Variants of PCR

Often only a small modification needs to be made to the 'standard' PCR protocol to achieve a desired goal:

One of the first adjustments made to PCR was the amplification of more than one target in a single tube. Multiplex-PCR can involve up to a dozen pairs of primers acting independently. This modification might be used simply to avoid having to prepare many individual reactions, or could allow the simultaneous analysis of multiple targets in a sample that has only a single copy of a genome. In testing for genetic disease mutations, six or more amplifications might be combined. In the standard protocol for DNA Fingerprinting, the 13 targets assayed are often amplified in groups of 3 or 4. Multiplex Ligation-dependent Probe Amplification (or MLPA) permits multiple targets to be amplified using only a single pair or primers, avoiding the resolution limitations of multiplex PCR

VNTR PCR involves few modifications to the basic PCR process, but instead targets areas of the genome that exhibit length variation. The analysis of the genotypes of the sample usually involves simple sizing of the amplification products by gel electrophoresis. Analysis of smaller VNTR segments known as Short Tandem Repeats (or STRs) is the basis for DNA Fingerprinting databases such as CODIS.

Asymmetric PCR is used to preferentially amplify one strand of the target DNA. It finds use in some types of sequencing and hybridization probing, where having only one of the two complementary strands of the product is advantageous. PCR is carried out as usual, but with a limiting amount of one of the primers. When it becomes depleted, continued replication leads to an arithmetic increase in extension of the other primer. A recent modification on this process, known as Linear-After-The-Exponential-PCR (or LATE-PCR), uses a limiting primer with a higher melting temperature Melting temperature (or Tm) than the excess primer to maintain reaction efficiency as the limiting primer concentration decreases mid-reaction. (Also see Overlap-extension PCR).

Some modifications are needed to perform long PCR. The original Klenow-based PCR process had trouble making a product larger than about 400 bp. However, early characterization of Taq polymerase showed that it could amplify targets up to several thousand bp long. Since then, modified protocols have allowed targets of over 50,000 bp to be amplified

Nested PCR, another early modification, can be used to increase the specificity of DNA amplification. Two sets of primers are used in two successive reactions. In the first, one pair of primers is used to generate DNA products, which may also contain products amplified from non-target areas. The products from the first PCR are then used to start a second, using one ('hemi-nesting') or two different primers whose binding sites are located (nested) within the first set. The specificity of all of the primers is combined, usually leading to a single product. Nested PCR is often more successful in specifically amplifying long DNA products than conventional PCR, but it requires more detailed knowledge of the sequence of the target.

Quantitative PCR (or Q-PCR) is used to measure the specific amount of target DNA (or RNA) in a sample. The normal PCR process is performed in a way that is largely qualitative - the amount of final product is only slightly proportional to the initial amount of target. By carefully running the amplification only within the phase of true exponential increase (avoiding the later 'plateau' phase), the amount of product is more proportional to the initial amount of target. Thermal cyclers have been developed which can monitor the amount of product during the amplification, allowing quantitation of samples containing a wide range of target copies. A method currently used is Quantitative Real-Time PCR. QRT-PCR methods use fluorescent dyes, such as Sybr Green, or fluorophore-containing DNA probes, such as TaqMan, to measure the amount of amplified product as the amplification progresses. It is often confusingly referred to as RT-PCR, the same acronym used for PCR combined with Reverse Transcriptase (see below), which itself might be used in conjunction with Q-PCR. More appropriate acronyms are QRT-PCR or RTQ-PCR.

Hot-start PCR is a technique that modifies the way that a PCR mixture is initially heated. During this step the polymerase is active, but the target has not yet been denatured and the primers may be able to bind to non-specific locations (or even to each other). The technique can be performed manually by heating the reaction components to the melting temperature (e.g. 95°C) before adding the polymerase. Alternatively, specialized systems have been developed that inhibit the polymerase's activity at ambient temperature, either by the binding of an antibody, or by the presence of covalently bound inhibitors that only dissociate after a high-temperature activation step. 'Hot-start/cold-finish PCR' is achieved with new hybrid polymerases that are inactive at ambient temperature and are only activated at elevated temperatures.

Another simple modification can also decrease non-specific amplification. In Touchdown PCR, the temperature used to anneal the primers is gradually decreased in later cycles. The annealing temperature in the early cycles is usually 3-5°C above the standard Tm of the primers used, while in the later cycles it is a similar amount below the Tm. The initial higher annealing temperature leads to greater specificity for primer binding, while the lower temperatures permit more efficient amplification to the end of the reaction.

Other common modifications to PCR allow it to amplify low copy targets. The original report on Taq polymerase showed how the use of up to 60 cycles could amplify targets diluted to just one copy per reaction tube. A later report showed how multiple genetic loci could be amplified and analyzed from a single sperm. Modified protocols have allowed the identification of just one copy of the HIV genome within the DNA of up to 70,000 host cells.

Assembly PCR (also known as Polymerase Cycling Assembly or PCA) is the artificial synthesis of long DNA structures by performing PCR on a pool of long oligonucleotides with short overlapping segments. The oligonucleotide building blocks alternate between sense and antisense directions, and the overlaps determine the order of oligonucleotides, thereby selectively producing the final long DNA product.

In Colony PCR, bacterial colonies are rapidly screened by PCR for correct DNA vector constructs. Colonies are sampled with a sterile toothpick and dabbed into a master mix. To free the DNA for amplification, PCR is either started with an extended time at 95°C (when standard polymerase is used), or with a shortened denaturation step at 100°C and special chimeric DNA polymerase. Colonies from the master mix that shows the desired product are then tested individually.

The Digital polymerase chain reaction simultaneously amplifies thousands of samples, each in a separate droplet within an emulsion.

Text Source: Wikipedia Liscence NGU

What Is a Primer ?

A primer is a strand of nucleic acid that serves as a starting point for DNA replication. They are required because the enzymes that catalyze replication, DNA polymerases, can only add new nucleotides to an existing strand of DNA. The polymerase starts replication at the 3'-end of the primer, and copies the opposite strand.

In most cases of natural DNA replication, the primer for DNA synthesis and replication is a short strand of RNA (which can be made de novo). This RNA is produced by primase, and is later removed and replaced with DNA by a repair polymerase.

Many of the laboratory techniques of biochemistry and molecular biology that involve DNA polymerase, such as DNA sequencing and the polymerase chain reaction (PCR), require primers. These primers are usually short, chemically synthesized oligonucleotides, with a length of about twenty bases. They are hybridized to a target DNA, which is then copied by the polymerase.

Uses of synthetic primers

DNA sequencing is used to determine the nucleotides in a DNA strand; the chain termination method (dideoxy sequencing or Sanger method) uses a primer as a start marker for the chain reaction.

In PCR, primers are used to determine the DNA fragment to be amplified by the PCR process. The length of primers is usually not more than 30 nucleotides, and they match exactly the beginning and the end of the DNA fragment to be amplified. They direct replication towards each other - the extension of one primer by polymerase then becomes the template for the other, leading to an exponential increase in the target segment.
It is worth noting that primers are not essentially always necessary for DNA synthesis and can in fact be used by viral polymerases, e.g. influenza, for RNA synthesis.

PCR primer design

The melting temperature of a primer is defined as the temperature at which 50% of that same DNA molecule species form a stable double helix and the other 50% have been separated to single strand molecules. The melting temperature required increases with the length of the primer. Primers that are too short would anneal at several positions on a long DNA template, which would result in non-specific copies. On the other hand, the length of a primer is limited by the temperature required to melt it. Melting temperatures that are too high, i.e., above 80°C, can also cause problems since the DNA polymerases used for PCR are less active at such temperatures. The optimum length of a primer is generally from 20 to 30 nucleotides with a melting temperature between about 55°C and 65°C.

Pairs of primers should have the similar melting temperatures as annealing in a PCR reaction occurs for both simultaneously. A primer with a Tm significantly higher than the reaction's annealing temperature may mishybridize and extend at an incorrect location along the DNA sequence, while Tm significantly lower than the annealing temperature may fail to anneal and extend at all.

Primer sequences need to be chosen to uniquely select for a region of DNA, avoiding the possibility of mishybridization to a similar sequence nearby. A commonly used method is BLAST search whereby all the possible regions to which a primer may bind can be seen. Both the nucleotide sequence as well as the primer itself can be BLAST searched. Alternatively use of software such as Beacon designer, may yeild to specific primers. Mononucleotide repeats should be avoided, as loop formation can occur and contribute to mishybridization. Primers should not easily anneal with other primers in the mixture (either other copies of same or the reverse direction primer); this phenomenon can lead to the production of 'primer dimer' products contaminating the mixture. Primers should also not anneal strongly to themselves, as internal hairpins and loops could hinder the annealing with the template DNA.

Degenerate primers

Sometimes degenerate primers are used. These are actually mixtures of similar, but not identical, primers. They may be convenient if the same gene is to be amplified from different organisms, as the genes themselves are probably similar but not identical. The other use for degenerate primers is when primer design is based on protein sequence. As several different codons can code for one amino acid, it is often difficult to deduce which codon is used in a particular case. Therefore primer sequence corresponding to the amino acid isoleucine might be "ATH", where A stands for adenine, T for thymine, and H for adenine, thymine, or cytosine, according to the genetic code for each codon, using the IUPAC symbols for degenerate bases. Use of degenerate primers can greatly reduce the specificity of the PCR amplification. The problem can be partly solved by using touchdown PCR.

Degenerate primers are widely used and extremely useful in the field of microbial ecology. They allow for the amplification of genes from thus far uncultivated microorganisms or allow the recovery of genes from organisms where genomic information is not available. Usually, degenerate primers are designed by aligning gene sequencing found in GenBank. Differences among sequences are accounted for by using IUPAC degeneracies for individual bases. PCR primers are then synthesized as a mixture of primers corresponding to all permutations.

Text Source: Wikipedia Liscence NGU

Real-time PCR

In molecular biology, real-time polymerase chain reaction, also called quantitative real time polymerase chain reaction (qPCR) or kinetic polymerase chain reaction, is a laboratory technique based on the polymerase chain reaction, which is used to amplify and simultaneously quantify a targeted DNA molecule. It enables both detection and quantification (as absolute number of copies or relative amount when normalized to DNA input or additional normalizing genes) of a specific sequence in a DNA sample.
The procedure follows the general principle of polymerase chain reaction; its key feature is that the amplified DNA is quantified as it accumulates in the reaction in real time after each amplification cycle. Two common methods of quantification are the use of fluorescent dyes that intercalate with double-stranded DNA, and modified DNA oligonucleotide probes that fluoresce when hybridized with a complementary DNA.
Frequently, real-time polymerase chain reaction is combined with reverse transcription polymerase chain reaction to quantify low abundance messenger RNA (mRNA), enabling a researcher to quantify relative gene expression at a particular time, or in a particular cell or tissue type.
Although real-time quantitative polymerase chain reaction is often marketed as RT-PCR, it should not be confused with reverse transcription polymerase chain reaction, also known as RT-PCR.
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Background

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Cells in all organisms regulate their cellular activities by activating or deactivating the expression of their genes. Gene expression is usually directly proportional to the number of copies of messenger RNA (mRNA) of a particular gene in a cell or tissue.
Traditionally, the expression level of a gene has been estimated by visualizing the abundance of its mRNA transcript in a sample with a technique called northern blotting. In this method, purified RNA is separated by agarose gel electrophoresis, transferred to a solid matrix (such as a nylon membrane), and probed with a specific DNA probe that is complementary to the gene of interest. Although this technique is still used to assess gene expression, it requires relatively large amounts of RNA and provides only qualitative or semiquantitative information of mRNA levels.
In order to robustly detect and quantify gene expression from small amounts of RNA, amplification of the gene transcript is necessary. The polymerase chain reaction is a common method for amplifying DNA; for mRNA-based PCR the RNA sample is first reverse transcribed to cDNA with reverse transcriptase.
Development of PCR technologies based on reverse transcription and fluorophores permits measurement of DNA amplification during PCR in real time, i.e., the amplified product is measured at each PCR cycle. The data thus generated can be analysed by computer software to calculate relative gene expression in several samples, or mRNA copy number. Real-time PCR can also be applied to the detection and quantification of DNA in samples to determine the presence and abundance of a particular DNA sequence in these samples.

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Real-time PCR using double-stranded DNA dyes

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A DNA-binding dye binds to all double-stranded (ds)DNA in a PCR reaction, causing fluorescence of the dye. An increase in DNA product during PCR therefore leads to an increase in fluorescence intensity and is measured at each cycle, thus allowing DNA concentrations to be quantified. However, dsDNA dyes such as SYBR Green will bind to all dsDNA PCR products, including nonspecific PCR products (such as "primer dimers"). This can potentially interfere with or prevent accurate quantification of the intended target sequence. The reaction is prepared as usual, with the addition of fluorescent dsDNA dye. The reaction is run in a thermocycler, and after each cycle, the levels of fluorescence are measured with a detector; the dye only fluoresces when bound to the dsDNA (i.e., the PCR product). With reference to a standard dilution, the dsDNA concentration in the PCR can be determined.
Like other real-time PCR methods, the values obtained do not have absolute units associated with it (i.e. mRNA copies/cell). As described above, a comparison of a measured DNA/RNA sample to a standard dilution will only give a fraction or ratio of the sample relative to the standard, allowing only relative comparisons between different tissues or experimental conditions. To ensure accuracy in the quantification, it is usually necessary to normalize expression of a target gene to a stably expressed gene (see below). This can correct possible differences in RNA quantity or quality across experimental samples.

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Fluorescent reporter probe method

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Using fluorescent reporter probes is the most accurate and most reliable of the methods, but also the most expensive. It uses a sequence-specific RNA or DNA-based probe to quantify only the DNA containing the probe sequence; therefore, use of the reporter probe significantly increases specificity, and allows quantification even in the presence of some non-specific DNA amplification. This potentially allows for multiplexing - assaying for several genes in the same reaction by using specific probes with different-coloured labels, provided that all genes are amplified with similar efficiency.
It is commonly carried out with an RNA-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. 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 reaction is prepared as usual 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 the real-time PCR thermocycler, and its geometric increase corresponding to exponential increase of the product is used to determine the threshold cycle (CT) in each reaction.

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Quantitation

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Quantitating gene expression by traditional methods presents several problems. Firstly, detection of mRNA on a Northern blot or PCR products on a gel or Southern blot is time-consuming and does not allow precise quantitation. Also, over the 20-40 cycles of a typical PCR reaction, the amount of product reaches a plateau determined more by the amount of primers in the reaction mix than by the input template/sample.
Relative concentrations of DNA present during the exponential phase of the reaction are determined by plotting fluorescence against cycle number on a logarithmic scale (so an exponentially increasing quantity will give a straight line). A threshold for detection of fluorescence above background is determined. The cycle at which the fluorescence from a sample crosses the threshold is called the cycle threshold, Ct. Since the quantity of DNA doubles every cycle during the exponential phase, relative amounts of DNA can be calculated, e.g. a sample whose Ct is 3 cycles earlier than another's has 23 = 8 times more template.
Amounts of RNA or DNA are then determined by comparing the results to a standard curve produced by RT-PCR of serial dilutions (e.g. undiluted, 1:4, 1:16, 1:64) of a known amount of RNA or DNA. As mentioned above, to accurately quantify gene expression, the measured amount of RNA from the gene of interest is divided by the amount of RNA from a housekeeping gene measured in the same sample to normalize for possible variation in the amount and quality of RNA between different samples. This normalization permits accurate comparison of expression of the gene of interest between different samples, provided that the expression of the reference (housekeeping) gene used in the normalization is 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.
Applications of real-time polymerase chain reaction

There are numerous applications for real-time polymerase chain reaction in the laboratory. It is commonly used for both diagnostic and research applications.
Diagnostically real-time PCR is applied to rapidly detect the presence of genes involved in infectious diseases, cancer and genetic abnormalities. In the research setting, real-time PCR is mainly used to provide highly sensitive 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.
Also, the technique is used in Environmental microbiology, for example to quantify resistance genes in water samples.

Text Source: Wikipedia Liscence NGU