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Polymerase Chain Reaction (PCR) Steps

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We owe the discovery of the polymerase chain reaction to Kary B Mullis in the year 1983. He was the actual proponent of PCR. Few people are aware that in 1971, Kleppe and the Nobel laureate Gobind Khorana published studies including a description of techniques that are now known to be the basis for nucleic acid replication. However, it is unfortunate that Kleppe and Khorana were ahead of their times. Oligonucleotide synthesis wasn't as simple as it is today; genes had not been sequenced and the idea of thermostable DNA polymerases had not been described. Hence, the credit for discovering the PCR remains with Kary Mullis.

The Polymerase Chain Reaction is essentially a cell-free method of DNA and RNA cloning. The DNA or RNA is isolated from the cell and replicated upto a million times. At the end, what you get is a greatly amplified fragment of DNA. The PCR is quick, reliable and sensitive and its variations have made it the basis of genetic testing.


"I was just driving and thinking about ideas and suddenly I saw it. I saw the polymerase chain reaction as clear as if it were up on a blackboard inside my head, so I pulled over and started scribbling." A chemist friend of his was asleep in the car. Mullis says that "Jennifer objected groggily to the delay and the light, but I exclaimed I had discovered something fantastic. Unimpressed, she went back to sleep."

Mullis kept scribbling calculations, right there in the car. He convinced the small California biotech company, Cetus, he was working for at that time, that he was up to something big. They finally listened. They sold the patent of PCR to Hoffman-LaRoche for a staggering $300 million - the maximum amount of money ever paid for a patent. Mullis meanwhile received a $10,000 bonus.


The purpose of PCR is to generate a huge number of copies of a segment of DNA, which could be a gene, a portion of a gene, or an intronic region. There are three major steps in a PCR, which are repeated for 30 or 40 cycles. This is done on an automated cycler, which can either heat or cool the tubes containing the reaction mixture, as required, in a very short period of time. There are three major steps in a PCR, which are repeated for 30 or 40 cycles.

Denaturation--During this process, the double stranded DNA melts and opens to form single stranded DNA. All enzymatic reactions, such as those carried over from a previous cycle, stop. This will be explained in the next paragraph. The temperature for denaturation is not fixed but it usually occurs at about 95°C. It is important to realize that the denaturation temperature is largely dependent on G:C (guanine:cytosine) content of the DNA fragment to be analyzed. This is reasonable when one considers that the G:C bond is a triple hydrogen bond and the AT bond is a double bond. Logic dictates that a triple bond should be 1.5 times harder to break than a double bond. Therefore, when the segment of DNA to be analyzed has a very high G:C content, the denaturation temperature can reach even upto 99°C.

Annealing--This requires temperatures lower than those required for denaturation. In this process, the primers anneal to that very specific segment of DNA that is to be amplified. The primers are jiggling around, caused by the Brownian motion. Ionic bonds are constantly formed and broken between the single stranded primer and the single stranded template. The more stable bonds last a little bit longer (primers that fit exactly) and on that little piece of what is now double stranded DNA (template and primer); the polymerase can attach and starts copying the template. Once there are a few bases built in, the ionic bond is so strong between the template and the primer, that it does not break anymore.

Extension--This is done at 72°C. This is the ideal temperature for working with polymerase. The primers, which are complementary to the template, already have a strong ionic attraction to the template. This force is stronger than the forces breaking these attractions i.e. the high temperature. Primers that are on positions with no exact match (non complementary) get loose again (because of the higher temperature) and don't give an extension of the fragment. The nucleotide bases are added from the 5' end to the 3' end. The phosphate group of the dNTPs is coupled with the hydroxyl group of the extending DNA strand. The extension time depends on two factors; the type of polymerase used and the length of the DNA fragment to be amplified. Usually, Taq polymerase adds dNTPs at the rate of about 1000 bases per minute.

It is important to realize that each component of the PCR including the input DNA, the oligonucleotide primers, the thermostable polymerase, the buffer and the cycling parameters has a profound impact on the sensitivity, specificity and fidelity of the reaction.

The three steps of the first cycle are shown, that is, denaturation, annealing and extension. At the end of the first cycle, two strands have been synthesized. At the end of the second cycle, four strands have been synthesized (the three steps of the cycle have not been shown). At the end of the third cycle, eight strands have been synthesized. The number of strands increases exponentially with each cycle.


  • The Polymerase Chain Reaction is essentially a cell-free method of cloning DNA and RNA.
  • There are three steps involved in every cycle; these are denaturation, annealing and extension.
  • At the end of each cycle, the DNA doubles. Therefore, theoretically, if there are 'n' cycles in a reaction, the number of DNA fragments at the end of the reaction will be 2n.


The components that are essential for a successful PCR are elaborated here.


This is that portion of the DNA/gene that is to be amplified. Usually the concentration is 100 ng genomic DNA per PCR reaction. However, this can vary depending on the target gene concentration and the source of DNA. The PCR reaction is inherently sensitive. It is not necessary for the template DNA to be abundant or highly purified. Higher amounts of template DNA can increase the yield of nonspecific PCR products, but if the fidelity of the reaction is crucial, one should limit both template DNA quantities as well as the number of PCR cycles.

DNA in solution may contain a large number of contaminants. These contaminants may inhibit the PCR. Some of these reagents are phenol, EDTA, and proteinase K, which can inhibit Taq DNA polymerase. However, isopropanol precipitation of DNA and washing of DNA pellets with 70% ethanol is usually effective in removing traces of contaminants from the DNA sample.

Effects of Fixation

This is of particular interest to the pathologist since he has to deal with formalin fixed tissue. DNA extracted from fresh tissue or cell suspensions forms an optimal template for PCR. The tissue is best stored at -70°C at which the nucleic acids can be stored indefinitely. A temperature of -20°C is sufficient to preserve the DNA for several months and at 4°C, the DNA can be stored for several weeks. At room temperature, the DNA has been successfully stored for hours to days; however, mitochondrial DNA is very sensitive to temperature and may degrade in thawed tissues.

DNA extracted from fixed tissue has been used successfully for PCR. The type of fixative and the duration of fixation are of critical importance. Non crosslinking fixatives like ethanol provide the best DNA. Formaldehyde is variable in its DNA yield. Carnoy's, Zenker's and Bouin's are poor fixatives as far as DNA preservation is concerned.

Not surprisingly, formaldehyde is the fixative which has been evaluated the most, because it is more commonly used worldwide. The studies have demonstrated that a successful PCR depends on the protocol to extract the DNA and the length of fixation. Formaldehyde reacts with DNA and proteins to form labile hydroxymethyl intermediates which give rise to a mixture of end products which include DNA-DNA and DNA-protein adducts. Purification of DNA from formalin fixed tissue, therefore, includes heating to reverse the hydroxymethyl additions and treatment with a proteinase to hydrolyze the covalently linked proteins. However, there is no way to reverse the DNA-DNA links and these links inhibit the DNA polymerases. This accounts for the low PCR yield which is seen with formalin fixed tissue. Usually, the PCR reaction with formalin fixed DNA as a template yields products which are not more than 600 bp in size.


  • Template DNA is required in a concentration of 100ng for each PCR reaction. Contaminants in DNA may inhibit the reaction.
  • Fixation of tissues provides DNA which is not as good as DNA obtained from fresh/ frozen tissues.
  • Different fixatives give different DNA yields. Alcohol is the best fixative and Carnoy's, Zenker's and Bouin's are poor fixatives as far as DNA preservation is concerned. Formalin is intermediate in DNA yield.
  • Purification of DNA from formalin fixed tissue involves heating to reverse the attachment of hydroxymethyl intermediates and treatment with a proteinase to hydrolyze the covalently linked proteins.
  • The DNA obtained after fixation can be used for reactions in which the PCR product is not more than 600 bp.


The purpose of using buffers in PCR is to provide optimum pH and potassium ion concentration for the DNA polymerase enzyme (usually obtained from bacteria 'Thermus aquaticus') to function. Most buffers are available in a 10X concentration and require dilution before use. Although most protocols recommend the final buffer concentration of 1X, a concentration of 1.5X might result in increased PCR product yield.

The PCR buffer contains many components. Some important ones are discussed here:

Divalent and monovalent cations - These are required by all thermostable DNA polymerases. Mg2+ is the divalent cation that is usually present in most of the PCR buffers. Some polymerases also work with buffers containing Mn2+. Calcium containing buffers are ineffective and therefore, rarely used. Buffers can be divided into first and second generation buffers on the basis of their ionic component. The second generation buffers, as opposed to first generation buffers, also contain (NH4)2SO4 and permit consistent PCR product yield and specificity over a wide range of magnesium concentration (1.0 to 4.0 mM MgCl2). The overall specificity and yield of PCR products is better with second generation buffers, as compared with first generation PCR buffers. Buffers also contain KCl. Salts like KCl and NaCl may help to facilitate primer annealing, but concentration of 50 mM will inhibit Taq polymerase activity. Interactions between K+ and NH4+ allow specific primer hybridization over a broad range of temperatures. Magnesium is one of the most important components of the buffer. Mg2+ ions form a soluble complex with dNTPs which is essential for dNTP incorporation; they also stimulate polymerase activity and influence the annealing efficiency of primer to template DNA. The concentration of MgCl2 can have a dramatic effect on the specificity and yield of PCR products. Optimal concentration of MgCl2 is between 1.0 to 1.5 mM for most reactions. Low MgCl2 concentration helps to eliminate non-specific priming and formation of background PCR products. This is desirable when fidelity of DNA synthesis is critical. At the same time, however, too few Mg2+ ions can result in low yield of PCR products. High MgCl2 concentration helps to stabilize interaction of the primers with their intended template, but can also result in nonspecific binding and formation of non specific PCR products. It is important to be aware that many PCR buffers (often sold in 10X stocks) already contain some amount of MgCl2. Therefore, the addition of further amounts must be carefully monitored. In the best possible scenario, the PCR would work well with the amount of Mg2+ already present in the buffer solution. However, if this does not occur, it is necessary to standardize the amount of Mg2+ in the reaction mix. This can be difficult because the dNTPs and the oligonucleotide primers bind to Mg2+. Therefore, the molar concentration of Mg2+ must exceed the molar concentration of the phosphate groups contributed by dNTPs and the primers. As a rule of thumb, the magnesium concentration in the reaction mixture is generally 0.5 to 2.5 mM greater than the concentration of dNTPs. The optimal concentration of Mg2+ should, therefore, be standardized for each reaction.

Tris-Cl - The concentration of tris-Cl is adjusted so that the pH of the reaction mixture is maintained between 8.3 and 8.8 at room temperature. In standard PCR reactions, it is usually present in a concentration of 10mM. When incubated at 72°C which is the temperature for extension, the pH of the reaction mixture falls by more than a full unit, producing a buffer whose pH is 7.2.

Other components - Some buffers also contain components like BSA (Bovine serum albumin) and DMSO (dimethyl sulphoxide). BSA reduces the amount of template sticking to the side of the tube, making it available for amplification and reducing the risk of primer dimer. Primer dimers are products obtained when the primers anneal to each other instead to to the template DNA. DMSO has been shown to facilitate DNA strand separation (in GC rich difficult secondary structures) because it disrupts base pairing and has been shown to improve PCR efficiency.

In effect, it is wise not to tamper with the buffer provided with the Taq polymerase. The buffer is usually standardized for the vial of Taq and there is no need to add additional MgCl2 or stabilizers like DMSO and BSA. However, some Taq buffers come with the buffer in one vial and MgCl2 in a separate vial. Under such circumstances, it is advisable to start with 1µL of MgCl2 and increase its concentration in aliquots of 0.5 µL, if the initial reaction fails.


  • The PCR buffer contains divalent and monovalent cations, Tris Cl and other components.
  • The PCR buffer is used to give the correct pH and potassium concentration for the DNA polymerase to function.
  • The most common divalent ion used is magnesium in the form of MgCl2. MgCl2 concentration is vital for PCR.
  • Tris Cl is used to maintain the pH between 8.3 and 8.8 at room temperature.
  • Salts like NaCl and KCl may facilitate primer annealing
  • Other components like BSA and DMSO help to increase the sensitivity and specificity of the reaction.


What are Oligonucleotide Primers?

PCR primers are short fragments of single stranded DNA (17-30 nucleotides in length) that are complementary to DNA sequences that flank the target region of interest. The purpose of PCR primers is to provide a free 3'-OH group to which the DNA polymerase can add dNTPs.

There are two primers used in the reaction. The forward primer anneals to the DNA minus strand and directs synthesis in a 5' to 3' direction. The sequence of primers is always represented in a 5' to a 3' direction. The reverse primer anneals to the other strand of the DNA.

How to design a primer?

The predominant goal kept in mind while designing a primer is specificity. Each member of the primer must anneal in a stable fashion to its target sequence in the template DNA. The longer the primer, the higher is its specificity. Unfortunately, the longer the primer, the less likely it is to anneal to a particular sequence in the template DNA. Conversely, if the primer length is small, it is likely to anneal, but its specificity will be poor. A compromise is reached by designing primers between 20 and 25 nucleotides long. Inclusion of less than 17 nucleotides often leads to non specific annealing, while presence of more than 25 nucleotides may not allow annealing to occur at all.

Remember that the DNA sequence in the human genome appears to be a random sequence of nucleotides. When designing primers, it is important to calculate the probability that a sequence exactly complementary to a string of nucleotides in the human genome will occur by chance. Several formulae are designed to calculate such probabilities. However, mathematical expressions are not necessarily correct and in this case, the predictions maybe wildly wrong. The distribution of codons is non random with repetitive DNA sequences and gene families. It is advisable to use primers longer than the statistically indicated minimum. It is also advisable to scan DNA databases to check if the proposed sequence occurs only in the desired gene.

For a practicing pathologist, it is best not to attempt designing of primers. What a pathologist requires is the primer sequence for an established test. If, for example, a pathologist requires primer sequence for the diagnosis of sickle cell anemia, all he has to do is search the web for papers related to molecular testing of sickle cell anemia. The primer sequences will be provided in the paper. Custom made primers can be commercially synthesized. Several biotechnology companies provide this facility. Before the primers are ordered, it is essential to check that the sequence is correct and that there are no missing nucleotides in the sequence. That is where, BLAST is invaluable.

Before the intricacies of the BLAST search are elaborated upon, it is necessary to mention that designing a primer does not depend only on the sequence of nucleotides. Other factors like the GC content and melting point are also important considerations. They will be dealt with later in the chapter.

BLAST and its uses

BLAST is an acronym for Basic Local Alignment Search Tool. It is an algorithm comparing information about primary biological sequences with a library or database of sequences.

A BLAST can be performed for different organisms, but in this book, we will concern ourselves with nucleotide BLAST in humans only. BLAST searches the database for sequences similar to the sequence of interest (the "query" sequence) by using a 2-step approach. The basic concept is that the higher the number of similar segments between two sequences, and the longer the length of similar segments, the less divergent the sequences are, and therefore, likely to be more genetically related (homologous).

Before perfoming a BLAST search the oligonucleotide sequence is first identified. The sequence is fed into the programme. BLAST first searches for short regions of a given length called "words" (W). It then searches for substrings which are compared to the query sequence. The program then aligns with sequences in the database ("target sequences"), using a substitution matrix.

For every pair of sequences (query and target) that have a word or words in common, BLAST extends the search in both directions to find alignments that score greater (are more similar) than a certain score threshold (S). These alignments are called high scoring pairs or HSPs; the maximal scoring HSPs are called maximum segment pairs (MSPs).

The BLAST search as outlined in fig 7.2 shows the results of the search. If we scroll down further, we can see the sequences producing significant alignments. Note that in this BLAST search, there are 49 BLAST hits in the query sequence.

In the list shown in figure 7.2, there is a list of hits starting with the best (most similar). To the right of the screen is the E-value. This is the expected number of chance alignments; the lower the E value, the more significant the score. First in the list is the sequence finding itself, which obviously has the best score. To the left is the accession number. This refers to a unique code that identifies a sequence in a database.

It is important to know that there is no set cut-off that determines whether a match is significant or "similar enough". This must be determined according to the goals of the project.

The sequences provided in the figure 7.2 show a significant alignment with Pseudomonas japonica. It shows a high score (bits) and a low E-value. Note that the lower the E value, the greater the likelihood that the sequence is a good match.


BLAST output can be delivered in a variety of formats. These formats include HTML, plain text and XML formatting. For the NCBI's web-page, the default format for output is HTML. When performing a BLAST on NCBI (National Centre for Biotechnology Information), the results are displayed in a graphical format showing the following:

  1. The hits found
  2. A tabular form showing sequence identifiers for the hits with scoring related data
  3. Alignments for the sequence of interest and the hits received with corresponding BLAST scores for these.

The easiest to read and most informative of these is probably the table. The main idea of BLAST is that there are often high-scoring segment pairs (HSP) in a statistically significant alignment. BLAST searches for these high scoring sequence alignments between the query sequence and the sequences in the database. The speed and relatively good accuracy of BLAST are among the key technical innovations of the BLAST programs.

Sequence of events to be followed when performing a BLAST search.:

  • Go to PUBMED (http://www.ncbi.nlm.nih.gov/pubmed/)
  • Scroll down to reach a heading called 'POPULAR'
  • Under 'POPULAR' click on 'BLAST'
  • Click on 'nucleotide blast'
  • Under the heading, enter accession number(s), gi(s), or FASTA sequence(s), type or paste the sequence that you want matched.
  • Click BLAST
  • Wait for the results. Analyse the nucleotide sequence as it appears.

Calculation of Melting Temperature

The melting temperature or Tm is a measure of stability of the duplex formed by the primer and the complementary target DNA sequence and is an important consideration in primer design. Tm corresponds to the midpoint in transition of DNA from the double stranded to its single stranded form. A higher Tm permits an increased annealing temperature that makes sure that the annealing between the target DNA and the primer is specific. The Tm is dependent on the length of the oligonucleotides and the G+C content of the primer. The formula for calculation of Tm is given in table 7.1.

Table 7.1: Formula for calculation of the melting temperature.

Length of Primer

Tm (°C)

Less than 20 nucleotides long

2(effective length*)

20 to 35 nucleotides long

22 + 1.46(effective length)

*Effective length = 2(number of G+C) + number of (A + T)

Primers are usually designed to avoid matching repetitive DNA sequences. This includes repeats of a single nucleotide.. The two primers in a PCR reaction are not homologous to each other and their complementarity can lead to formation of spurious amplification artifacts called primer dimers. The 3' end of a primer is most critical for initiating polymerization.

The rules for selecting primers in addition to those already mentioned are as follows:

The C and G nucleotides should be distributed uniformly throughout the primer and comprise approximately 40% of the bases. More than three G or C nucleotides at the 3'-end of the primer should be avoided, as nonspecific priming may occur.

The primer should be neither self-complementary nor complementary to any other primer in the reaction mixture, in order to avoid formation of primer-dimer or hairpin-like structure.

All possible sites of complementarity between the primer and the template DNA should be noted.

The melting temperature of flanking primers should not differ by more than 5°C. Therefore, the G+C content and length must be chosen accordingly (a higher G+C content means a higher melting temperature).

The PCR annealing temperature (TA) should be approximately 5°C lower than the primer melting temperature.

G+C content in each primer should not be more than 60% to avoid formation of internal secondary structures and long stretches of any one base.

Primer extension will occur during the annealing step. Primers are always present in an excess concentration in conventional (symmetric) PCR amplification and, typically, are within the range of 0.1M to 1M. It is generally advisable to use purified oligomers of the highest chemical integrity.

Primer Dimers

A Primer Dimer (PD) consists of primer molecules that have attached or hybridized to each other because of strings of complementary bases in the primers. As a result, the DNA polymerase amplifies the PD, leading to competition for PCR reagents, thus potentially inhibiting amplification of the DNA sequence targeted for PCR amplification.

In the first step of primer dimer formation, two primers anneal at their respective 3' ends. The DNA polymerase will bind and extend the primers. In the third step, a single strand of the product of step II is used as a template to which fresh primers anneal leading to synthesis of more PD product.

Primer dimers may be visible after gel electrophoresis of the PCR product. In ethidium bromide stained gels, they are typically seen as 30-50 base-pair (bp) bands or smears of moderate to high intensity. They can be easily distinguished from the band of the target sequence, which is typically longer than 50 bp.

One approach to prevent PD formation consists of physical-chemical optimization of the PCR system, i.e., changing the concentration of primers, MgCl2, nucleotides, ionic strength and temperature of the reaction. Reducing PD formation may also result in reduced PCR efficiency. To overcome this limitation, other methods aim to reduce the formation of PDs only. These include primer design, and use of different PCR enzyme systems or reagents.


  • Oligonucleotide primers are short fragments of single stranded DNA (17-30 nucleotides in length) that are complementary to DNA sequences that flank the target region of interest. They dictate which region of DNA in the PCR will be amplified.
  • Primer sequences can be obtained by reviewing previously published literature. A confirmation of the sequence can be done by using BLAST (Basic Local Alignment Search Tool).
  • The melting temperature is the midpoint in the observed transition from a double stranded to a single stranded form. A higher annealing temperature ensures that the annealing between the target DNA and the primer is specific.
  • A primer dimer consists of primer molecules that have attached or hybridized to each other because of strings of complementary bases in the primers. Taq polymerase amplifies the primer dimer leading to competition for the PCR products.
  • Several methods are used to reduce primer dimer formation including changing the concentrations of primers, MgCl2, nucleotides, ionic strength and temperature of the reaction.


The initial PCR reaction used the Klenow fragment of Escherichia coli DNA polymerase. However, this was unstable at high temperatures and it was necessary to add a fresh aliquot of enzyme after every denaturation step. The annealing and extension temperatures had to be kept low and as a result, there was formation of non specific products in abundance. The discovery of the thermostable Taq DNA polymerases ensured that the PCR did not remain a laboratory curiosity. The extension and annealing temperatures could now be kept high and the formation of non specific products was greatly reduced.

Taq became famous for its use in the polymerase chain reaction and was called the 'Molecule of the Year' by the journal 'Science'.

Why Taq?

Taq is the enzyme of choice in PCR because of the following reasons:

Taq works best at 75°C--80°C, allowing the elongation step to occur at temperatures which make non-Watson-Crick base pairing a rare event.

It can add upto 1,000 nucleoside triphosphates to a growing DNA strand.

Taq has a half-life of 40 minutes at 95°C and 9 minutes at 97.5°C, and can replicate a 1000 base pair strand of DNA in less than 10 seconds at 72°C.

Because of all these properties, Taq is the enzyme of choice in the PCR.

How does Taq polymerase act?

The first requirement is a primer. The primer is annealed to the template strand having free hydroxyl group at its 3' end. During the extension phase, the Taq synthesizes a new DNA strand complementary to the template by adding dNTPs in a 5' to 3' direction condensing the 5' phosphate group of the dNTPs with the 3' hydroxyl group of the end of the extending DNA strand. Since Taq works best between 70°C- 80°C, a temperature of 72°C is usually chosen as the optimum annealing temperature.

Where does Taq come from?

In Thermus aquaticus, Taq polymerase is expressed at very low levels and commercial production is not economically viable. However, the enzyme can now be produced from different versions of the engineered Taq gene so as to obtain high levels of expression in E coli.

What other polymerases are available for use in PCR?

Taq is not the only polymerase; other polymerases are available but Taq is the one that is generally used in a PCR. A few other polymerases with their uses are as follows:

PFU DNA polymerase -Found in Pyrococcus furiosus, it functions in vivo to replicate the organism's DNA. The main difference between Pfu and alternative enzymes is the Pfu's superior thermostability and 'proofreading' properties compared to other thermostable polymerases. Unlike Taq DNA polymerase, Pfu DNA polymerase possesses 3' to 5' exonuclease proofreading activity, meaning that it works its way along the DNA from the 3' end to the 5' end and corrects nucleotide-misincorporation errors. This means that Pfu DNA polymerase-generated PCR fragments will have fewer errors than Taq-generated PCR inserts. As a result, Pfu is more commonly used for molecular cloning of PCR fragments than the historically popular Taq. However, Pfu is slower and typically requires 1-2 minutes to amplify 1kb of DNA at 72° C. Pfu can also be used in conjunction with Taq polymerase to obtain the fidelity of Pfu with the speed of Taq polymerase activity.

TFL DNA polymerase - Obtained from Thermus flavus, it is useful for the amplification of large segments of DNA.


All DNA polymerases have an intrinsic error rate that is highly dependant on the buffer composition, pH of the buffer, dNTP concentration and the sequence of the template itself. The types of errors that are introduced are frameshift mutations, single base pair substitutions, and spontaneous rearrangements. Therefore, the PCR reaction generates a product that is very similar, but in many cases, not identical to the original sequence. The quantity of dissimilar product obtained is obviously related to the cycle in which the mismatch took place. Under normal circumstances, this does not make any difference; however, these errors may become significant during sequencing when the role of fidelity comes into play.

Fidelity is the ability of the polymerases to avoid the incorporation of wrong nucleotides during the reaction. Under normal circumstances, it really does not make a difference if a wrong nucleotide is incorporated because the size of the PCR product remains the same and that is what we have to look for. However, there are some polymerases like Pfu which have a high fidelity. In addition to reading from the 5' to the 3' direction, they can also read from the 3' to the 5' direction and correct the wrong nucleotides which have been incorporated. This is called proofreading.

One of the major disadvantages of Taq polymerases is related to its low replication fidelity. As Taq does not have 3' to 5' exonuclease proofreading mechanism to replace an accidental mismatch in the newly synthesized DNA strand, it produces more errors than proofreading polymerases such as Pfu. Pfu on the other hand has an active 3' to 5' proofreading exonuclease activity and thus, generates a more specific PCR product. This is particularly important in further downstream applications like cloning and sequencing.


Taq is always stored in glycerol, which is a viscous fluid and rises slowly in the pipette. When added to the reaction mixture, Taq, therefore, settles at the bottom in a viscous swirl and can be easily visualized in the tube. One should always remember that it is important to mix Taq well. If mixed well with the rest of the reagents, the viscous fluid at the bottom of the tube is no longer visible.

Since Taq is stored in glycerol, it does not freeze. One should not bother trying to thaw Taq, as this will only reduce the enzyme activity. As far as possible, Taq should not be exposed to room temperature; it should be kept on ice when being aliquoted and otherwise stored at -20°C.


  • Taq polymerase is an enzyme obtained from Thermus aquaticus. It works best at 75°C - 80°C. It can replicate a 1000 base pair strand of DNA in less than 10 seconds at 72°C.
  • It synthesizes a new DNA strand complementary to the template by adding dNTP's that are complementary to the template in a 5' to 3' direction.
  • Other polymerases are Pfu from Pyrosus furiosus and Tfl from Thermus flavus.
  • Fidelity is the ability of the polymerase to avoid the incorporation of wrong nucleotides during the reaction. This is done by reading nucleotides from the 3' to the 5' direction in addition to the normal 5' to 3' direction. This can therefore, correct the wrong nucleotides which have been incorporated. This is called proofreading.
  • Taq is always stored in glycerol. Since Taq is stored in glycerol, it does not freeze. Taq should not be exposed to room temperature; it should be kept on ice when it is being aliquoted and stored at -20°C when not in use.


Standard PCR requires equimolar concentrations of dATP, dCTP, dGTP and dTTP. dATP, dCTP, dGTP and dTTP are different types of dNTPs. Concentrations of 200 to 250 µM of each dNTP are recommended for Taq polymerase in reactions containing 1.5 mM MgCl2. Higher concentrations of dNTPs are inhibitory because they sequester MgCl2. The stock solution should be free from pyrophosphates which inhibit PCR. The stock solution contains NaOH which adjusts the pH of the solution to 8.1. This protects dNTP from damage during freezing and thawing. To be on the safe side, the dNTPs should be stored in small aliquots and discarded after two to three cycles of freezing and thawing. If the vials are stored for a long time, water evaporates and it maybe necessary to centrifuge the mixture in order to avoid changes in concentration.


The denaturation temperature is critically dependant on the G:C content of the DNA fragment to be amplified. For calculation of the denaturation temperature, see the calculations for melting temperature above. (Page 10 - Calculation of melting temperatures)

The annealing temperature is the most critical temperature in the reaction. It is usually 5°C below the calculated Tm of the primer. In some reactions, the annealing temperature maybe kept close to Tm in order to avoid formation of non specific products; however the annealing temperature should be carefully titrated to create balance between the production of non specific products and no formation of products at all.

The extension phase reactions are usually performed at 70°C - 75°C and are related to the optimum temperature reaction of the thermostable polymerase.

A typical PCR reaction involves 25 to 35 cycles. Higher number of cycles does not lead to a significant increase in the amount of the PCR product. However, some PCRs can stretch up to 40 cycles.


  • Standard PCRs contain equimolar concentration of dATP, dCTP, dGTP and dTTP.
  • Concentrations of 200 to 250 µM of each dNTP are recommended.
  • The cycling parameters include the denaturation temperature, the annealing and the extension temperatures.
  • Each PCR has 25 to 35 cycles. A greater number of cycles does not significantly increase the PCR product.


  • Simple, Quick and Inexpensive--The reaction is simple. It is also much faster. Visualization is done using ethidium bromide. This eliminates the need for radioactivity, which is used in hybridization techniques.
  • High Sensitivity and Specificity--The PCR can detect one abnormal cell in a background of 105 abnormal cells. It can also be used to analyze single copy genes from individual cells. The PCR can detect a broad range of genetic abnormalities ranging from gross structural alterations like translocations and deletions to point mutations within a specific gene. The primer design is important and it dictates the sensitivity of the reaction. However, for the pathologist, the sensitivity and specificity does not translate to diagnostic sensitivity and specificity. The sensitivity of the PCR may not reflect the diagnostic utility of this method in clinical practice.
  • Ease of PCR product labeling--Direct labeling of PCR product DNA is easily accomplished using a flurochrome which can be attached to the 5' end of either or both the oligonucleotide primers. The PCR can also be labeled using radiolabelled dNTPs (32P or 35S). This allows further evaluation of the PCR product for mutations by techniques like Single Strand Conformational Polymorphism (SSCP) and Denaturing Gradient Gel Electrophoresis.
  • Phenotpe Genotype Correlations--Microdissection and precise collection of individual cells by laser capture microdissection technique allows only the affected cells to be removed for analysis. This allows considerable genotype phenotype correlation. In situ PCR performed on the tissue itself is perhaps the best method for correlating genotype and phenotype.


  • PCR analyses only the target region - Unlike conventional cytogenetics or southern and northern hybridization, the PCR fails to identify structural changes that do not alter the sequence of the target gene itself.
  • Inability to analyse large mutations - Some mutations including large insertions or inversions alter the structure of the target region in such a way that it cannot be amplified. Mutations that damage the primer binding site also do not allow amplification.
  • Amplification bias - In PCR, some templates are amplified in preference to others. This is because of factors such as the length of amplicons, random variation in the target number, and random variation in the PCR efficiency with each cycle. These factors can cause 10 to 30 fold differences in the amplification efficiency and this is sufficient to interfere with quantitative PCR and loss of heterozygosity analysis.
  • Spurious PCR products - Formation of spurious PCR products occurs at low levels even in reactions using well designed primers and stringent PCR conditions. These spurious PCR products consist of fragments of irrelevant genes.
  • Technical issues - Non specific PCR inhibitors include detergents, phenol, heparin, dyes like bromophenol blue and hemoglobin. The most important problem is degradation of nucleic acids especially when extracted from fixed tissues. The risk of cross contaminating with another sample is very high in PCR. This can be avoided by strict attention to laboratory techniques, physical separation at various stages in the technique, use of aerosol barrier tips, regular UV radiation of the laboratory workbenches to degrade any contaminating DNA and the use of positive and negative controls.


With PCR, unfortunately, things tend to go wrong and very often one is not really sure why things have gone wrong. Given here is a comprehensive checklist which can guide the user if there is trouble.

Many short non specific products are formed - This could occur primarily because of three problems; the first is that the annealing temperature is very low and as a result, the primers have annealed to multiple regions of the DNA. The second problem could be that there is a problem with the magnesium, potassium or dNTPs concentrations. The third problem could be that the primers are incorrectly selected. Troubleshooting would therefore include:

  • Increasing annealing temperature and/or annealing time
  • Increasing extension time and/or extension temperature.
  • Decreasing the amount of buffer thereby decreasing the KCl concentration
  • Increasing MgCl2 concentration up to 3-4.5 mM but keeping dNTP concentration constant
  • Taking less primer and/or less DNA template and/or less Taq polymerase
  • Rechecking the primers using a BLAST search

Many long non specific products are formed - This could occur because of high annealing temperature; incorrect magnesium, potassium and dNTP concentrations; or incorrect selection of primers. Troubleshooting would therefore include:

  • Decreasing annealing time and/or extension time and/or extension temperature
  • Increasing annealing temperature
  • Increasing the amount of buffer and thereby increasing the KCl concentration
  • Increasing MgCl2 concentration up to 3-4.5 mM but keeping dNTP concentration constant.
  • Taking less primer and/or DNA template and/or less Taq polymerase
  • Rechecking the primers using a BLAST search
  • No product is formed at all - The problems could be in the reagents, the template or the cycling parameters.

In the template, it is likely that secondary structures have formed. Under such circumstances, adjuvants like DMSO, BSA or glycerol need to be added. One can also try a hot start or touch down PCR. It is also likely that the DNA was dirty or it contained inhibitors. The ethanol should have completely evaporated. The DNA can also be diluted if there is a suspicion of inhibitors in the DNA.

In the cycling parameters, it is likely that the annealing temperature was not optimal. A gradient PCR with 2°C increments can be tried. Also, it may be possible that there were inadequate number of cycles or a short extension time; for long products (>2kb), extension time (in minutes) should be approximately equal to the number of kb in the amplicon. In such cases, these need to be increased.

In the reagents, the problem could be in the divalent ion concentrations. Increments of 0.5mM in MgCl2 gradient can be tried. The final concentration should be kept between 1.5 and 4.0 mM. Instead of a KCl based buffer, an NH4 based buffer can be tried for greater yield. An increase in the template, primer and Taq concentrations can also be attempted.

It is also possible that the primers were poorly designed. If so, they need to be redesigned and a BLAST search needs to be done to confirm that the primers are suitable.

Finally, it is possible that there was an overabundance of primer or template; in either case, they need to be decreased.

The reaction was working before but there is no product formed now - This is perhaps the most frustrating problem for a person who is operating the PCR. It is necessary to check all the reagents once again. The dNTP solution should be changed since dNTPs are very prone to freeze thaw cycles. If the primers are new, the sequence should be checked for correctness by using BLAST. The primer and template amounts should be increased and the reaction is run again. A decrease in either the primer or the template can cause the PCR reaction to fail. Finally, the annealing temperature should be decreased by 5°C to 10°C. If still no products are formed, it is necessary to check all the reagents once again. If there are non specific products, reset the annealing temperature and start again.

A weak PCR product is formed - It is advisable to decrease the annealing temperature to the lowest possible level. The amount of Taq, DNA template and primers can be increased. Adjuvants like BSA, DMSO or glycerol can be used. The primer sequences should be rechecked for confirming that the sequences are correct.

Smeared product - There could be a smear instead of a single band. In such cases, it maybe necessary to increase the extension time. It is also possible that the reagents are contaminated; in that case, it is necessary to have fresh aliquots of reagents. It is also important to clean and sterilize the pipettes and use filter tips to avoid contamination. The workbench should be clean and the pre and post PCR areas clearly separated.

Wrong size band amplified - It is advisable to check the gene for isoforms or splice variants. It is important to use the same thermal cycler for optimization and all future experiments. Different cyclers can vary in ramping speeds and temperature. Degraded dNTPs are very susceptible to freeze thawing. Hence, they should be replaced with fresh aliquots. It is also important to check any new components that have been added (eg. new batch of primers)


The advantages of PCR are: 1. It is simple, quick and inexpensive 2. It has high sensitivity and specificity 3. As compared to other methods, there is an ease of PCR product labeling 4. If only the target DNA is analysed, it allows for considerable phenotpe genotype correlations

The limitations of PCR are: 1. It analyses only the target region 2. It cannot analyse large mutations 3. There is often an amplification bias in PCR 4. PCR often forms spurious products 5. Several technical issues are involved in PCR. These include the presence of inhibitors, degradation of nucleic acids and cross contamination

Considerable problems may occur during a PCR. These include: 1. Many short non specific products are formed 2. Many long non specific products are formed 3. No product is formed at all 4. A reaction that was working properly, suddenly stops working 5. A weak or a smeared product is formed 6. A wrong area of the DNA is amplified.

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