.[31]

According to research of Ruijter et al.[36] MAK2 assumes constant amplification efficiency during the PCR reaction. However, theoretical analysis of polymerase chain reaction, from which MAK2 was derived, has revealed that amplification efficiency is not constant throughout PCR. While MAK2 quantification provides reliable estimates of target DNA concentration in a sample under normal qPCR conditions, MAK2 does not reliably quantify target concentration for qPCR assays with competimeters.

Applications[edit]

There are numerous applications for quantitative polymerase chain reaction in the laboratory. It is commonly used for both diagnostic and basic research. Uses of the technique in industry include the quantification of microbial load in foods or on vegetable matter, the detection of GMOs (Genetically modified organisms) and the quantification and genotyping of human viral pathogens.

Quantification of gene expression[edit]

Quantifying gene expression by traditional DNA detection methods is unreliable. Detection of mRNA on a Northern blot or PCR products on a gel or Southern blot does not allow precise quantification.[37] For example, over the 20-40 cycles of a typical PCR, the amount of DNA product reaches a plateau that is not directly correlated with the amount of target DNA in the initial PCR.[citation needed]

Real-time PCR can be used to quantify nucleic acids by two common methods: relative quantification and absolute quantification.[38] Absolute quantification gives the exact number of target DNA molecules by comparison with DNA standards using a calibration curve. It is therefore essential that the PCR of the sample and the standard have the same amplification efficiency.[4] Relative quantification is based on internal reference genes to determine fold-differences in expression of the target gene. The quantification is expressed as the change in expression levels of mRNA interpreted as complementary DNA (cDNA, generated by reverse transcription of mRNA). Relative quantification is easier to carry out as it does not require a calibration curve as the amount of the studied gene is compared to the amount of a control reference gene.

As the units used to express the results of relative quantification are unimportant the results can be compared across a number of different RT-Q-PCR. The reason for using one or more housekeeping genes is to correct non-specific variation, such as the differences in the quantity and quality of RNA used, which can affect the efficiency of reverse transcription and therefore that of the whole PCR process. However, the most crucial aspect of the process is that the reference gene must be stable.[39]

The selection of these reference genes was traditionally carried out in molecular biology using qualitative or semi-quantitative studies such as the visual examination of RNA gels, Northern blot densitometry or semi-quantitative PCR (PCR mimics). Now, in the genome era, it is possible to carry out a more detailed estimate for many organisms using DNA microarrays.[40] However, research has shown that amplification of the majority of reference genes used in quantifying the expression of mRNA varies according to experimental conditions.[41][42][43] It is therefore necessary to carry out an initial statistically sound methodological study in order to select the most suitable reference gene.

A number of statistical algorithms have been developed that can detect which gene or genes are most suitable for use under given conditions. Those like geNORM or BestKeeper can compare pairs or geometric means for a matrix of different reference genes and tissues.[44][45]

Diagnostic uses[edit]

Diagnostic qualitative PCR is applied to rapidly detect nucleic acids that are diagnostic of, for example, infectious diseases, cancer and genetic abnormalities. The introduction of qualitative PCR assays to the clinical microbiology laboratory has significantly improved the diagnosis of infectious diseases,[46] and is deployed as a tool to detect newly emerging diseases, such as new strains of flu, in diagnostic tests.[47]

Microbiological uses[edit]

Quantitative PCR is also used by microbiologists working in the fields of food safety, food spoilage and fermentation and for the microbial risk assessment of water quality (drinking and recreational waters) and in public health protection.[48]

Uses in research[edit]

In research settings, quantitative PCR is mainly used to provide quantitative measurements of gene transcription. The technology may be used in determining how the genetic expression of a particular gene changes over time, such as in the response of tissue and cell cultures to an administration of a pharmacological agent, progression of cell differentiation, or in response to changes in environmental conditions. It is also used for the determination of zygosity of transgenic animals used in research.

Detection of phytopathogens[edit]

The agricultural industry is constantly striving to produce plant propagules or seedlings that are free of pathogens in order to prevent economic losses and safeguard health. Systems have been developed that allow detection of small amounts of the DNA of Phytophthora ramorum, an oomycete that kills Oaks and other species, mixed in with the DNA of the host plant. Discrimination between the DNA of the pathogen and the plant is based on the amplification of ITS sequences, spacers located in ribosomal RNA gene’s coding area, which are characteristic for each taxon.[49] Field-based versions of this technique have also been developed for identifying the same pathogen.[50]

Detection of genetically modified organisms[edit]

qPCR using reverse transcription (RT-qPCR) can be used to detect GMOs given its sensitivity and dynamic range in detecting DNA. Alternatives such as DNA or protein analysis are usually less sensitive. Specific primers are used that amplify not the transgene but the promoter, terminator or even intermediate sequences used during the process of engineering the vector. As the process of creating a transgenic plant normally leads to the insertion of more than one copy of the transgene its quantity is also commonly assessed. This is often carried out by relative quantification using a control gene from the treated species that is only present as a single copy.[51][52]

Clinical quantification and genotyping[edit]

Viruses can be present in humans due to direct infection or co-infections which makes diagnosis difficult using classical techniques and can result in an incorrect prognosis and treatment. The use of qPCR allows both the quantification and genotyping (characterization of the strain, carried out using melting curves) of a virus such as the Hepatitis B virus.[53] The degree of infection, quantified as the copies of the viral genome per unit of the patient’s tissue, is relevant in many cases; for example, the probability that the type 1 herpes simplex virus reactivates is related to the number of infected neurons in the ganglia.[54] This quantification is carried out either with reverse transcription or without it, as occurs if the virus becomes integrated in the human genome at any point in its cycle, such as happens in the case of HPV (human papillomavirus), where some of its variants are associated with the appearance of cervical cancer.[55]

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Bibliography[edit]

External links[edit]


Category:Molecular biology Category:Laboratory techniques Category:Polymerase chain reaction Category:Real-time technology

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