PCR

"PCR" redirects here. For other uses, see PCR (disambiguation).

A strip of eight PCR tubes, each containing a 100μl reaction.

The polymerase chain reaction (PCR) is a technique widely used in molecular biology. It derives its name from one of its key components, a DNA polymerase used to amplify a piece of DNA by in vitro enzymatic replication. As PCR progresses, the DNA thus generated is itself used as a template for replication. This sets in motion a chain reaction in which the DNA template is exponentially amplified. With PCR it is possible to amplify a single or few copies of a piece of DNA across several orders of magnitude, generating millions or more copies of the DNA piece. PCR can be extensively modified to perform a wide array of genetic manipulations.

Almost all PCR applications employ a heat-stable DNA polymerase, such as Taq polymerase, an enzyme originally isolated from the bacterium Thermus aquaticus. This DNA polymerase enzymatically assembles a new DNA strand from DNA building blocks, the nucleotides, using single-stranded DNA as template and DNA oligonucleotides (also called DNA primers) required for initiation of DNA synthesis. The vast majority of PCR methods use thermal cycling, i.e., alternately heating and cooling the PCR sample to a defined series of temperature steps. These thermal cycling steps are necessary to physically separate the strands (at high temperatures) in a DNA double helix (DNA melting) used as template during DNA synthesis (at lower temperatures) by the DNA polymerase to selectively amplify the target DNA. The selectivity of PCR results from the use of primers that are complementary to the DNA region targeted for amplification under specific thermal cycling conditions.

Developed in 1983 by Kary Mullis,[1] PCR is now a common and often indispensable technique used in medical and biological research labs for a variety of applications.[2][3] These include DNA cloning for sequencing, DNA-based phylogeny, or functional analysis of genes; the diagnosis of hereditary diseases; the identification of genetic fingerprints (used in forensic sciences and paternity testing); and the detection and diagnosis of infectious diseases. In 1993 Mullis won the Nobel Prize in Chemistry for his work on PCR.[4] PCR is used to amplify specific regions of a DNA strand (the DNA target). This can be a single gene, a part of a gene, or a non-coding sequence. Most PCR methods typically amplify DNA fragments of up to 10 kilo base pairs (kb), although some techniques allow for amplification of fragments up to 40 kb in size.[5]

A basic PCR set up requires several components and reagents.[6] These components include:

DNA template that contains the DNA region (target) to be amplified. Two primers, which are complementary to the DNA regions at the 5' (five prime) or 3'

(three prime) ends of the DNA region. A DNA polymerase such as Taq polymerase or another DNA polymerase with a

temperature optimum at around 70°C. Deoxynucleoside triphosphates (dNTPs; also very commonly and erroneously called

deoxynucleotide triphosphates), the building blocks from which the DNA polymerases synthesizes a new DNA strand.

Buffer solution , providing a suitable chemical environment for optimum activity and stability of the DNA polymerase.

Divalent cations, magnesium or manganese ions; generally Mg2+ is used, but Mn2+ can be utilized for PCR-mediated DNA mutagenesis, as higher Mn2+ concentration increases the error rate during DNA synthesis[7]

Monovalent cation potassium ions.

The PCR is commonly carried out in a reaction volume of 10-200 μl in small reaction tubes (0.2-0.5 ml volumes) in a thermal cycler. The thermal cycler heats and cools the reaction tubes to achieve the temperatures required at each step of the reaction (see below). Many modern thermal cyclers make use of the Peltier effect which permits both heating and cooling of the block holding the PCR tubes simply by reversing the electric current. Thin-walled reaction tubes permit favorable thermal conductivity to allow for rapid thermal equilibration. Most thermal cyclers have heated lids to prevent condensation at the top of the reaction tube. Older thermocyclers lacking a heated lid require a layer of oil on top of the reaction mixture or a ball of wax inside the tube.

[edit] Procedure

Figure 2: Schematic drawing of the PCR cycle. (1) Denaturing at 94-96°C. (2) Annealing at ~65°C (3) Elongation at 72°C. Four cycles are shown here. The blue lines represent the DNA template to which primers (red arrows) anneal that are extended by the DNA polymerase (light green circles), to give shorter DNA products (green lines), which themselves are used as templates as PCR progresses.

The PCR usually consists of a series of 20 to 40 repeated temperature changes called cycles; each cycle typically consists of 2-3 discrete temperature steps. Most commonly PCR is carried out with cycles that have three temperature steps (Fig. 2). The cycling is often preceded by a single temperature step (called hold) at a high temperature (>90°C), and followed by one hold at the end for final product extension or brief storage. The temperatures used and the length of time they are applied in each cycle depend on a variety of parameters. These include the enzyme used for DNA synthesis, the concentration of divalent ions and dNTPs in the reaction, and the melting temperature (Tm) of the primers.[8]

Initialization step: This step consists of heating the reaction to a temperature of 94-96°C (or 98°C if extremely thermostable polymerases are used), which is held for 1-9 minutes. It is only required for DNA polymerases that require heat activation by hot-start PCR.[9]

Denaturation step : This step is the first regular cycling event and consists of heating the reaction to 94-98°C for 20-30 seconds. It causes melting of DNA template and primers by disrupting the hydrogen bonds between complementary bases of the DNA strands, yielding single strands of DNA.

Annealing step : The reaction temperature is lowered to 50-65°C for 20-40 seconds allowing annealing of the primers to the single-stranded DNA template. Typically the annealing temperature is about 3-5 degrees Celsius below the Tm of the primers used. Stable DNA-DNA hydrogen bonds are only formed when the primer sequence very closely matches the template sequence. The polymerase binds to the primer-template hybrid and begins DNA synthesis.

Extension/elongation step: The temperature at this step depends on the DNA polymerase used; Taq polymerase has its optimum activity temperature at 75-80°C,[10][11] and commonly a temperature of 72°C is used with this enzyme. At this step the DNA polymerase synthesizes a new DNA strand complementary to the DNA template strand by adding dNTPs that are complementary to the template in 5' to 3' direction, condensing the 5'-phosphate group of the dNTPs with the 3'-hydroxyl group at the end of the nascent (extending) DNA strand. The extension time depends both on the DNA polymerase used and on the length of the DNA fragment to be amplified. As a rule-of-thumb, at its optimum temperature, the DNA polymerase will polymerize a thousand bases per minute. Under optimum conditions, i.e., if there are no limitations due to limiting substrates or reagents, at each extension step, the amount of DNA target is doubled, leading to exponential (geometric) amplification of the specific DNA fragment.

Final elongation: This single step is occasionally performed at a temperature of 70-74°C for 5-15 minutes after the last PCR cycle to ensure that any remaining single-stranded DNA is fully extended.

Final hold: This step at 4-15°C for an indefinite time may be employed for short-term storage of the reaction.

Figure 3: Ethidium bromide-stained PCR products after gel electrophoresis. Two sets of primers were used to amplify a target sequence from three different tissue samples. No amplification is present in sample #1; DNA bands in sample #2 and #3 indicate successful amplification of the

target sequence. The gel also shows a positive control, and a DNA ladder containing DNA fragments of defined length for sizing the bands in the experimental PCRs.

To check whether the PCR generated the anticipated DNA fragment (also sometimes referred to as the amplimer or amplicon), agarose gel electrophoresis is employed for size separation of the PCR products. The size(s) of PCR products is determined by comparison with a DNA ladder (a molecular weight marker), which contains DNA fragments of known size, run on the gel alongside the PCR products (see Fig. 3).

[edit] PCR stages

The PCR process can be divided into three stages:

Exponential amplification: At every cycle, the amount of product is doubled (assuming 100% reaction efficiency). The reaction is very specific and precise.[citation needed]

Levelling off stage: The reaction slows as the DNA polymerase loses activity and as consumption of reagents such as dNTPs and primers causes them to become limiting.

Plateau: No more product accumulates due to exhaustion of reagents and enzyme.

[edit] PCR optimization

Main article: PCR optimization

In practice, PCR can fail for various reasons, in part due to its sensitivity to contamination causing amplification of spurious DNA products. Because of this, a number of techniques and procedures have been developed for optimizing PCR conditions.[12][13] Contamination with extraneous DNA is addressed with lab protocols and procedures that separate pre-PCR mixtures from potential DNA contaminants.[6] This usually involves spatial separation of PCR-setup areas from areas for analysis or purification of PCR products, and thoroughly cleaning the work surface between reaction setups. Primer-design techniques are important in improving PCR product yield and in avoiding the formation of spurious products, and the usage of alternate buffer components or polymerase enzymes can help with amplification of long or otherwise problematic regions of DNA.

[edit] Application of PCR

[edit] Isolation of genomic DNA

PCR allows isolation of DNA fragments from genomic DNA by selective amplification of a specific region of DNA. This use of PCR augments many methods, such as generating hybridization probes for Southern or northern hybridization and DNA cloning, which require larger amounts of DNA, representing a specific DNA region. PCR supplies these techniques with

high amounts of pure DNA, enabling analysis of DNA samples even from very small amounts of starting material.

Other applications of PCR include DNA sequencing to determine unknown PCR-amplified sequences in which one of the amplification primers may be used in Sanger sequencing, isolation of a DNA sequence to expedite recombinant DNA technologies involving the insertion of a DNA sequence into a plasmid or the genetic material of another organism. Bacterial colonies (E.coli) can be rapidly screened by PCR for correct DNA vector constructs[14]. PCR may also be used for genetic fingerprinting; a forensic technique used to identify a person or organism by comparing experimental DNAs through different PCR-based methods.

Some PCR 'fingerprints' methods have high discriminative power and can be used to identify genetic relationships between individuals, such as parent-child or between siblings, and are used in paternity testing (Fig. 4). This technique may also be used to determine evolutionary relationships among organisms.

Figure 4: Electrophoresis of PCR-amplified DNA fragments. (1) Father. (2) Child. (3) Mother. The child has inherited some, but not all of the fingerprint of each of its parents, giving it a new, unique fingerprint.

[edit] Amplification and quantitation of DNA

Because PCR amplifies the regions of DNA that it targets, PCR can be used to analyze extremely small amounts of sample. This is often critical for forensic analysis, when only a trace amount of DNA is available as evidence. PCR may also be used in the analysis of ancient DNA that is tens of thousands of years old. These PCR-based techniques have been successfully used on animals, such as a forty-thousand-year-old mammoth, and also on human DNA, in applications ranging from the analysis of Egyptian mummies to the identification of a Russian Tsar.[15]

Quantitative PCR methods allow the estimation of the amount of a given sequence present in a sample – a technique often applied to quantitatively determine levels of gene expression. Real-time PCR is an established tool for DNA quantification that measures the accumulation of DNA product after each round of PCR amplification.

See also Use of DNA in forensic entomology

[edit] PCR in diagnosis of diseases

PCR allows early diagnosis of malignant diseases such as leukemia and lymphomas, which is currently the highest developed in cancer research and is already being used routinely.[citation needed] PCR assays can be performed directly on genomic DNA samples to detect translocation-specific malignant cells at a sensitivity which is at least 10,000 fold higher than other methods.[citation needed]

PCR also permits identification of non-cultivatable or slow-growing microorganisms such as mycobacteria, anaerobic bacteria, or viruses from tissue culture assays and animal models. The basis for PCR diagnostic applications in microbiology is the detection of infectious agents and the discrimination of non-pathogenic from pathogenic strains by virtue of specific genes.[citation

needed]

Viral DNA can likewise be detected by PCR. The primers used need to be specific to the targeted sequences in the DNA of a virus, and the PCR can be used for diagnostic analyses or DNA sequencing of the viral genome. The high sensitivity of PCR permits virus detection soon after infection and even before the onset of disease. Such early detection may give physicians a significant lead in treatment. The amount of virus ("viral load") in a patient can also be quantified by PCR-based DNA quantitation techniques (see below).

[edit] Variations on the basic PCR technique

Main article: Variants of PCR Allele-specific PCR: This diagnostic or cloning technique is used to identify or utilize

single-nucleotide polymorphisms (SNPs) (single base differences in DNA). It requires prior knowledge of a DNA sequence, including differences between alleles, and uses primers whose 3' ends encompass the SNP. PCR amplification under stringent conditions is much less efficient in the presence of a mismatch between template and primer, so successful amplification with an SNP-specific primer signals presence of the specific SNP in a sequence.[16] See SNP genotyping for more information.

Assembly PCR or Polymerase Cycling Assembly (PCA): Assembly PCR is the artificial synthesis of long DNA sequences by performing PCR on a pool of long oligonucleotides with short overlapping segments. The oligonucleotides alternate between sense and antisense directions, and the overlapping segments determine the order of the PCR fragments thereby selectively producing the final long DNA product.[17]

Asymmetric PCR: Asymmetric PCR is used to preferentially amplify one strand of the original DNA more than the other. It finds use in some types of sequencing and hybridization probing where having only one of the two complementary stands is required. PCR is carried out as usual, but with a great excess of the primers for the chosen strand. Due to the slow (arithmetic) amplification later in the reaction after the limiting primer has been used up, extra cycles of PCR are required.[18] A recent modification on this process, known as Linear-After-The-Exponential-PCR (LATE-PCR), uses a limiting primer with a higher melting temperature (Melting temperature|Tm) than the excess primer to maintain reaction efficiency as the limiting primer concentration decreases mid-reaction.[19]

Helicase-dependent amplification : This technique is similar to traditional PCR, but uses a constant temperature rather than cycling through denaturation and annealing/extension cycles. DNA Helicase, an enzyme that unwinds DNA, is used in place of thermal denaturation.[20]

Hot-start PCR: This is a technique that reduces non-specific amplification during the initial set up stages of the PCR. The technique may be performed manually by heating the reaction components to the melting temperature (e.g., 95˚C) before adding the polymerase.[21] Specialized enzyme systems have been developed that inhibit the polymerase's activity at ambient temperature, either by the binding of an antibody [9] 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 instantly activated at elongation temperature.

Intersequence-specific (ISSR) PCR: a PCR method for DNA fingerprinting that amplifies regions between some simple sequence repeats to produce a unique fingerprint of amplified fragment lengths.[22]

Inverse PCR : a method used to allow PCR when only one internal sequence is known. This is especially useful in identifying flanking sequences to various genomic inserts. This involves a series of DNA digestions and self ligation, resulting in known sequences at either end of the unknown sequence.[23]

Ligation-mediated PCR: This method uses small DNA linkers ligated to the DNA of interest and multiple primers annealing to the DNA linkers; it has been used for DNA sequencing, genome walking, and DNA footprinting.[24]

Methylation-specific PCR (MSP): The MSP method was developed by Stephen Baylin and Jim Herman at the Johns Hopkins School of Medicine,[25] and is used to detect methylation of CpG islands in genomic DNA. DNA is first treated with sodium bisulfite, which converts unmethylated cytosine bases to uracil, which is recognized by PCR primers as thymine. Two PCRs are then carried out on the modified DNA, using primer sets identical except at any CpG islands within the primer sequences. At these points, one primer set recognizes DNA with cytosines to amplify methylated DNA, and one set recognizes DNA with uracil or thymine to amplify unmethylated DNA. MSP using qPCR can also be performed to obtain quantitative rather than qualitative information about methylation.

Miniprimer PCR: Miniprimer PCR uses a novel thermostable polymerase (S-Tbr) that can extend from short primers ("smalligos") as short as 9 or 10 nucleotides, instead of the approximately 20 nucleotides required by Taq. This method permits PCR targeting smaller primer binding regions, and is particularly useful to amplify unknown, but conserved, DNA sequences, such as the 16S (or eukaryotic 18S) rRNA gene. 16S rRNA miniprimer PCR was used to characterize a microbial mat community growing in an extreme environment, a hypersaline pond in Puerto Rico. In that study, deeply divergent sequences were discovered with high frequency and included representatives that defined two new division-level taxa, suggesting that miniprimer PCR may reveal new dimensions of microbial diversity.[26] By enlarging the "sequence space" that may be queried by PCR primers, this technique may enable novel PCR strategies that are not possible within the limits of primer design imposed by Taq and other commonly used enzymes.

Multiplex Ligation-dependent Probe Amplification (MLPA): permits multiple targets to be amplified with only a single primer pair, thus avoiding the resolution limitations of multiplex PCR (see below).

Multiplex-PCR: The use of multiple, unique primer sets within a single PCR mixture to produce amplicons of varying sizes specific to different DNA sequences. By targeting multiple genes at once, additional information may be gained from a single test run that otherwise would require several times the reagents and more time to perform. Annealing temperatures for each of the primer sets must be optimized to work correctly within a single reaction, and amplicon sizes, i.e., their base pair length, should be different enough to form distinct bands when visualized by gel electrophoresis.

Nested PCR : increases the specificity of DNA amplification, by reducing background due to non-specific amplification of DNA. Two sets of primers are being used in two successive PCRs. In the first reaction, one pair of primers is used to generate DNA products, which besides the intended target, may still consist of non-specifically amplified DNA fragments. The product(s) are then used in a second PCR with a set of primers whose binding sites are completely or partially different from and located 3' of each of the primers used in the first reaction. Nested PCR is often more successful in specifically amplifying long DNA fragments than conventional PCR, but it requires more detailed knowledge of the target sequences.

Overlap-extension PCR : is a genetic engineering technique allowing the construction of a DNA sequence with an alteration inserted beyond the limit of the longest practical primer length.

Quantitative PCR (Q-PCR): is used to measure the quantity of a PCR product (preferably real-time). It is the method of choice to quantitatively measure starting amounts of DNA, cDNA or RNA. Q-PCR is commonly used to determine whether a DNA sequence is present in a sample and the number of its copies in the sample. The method with currently the highest level of accuracy is Quantitative real-time PCR. It is often confusingly known as RT-PCR (Real Time PCR) or RQ-PCR. QRT-PCR or RTQ-PCR are more appropriate contractions. RT-PCR commonly refers to reverse transcription PCR (see below), which is often used in conjunction with Q-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 in real time.

RT-PCR : (Reverse Transcription PCR) is a method used to amplify, isolate or identify a known sequence from a cellular or tissue RNA. The PCR is preceded by a reaction using reverse transcriptase to convert RNA to cDNA. RT-PCR is widely used in expression profiling, to determine the expression of a gene or to identify the sequence of an RNA transcript, including transcription start and termination sites and, if the genomic DNA sequence of a gene is known, to map the location of exons and introns in the gene. The 5' end of a gene (corresponding to the transcription start site) is typically identified by an RT-PCR method, named RACE-PCR, short for Rapid Amplification of cDNA Ends.

Solid Phase PCR: encompasses multiple meanings, including Polony Amplification (where PCR colonies are derived in a gel matrix, for example), 'Bridge PCR' (the only primers present are covalently linked to solid support surface), conventional Solid Phase PCR (where Asymmetric PCR is applied in the presence of solid support bearing primer with sequence matching one of the aqueous primers) and Enhanced Solid Phase PCR[27] (where conventional Solid Phase PCR can be improved by employing high Tm solid support primer with application of a thermal 'step' to favour solid support priming).

TAIL-PCR: Thermal asymmetric interlaced PCR is used to isolate unknown sequence flanking a known sequence. Within the known sequence TAIL-PCR uses a nested pair of primers with differing annealing temperatures; a degenerate primer is used to amplify in the other direction from the unknown sequence.[28]

Touchdown PCR : a variant of PCR that aims to reduce nonspecific background by gradually lowering the annealing temperature as PCR cycling progresses. The annealing temperature at the initial cycles is usually a few degrees (3-5˚C) above the Tm of the primers used, while at the later cycles, it is a few degrees (3-5˚C) below the primer Tm. The higher temperatures give greater specificity for primer binding, and the lower temperatures permit more efficient amplification from the specific products formed during the initial cycles.[29]

PAN-AC: This method uses isothermal conditions for amplification, and may be used in living cells.[30][31]

Universal Fast Walking: this method allows genome walking and genetic fingerprinting using a more specific 'two-sided' PCR than conventional 'one-sided' approaches (using only one gene-specific primer and one general primer - which can lead to artefactual 'noise') [32] by virtue of a mechanism involving lariat structure formation. Streamlined derivatives of UFW are LaNe RAGE (lariat-dependent nested PCR for rapid amplification of genomic DNA ends) [33], 5'RACE LaNe [34] and 3'RACE LaNe [35].

[edit] History

Main article: History of polymerase chain reaction

A 1971 paper in the Journal of Molecular Biology by Kleppe and co-workers first described a method using an enzymatic assay to replicate a short DNA template with primers in vitro.[36] However, this early manifestation of the basic PCR principle did not receive much attention, and the invention of the polymerase chain reaction in 1983 is generally credited to Kary Mullis.[37]

At the core of the PCR method is the use of a suitable DNA polymerase able to withstand the high temperatures of >90°C (>195°F) required for separation of the two DNA strands in the DNA double helix after each replication cycle. The DNA polymerases initially employed for in vitro experiments presaging PCR were unable to withstand these high temperatures.[2] So the early procedures for DNA replication were very inefficient, time consuming, and required large amounts of DNA polymerase and continual handling throughout the process.

A 1976 discovery of Taq polymerase a DNA polymerase purified from the thermophilic bacterium, Thermus aquaticus, which naturally occurs in hot (50 to 80 °C (120 to 175 °F)) environments[10] paved the way for dramatic improvements of the PCR method. The DNA polymerase isolated from T. aquaticus is stable at high temperatures remaining active even after DNA denaturation,[11] thus obviating the need to add new DNA polymerase after each cycle[3]. This allowed an automated thermocycler-based process for DNA amplification.

At the time he developed PCR in 1983, Mullis was working in Emeryville, California for Cetus Corporation, one of the first biotechnology companies. There, he was responsible for synthesizing short chains of DNA. Mullis has written that he conceived of PCR while cruising along the Pacific Coast Highway one night in his car.[38] He was playing in his mind with a new way of analyzing changes (mutations) in DNA when he realized that he had instead invented a method of amplifying any DNA region through repeated cycles of duplication driven by DNA polymerase.

In Scientific American, Mullis summarized the procedure: "Beginning with a single molecule of the genetic material DNA, the PCR can generate 100 billion similar molecules in an afternoon. The reaction is easy to execute. It requires no more than a test tube, a few simple reagents, and a source of heat."[39] He was awarded the Nobel Prize in Chemistry in 1993 for his invention,[4] seven years after he and his colleagues at Cetus first put his proposal to practice. However, some controversies have remained about the intellectual and practical contributions of other scientists to Mullis' work, and whether he had been the sole inventor of the PCR principle. (see main article: Kary Mullis)

[edit] Patent wars

The PCR technique was patented by Cetus Corporation, where Mullis worked when he invented the technique in 1983. The Taq polymerase enzyme was also covered by patents. There have been several high-profile lawsuits related to the technique, including an unsuccessful lawsuit brought by DuPont. The pharmaceutical company Hoffmann-La Roche purchased the rights to the patents in 1992 and currently holds those that are still protected.

A related patent battle over the Taq polymerase enzyme is still ongoing in several jurisdictions around the world between Roche and Promega. The legal arguments have extended beyond the life of the original PCR and Taq polymerase patents, which expired on March 28, 2005 [40]

Kary Banks Mullis, Ph.D. (born December 28, 1944) is an American biochemist and Nobel laureate.

Dr Mullis was awarded the Nobel Prize in Chemistry in 1993 for his development of the Polymerase Chain Reaction (PCR), a central technique in biochemistry and molecular biology which allows the amplification of specific DNA sequences. Dr Mullis subsequently was awarded the Japan Prize that same year.

Contents

[hide] 1 Early life and education 2 Career

o 2.1 PCR and other inventions o 2.2 Accreditation of the PCR technique

3 Controversies o 3.1 HIV link to AIDS o 3.2 Global warming

4 Personal life 5 Use of LSD 6 Books authored

o 6.1 Awards and honors 7 See also 8 References 9 Additional sources

10 External links

[edit] Early life and education

Mullis was born in Lenoir, North Carolina, near the Blue Ridge Mountains,[1] on December 28, 1944. His family had a background in farming in the rural area, and as a child Mullis studied the diverse organisms of nearby farms.[2] He grew up in Columbia, South Carolina,[2] where he attended Dreher High School.

Mullis earned a Bachelor of Science degree in chemistry [1] from the Georgia Institute of Technology in Atlanta in 1966 and received a Ph.D. in biochemistry from the University of California, Berkeley in 1973; his research focused on synthesis and structure of proteins.[2] Following his graduation, Mullis became a postdoctoral fellow in paediatric cardiology at the University of Kansas Medical School, going on to complete two years of postdoctoral work in pharmaceutical chemistry at the University of California, San Francisco.

[edit] Career

In 1979, Mullis joined the biotechnology company Cetus Corporation of Emeryville, California,[2] where he worked as a DNA chemist for seven years. In 1983, while synthesizing oligonucleotides, Mullis invented the technique known as the polymerase chain reaction.[3] He then proceeded to Xytronyx Inc. in 1986, where he was appointed the director of molecular biology, before moving on to serve as a nucleic acid chemistry consultant for multiple corporations.[4]

In 1992, Mullis founded a business with the intent to sell pieces of jewelry containing the amplified DNA of famous people, such as musicians, to young people.[5]

[edit] PCR and other inventions

Main articles: Taq Polymerase and History of polymerase chain reaction

In 1983, Mullis was working for Cetus. That spring, while driving his scooter and not watching the lines on the highway,[6] Mullis conceived of the idea of using a pair of primers to bracket the desired sequence and copying it using DNA polymerase, but the polymerase was destroyed with each thermal cycle and had to be replaced. In 1986, he started to use Thermophilus aquaticus (Taq) DNA polymerase to amplify segments of DNA. The Taq polymerase was heat resistant and would only need to be added once, thus making the technique dramatically more affordable and subject to automation. This has created revolutions in biochemistry, molecular biology, genetics, medicine and forensics.

Mullis has also invented a UV-sensitive plastic that changes color in response to light, and most recently has been working on an approach for mobilizing the immune system to neutralize invading pathogens and toxins, leading to the formation of his current venture, Altermune LLC. This work is now being funded by DARPA. Mullis described this idea this way:

It is a method using specific synthetic chemical linkers to divert an immune response from its nominal target to something completely different which you would right now like to be temporarily immune to. Let's say you just got exposed to a new strain of the flu. You're already immune to alpha-1,3-galactosyl-galactose bonds. All humans are. Why not divert a fraction of those antibodies to the influenza strain you just picked up? A chemical linker synthesized with an alpha-1,3-gal-gal bond on one end and a DNA aptamer devised to bind specifically to the strain of influenza you have on the other end will link anti-alpha-Gal antibodies to the influenza virus and presto!--you have fooled your immune system into attacking the new virus.[1]

[edit] Accreditation of the PCR technique

Some controversy surrounds the balance of credit that should be given to Mullis versus the team at Cetus.[citation needed] In practice, credit has accrued to both the inventor and the company (although not its individual workers) in the form of a Nobel Prize and a $10,000 Cetus bonus for Mullis and $300 million for Cetus when the company sold the patent to Roche Molecular Systems.

The main principles of PCR were described in 1971 by Kjell Kleppe, a Norwegian scientist, and some have asserted that Kleppe has a better claim to the invention.[citation needed] Together with 1968 Nobel Prize laureate H. Gobind Khorana, Kleppe released a 20-page research paper on PCR in the 1971 Journal of Molecular Biology. As early as June 18, 1969, Kleppe presented his work at the Gordon Conference in New Hampshire. Using repair replication (the principle of PCR), he duplicated and then quadrupled a small synthetic molecule with the help of two primers and DNA-polymerase. Among the attendees[7] was Stuart Linn, who then used Kleppe's material in his own teachings to his students, including Mullis.

The suggestion that Mullis was solely responsible for the idea of using Taq polymerase in the PCR process has been refuted by his co-workers at the time.[citation needed] However, other scientists have said that "the full potential [of PCR] was not realized" until Mullis' work in 1983,[8] and at least one book has reported that Mullis' colleagues failed to see the potential of the technique when he presented it to them.[6]

The anthropologist Paul Rabinow wrote a book on the history of the PCR method in 1996 in which he questioned whether or not Mullis "invented" PCR or "merely" came up with the concept of it. Rabinow, a Foucault scholar interested in issues of the production of knowledge, used the topic to argue against the idea that scientific discovery is the product of individual work, writing, "Committees and science journalists like the idea of associating a unique idea with a unique person, the lone genius. PCR is, in fact, one of the classic examples of teamwork."[9]

[edit] Controversies

[edit] HIV link to AIDS

Mullis has also drawn controversy[citation needed] for his past association with Peter Duesberg and his controversial position on HIV and its link to AIDS.[10] As the recipient of a Nobel Prize for the PCR technique that is used to measure viral load in people with AIDS. Mullis wrote in an introduction to Duesberg's Inventing the Aids Virus (1997), "No one has ever proven that HIV causes AIDS. We have not been able to discover any good reasons why most of the people on earth believe that AIDS is a disease caused by a virus called HIV."[11]

Mullis has said of HIV:

"If HIV has been here all along and it can be passed from mother to child, wouldn't it make sense to test for the antibodies in the mothers of anyone who is positive to HIV, especially if that individual is not showing any signs of disease?... If an HIV-positive woman develops uterine cancer, for example, she is considered to have AIDS. If she is not HIV-positive, she simply has uterine cancer. An HIV-positive man with tuberculosis has AIDS; if he tests negative he simply has tuberculosis. If he lives in Kenya or Colombia, where the test for HIV antibodies is too expensive, he is simply presumed to have the antibodies and therefore AIDS, and therefore he can be treated in the World Health Organization's clinic. It's the only medical help available in some places."[12]

[edit] Global warming

Mullis is skeptical about the concern over global warming, disagreeing with the theory that humans are a factor and also disagrees with the idea that CFCs cause ozone depletion.[13]

[edit] Personal life

Mullis enjoys beach surfing.[14]

[edit] Use of LSD

In a Q&A interview published in the September 1994 issue of California Monthly, Mullis said, "Back in the 1960s and early '70s I took plenty of LSD. A lot of people were doing that in Berkeley back then. And I found it to be a mind-opening experience. It was certainly much more

important than any courses I ever took." [15] During a symposium held for centenarian Albert Hofmann, Hofmann revealed that he was told by Nobel-prize-winning chemist Kary Mullis that LSD had helped him develop the polymerase chain reaction that helps amplify specific DNA sequences.[16]

[edit] Books authored

The Polymerase Chain Reaction, 1994, with Richard A. Gibbs Dancing Naked in the Mind Field. 1998, Vintage Books.

Mullis wrote the 1998 autobiography Dancing Naked in the Mind Field, which gives an account of his initial invention of PCR, as well as providing insights into the opinions and experiences of the author. Several examples of supposedly atypical behavior for a scientist, including the use of LSD, belief in astrology, and the belief in an extraterrestrial encounter, are also chronicled within the book.

[edit] Awards and honors

1990 - William Allan Memorial Award of the American Society of Human Genetics | Preis Biochemische Analytik of the German Society of Clinical Chemistry and Boehringer Mannheim

1991 - National Biotechnology Award | Gairdner Award | R&D Scientist of the Year 1992 - California Scientist of the Year Award 1993 - Nobel Prize in Chemistry | Japan Prize | Thomas A. Edison Award 1994 - Honorary degree of Doctor of Science from the University of South Carolina 1998 - Inducted into the National Inventors Hall of Fame [17] | 2004 - Honorary degree in Pharmaceutical Biotechnology from the University of

Bologna, Italy

Ronald H. Brown American Innovator Award[18]

Mullis has also received the John Scott Award, given by the City Trusts of Philadelphia to other Nobelists, as well as Thomas Edison and the Wright Brothers.[19]

The polymerase chain reaction (PCR) is a commonly used molecular biology tool for amplifying DNA, and various techniques for PCR optimization have been developed by molecular biologists to improve PCR performance and minimize failure.

Contents

[hide] 1 Contamination and PCR 2 Hairpins 3 Polymerase errors 4 Size and other limitations

5 Non-specific priming

6 References

[edit] Contamination and PCR

The PCR method is extremely sensitive, requiring only a few DNA molecules in a single reaction for amplification across several orders of magnitude. Therefore, adequate measures to avoid contamination from any DNA present in the lab environment (bacteria, viruses, or human sources) are required. Because products from previous PCR amplifications are a common source of contamination, many molecular biology labs have implemented procedures that involve dividing the lab into separate areas.[1] One lab area is dedicated to preparation and handling of pre-PCR reagents and the setup of the PCR reaction, and another area to post-PCR processing, such as gel electrophoresis or PCR product purification. For the setup of PCR reactions, many standard operating procedures involve using pipettes with filter tips and wearing fresh laboratory gloves, and in some cases a laminar flow cabinet with UV lamp as a work station (to destroy any extraneous DNA before PCR setup). Possible contamination with extraneous DNA or primer-multimer formation is routinely assessed with a (negative) control PCR reaction. This control reaction is set up in the same way as the experimental PCRs, but without template DNA added, and is performed alongside the experimental PCRs.

[edit] Hairpins

Secondary structures in the DNA can result in folding or knotting of DNA template or primers, leading to decreased product yield or failure of the reaction. Hairpins, which consist of internal folds caused by base-pairing between nucleotides in inverted repeats within single-stranded DNA, are common secondary structures and may result in failed PCRs.

Typically, primer design that includes a check for potential secondary structures in the primers, or addition of DMSO or glycerol to the PCR to minimize secondary structures in the DNA template[citation needed], are used in the optimization of PCRs that have a history of failure due to suspected DNA hairpins.

[edit] Polymerase errors

Taq polymerase lacks a 3' to 5' exonuclease activity. Thus, Taq has no error-proofreading activity, which consists of excision of any newly polymerized nucleotide base from the nascent (=extending) DNA strand that does not match with its opposite base in the complementary DNA strand. The lack in 3' to 5' proofreading of the Taq enzyme results in a high error rate (approximately 1 in 10,000 bases), which affects the fidelity of the PCR, especially if errors occur early in the PCR, causing accumulation of a large proportion of amplified DNA with incorrect sequence in the final product.

Several "high-fidelity" DNA polymerases, having engineered 3' to 5' exonuclease activity, have become available that permit more accurate amplification for use in PCRs for sequencing or

cloning of products. Examples of polymerases with 3' to 5' exonuclease activity include: KOD DNA polymerase, a recombinant form of Thermococcus kodakaraensis KOD1; Vent, which is extracted from Thermococcus litoralis; Pfu DNA polymerase, which is extracted from Pyrococcus furiosus; and Pwo, which is extracted from Pyrococcus woesii.[citation needed]

[edit] Size and other limitations

PCR works readily with a DNA template of up to two to three thousand base pairs in length. However, above this size, product yields often decrease, as with increasing length stochastic effects such as premature termination by the polymerase begin to affect the efficiency of the PCR. It is possible to amplify larger pieces of up to 50,000 base pairs with a slower heating cycle and special polymerases. These are polymerases fused to a processivity-enhancing DNA-binding protein, enhancing adherence of the polymerase to the DNA [2][3].

Other valuable properties of the chimeric polymerases TopoTaq and PfuC2 include enhanced thermostability, specificity and resistance to contaminants and inhibitors [4] [5] . They were engineered using the unique helix-hairpin-helix (HhH) DNA binding domains of topoisomerase V[6] from hyperthermophile Methanopyrus kandleri. Chimeric polymerases overcome many limitations of native enzymes and are used in direct PCR amplification from cell cultures and even food samples, thus by-passing laborious DNA isolation steps. A robust strand-displacement activity of the hybrid TopoTaq polymerase helps solving PCR problems with hairpins and G-loaded double helices, because helices with a high G-C context possess a higher melting temperature [7].

[edit] Non-specific priming

Non-specific binding of primers frequently occurs and can be due to repeat sequences in the DNA template, non-specific binding between primer and template, and incomplete primer binding, leaving the 5' end of the primer unattached to the template. Non-specific binding is also often increased when degenerate primers are used in the PCR. Manipulation of annealing temperature and magnesium ion (which stabilise DNA and RNA interactions) concentrations can increase specificity. Non-specific priming during reaction preparation at lower temperatures can be prevented by using "hot-start" polymerase enzymes whose active site is blocked by an antibody or chemical that only dislodges once the reaction is heated to 95˚C during the denaturation step of the first cycle.

Other methods to increase specificity include Nested PCR and Touchdown PCR.

Factors Affecting the PCR:

Denaturing Temperature and time

The specific complementary association due to hydrogen bonding of single-stranded nucleic acids is referred to as "annealing": two complementary sequences will form hydrogen bonds between their complementary bases (G to C, and A to T or U) and form a stable double-

stranded, anti-parallel "hybrid" molecule. One may make nucleic acid (NA) single-stranded for the purpose of annealing - if it is not single-stranded already, like most RNA viruses - by heating it to a point above the "melting temperature" of the double- or partially-double-stranded form, and then flash-cooling it: this ensures the "denatured" or separated strands do not re-anneal. Additionally, if the NA is heated in buffers of ionic strength lower than 150mM NaCl, the melting temperature is generally less than 100oC - which is why PCR works with denaturing temperatures of 91-97oC.

A more detailed treatment of annealing / hybridisation is given in an accompanying page, together with explanations of calculations of complexity, conditions for annealing / hybridisation, etc.

Taq polymerase is given as having a half-life of 30 min at 95oC, which is partly why one should not do more than about 30 amplification cycles: however, it is possible to reduce the denaturation temperature after about 10 rounds of amplification, as the mean length of target DNA is decreased: for templates of 300bp or less, denaturation temperature may be reduced to as low as 88oC for 50% (G+C) templates (Yap and McGee, 1991), which means one may do as many as 40 cycles without much decrease in enzyme efficiency.

"Time at temperature" is the main reason for denaturation / loss of activity of Taq: thus, if one reduces this, one will increase the number of cycles that are possible, whether the temperature is reduced or not. Normally the denaturation time is 1 min at 94oC: it is possible, for short template sequences, to reduce this to 30 sec or less. Increase in denaturation temperature and decrease in time may also work: Innis and Gelfand (1990) recommend 96oC for 15 sec.

Annealing Temperature and Primer Design

Primer length and sequence are of critical importance in designing the parameters of a successful amplification: the melting temperature of a NA duplex increases both with its length, and with increasing (G+C) content: a simple formula for calculation of the Tm is

Tm = 4(G + C) + 2(A + T)oC.

Thus, the annealing temperature chosen for a PCR depends directly on length and composition of the primer(s). One should aim at using an annealing temperature (Ta) about 5oC below the lowest Tm of ther pair of primers to be used (Innis and Gelfand, 1990). A more rigorous treatment of Ta is given by Rychlik et al. (1990): they maintain that if the Ta is increased by 1oC every other cycle, specificity of amplification and yield of products <1kb in length are both increased. One consequence of having too low a Ta is that one or both primers will anneal to sequences other than the true target, as internal single-base mismatches or partial annealing may be tolerated: this is fine if one wishes to amplify similar or related targets; however, it can lead to "non-specific" amplification and consequent reduction in yield of the desired product, if the 3'-most base is paired with a target. 

A consequence of too high a Ta is that too little product will be made, as the likelihood of primer annealing is reduced; another and important consideration is that a pair of primers with very different Tas may never give appreciable yields of a unique product, and may also result in inadvertent "asymmetric" or single-strand amplification of the most efficiently primed product strand.

Annealing does not take long: most primers will anneal efficiently in 30 sec or less, unless the Ta is too close to the Tm, or unless they are unusually long.

An illustration of the effect of annealing temperature on the specificity and on the yield of amplification of Human papillomavirus type 16 (HPV-16) is given below (Williamson and Rybicki, 1991: J Med Virol 33: 165-171).

Plasmid and biopsy sample DNA templates were amplified at different annealing temperatures as shown: note that while plasmid is amplified from 37 to 55oC, HPV DNA is only specifically

amplified at 50oC.

Primer Length

The optimum length of a primer depends upon its (A+T) content, and the Tm of its partner if one runs the risk of having problems such as described above. Apart from the Tm, a prime consideration is that the primers should be complex enough so that the likelihood of annealing to sequences other than the chosen target is very low. (See hybridn.doc).

For example, there is a ¼ chance (4-1) of finding an A, G, C or T in any given DNA sequence; there is a 1/16 chance (4-2) of finding any dinucleotide sequence (eg. AG); a 1/256 chance of finding a given 4-base sequence. Thus, a sixteen base sequence will statistically be present only once in every 416 bases (=4 294 967 296, or 4 billion): this is about the size of the human

or maize genome, and 1000x greater than the genome size of E. coli. Thus, the association of a greater-than-17-base oligonucleotide with its target sequence is an extremely sequence-specific process, far more so than the specificity of monoclonal antibodies in binding to specific antigenic determinants. Consequently, 17-mer or longer primers are routinely used for amplification from genomic DNA of animals and plants. Too long a primer length may mean that even high annealing temperatures are not enough to prevent mismatch pairing and non-specific priming.

Degenerate Primers

For amplification of cognate sequences from different organisms, or for "evolutionary PCR", one may increase the chances of getting product by designing "degenerate" primers: these would in fact be a set of primers which have a number of options at several positions in the sequence so as to allow annealing to and amplification of a variety of related sequences. For example, Compton (1990) describes using 14-mer primer sets with 4 and 5 degeneracies as forward and reverse primers, respectively, for the amplification of glycoprotein B (gB) from related herpesviruses. The reverse primer sequence was as follows:

TCGAATTCNCCYAAYTGNCCNT

where Y = T + C, and N = A + G + C + T, and the 8-base 5'-terminal extension comprises a EcoRI site (underlined) and flanking spacer to ensure the restriction enzyme can cut the product (the New England Biolabs catalogue gives a good list of which enzymes require how long a flanking sequence in order to cut stub ends). Degeneracies obviously reduce the specificity of the primer(s), meaning mismatch opportunities are greater, and background noise increases; also, increased degeneracy means concentration of the individual primers decreases; thus, greater than 512-fold degeneracy should be avoided. However, I have used primers with as high as 256- and 1024-fold degeneracy for the successful amplification and subsequent direct sequencing of a wide range of Mastreviruses against a background of maize genomic DNA (Rybicki and Hughes, 1990).

Primer sequences were derived from multiple sequence alignments; the mismatch positions were used as 4-base degeneracies for the primers (shown as stars; 5 in F and 4 in R), as shown above.  Despite their degeneracy, the primers could be used to amplify a 250 bp sequence from viruses differing in sequence by as much as 50% over the target sequence, and 60% overall.  They could also be used to very sensitively detect the presence of Maize streak virus DNA against a background of maize genomic DNA, at dilutions as low as 1/109 infected sap / healthy sap (see below).

The identification of novel members of gene families by PCR using degenerate primers has been considered more of an art than a science, so much so that the methods books I've come across have been too timid to discuss the considerations that go into the design of this experiment, much less give a protocol for its execution. At the risk of leading my readers on wild goose chases, I'm committing my methods to paper. The following is based on my reading of the recent literature (e.g. Buck and Axel, Cell 65: 175-187, 1991; Riddle et al., Cell 75: 1401-1416, 1993; Krauss et al., Cell 75:1431-1444, 1993), discussions with several other successful practitioners of the art, and my own experience isolating vertebrate homologs of the C. elegans egl-10 gene (Koelle and Horvitz, Cell 84: 1-20, 1996).

Primer design This is the most important factor in the success of the experiment, and deserves careful deliberation. I suggest diagramming out an alignment of the existing members of your gene family, highlighting conserved residues, and labeling each important position in the alignment with the number of codons that encode the amino acid(s) at that position. An example based on the original members of the egl-10 gene family is included below, and cited in the following discussion.

Primer degeneracy

In the early days of degenerate PCR some novel genes were successfully amplified using primer pools that were over 1000-fold degenerate. However, primers of such high degeneracy appear not to have been generally successful (with some exceptions, e.g. Giovane et al., Genes Dev. 8: 1502-1513, 1994), and most recent successes have come using primer pools of 100-fold degeneracy or less. Five methods for reducing the degeneracy of the primers are discussed

below: 1) judicious selection of the primer sites The positioning of the primers is a compromise between placing them at the codons for the most conserved amino acids, and placing them at the codons for less conserved amino acids whose degeneracy may be lower. Consider the case of the 3' primers ("3T and 3A") in the example shown below for the egl-10 gene family. At first it might seem more sensible to place these primers 3 codons to the right, where there is a block of 5 out of 6 absolutely conserved amino acids:

SY(P/Q)

RFL. Unfortunately, this block of amino acids is encoded by 5184 different DNA sequences. The actual primers used were placed 3 codons to the left. At this position only 3 out of 6 amino acids are absolutely conserved:

M(E/K)(K/N)(D/N)

SY. However, this block of amino acids is encoded by only 768 different DNA sequences.2) the use of inosine as a "neutral" base Inosine is a purine (which occurs naturally in tRNAs) that can form base pairs with cytidine, thymidine, and adenosine (although the inosine:adenosine pairing presumably doesn't fit quite correctly in double stranded DNA, so there may be an energetic penalty to pay when the helix bulges out at this purine:purine pairing). Recently, most people have been using inosine in their primers at positions where any of the four bases might be required. Each use of inosine thus reduces the degeneracy of the primer pool 4-fold. However, you risk the occurrence of I:G mismatches, and therefore must assume that exact base pairing at other positions in the primer will overcome such a problem. Most oligo synthesis facilities will make inosine-containing oligos, no problem. I had excellent luck with inosine-containing primers with the egl-10 gene family, except in the case of the primer "5out", a 20mer containing 5 inosines, (including 2 near the 3' end of the primer) which failed to amplify products even from a cloned egl-10 cDNA. So, perhaps 5 out of 20 inosines is too many.Using inosine in the primers requires that the DNA polymerase used in the PCR reaction be capable of synthesizing DNA over an inosine-containing template. Taq polymerase is capable of doing this, but some others (e.g. Vent) appear not to be able to.3) using multiple separate oligo pools at a single position In an effort to use primer pools with the lowest possible degeneracy, it is sometimes useful to synthesize primers over a particular stretch of codons as two or more separate pools, each of which will have lower degeneracy than you would get by synthesizing a single pool including all of the same codons. The pools are then used separately to carry out PCR reactions. For example, the primer pools "3T" and "3A" in the egl-10 example below are identical, except at their serine codons. Sadly, serine is encoded by 6 different codons, TC(A/G/C/T) and AG(T/C). Synthesizing a single pool covering all these possibilities might require a high degeneracy and

would necessarily include some non-serine codons. By splitting into two pools (one nondegenerate containing an inosine, the other 2-fold degenerate) I was able to keep the degeneracy low, and avoid all non-serine codons. Another example is shown in the case of primers "5inE" and "5inR", which again are identical except at one codon. 4) including partial codons at the ends of the primers The various codons encoding an amino acid or a set of similar amino acids are often identical at their first (and maybe second) positions, but different at their third position. You can take advantage of this by synthesizing only the first or first and second positions of the 3' most codon covered by your primer pools, thus giving you one or two extra positions of exact match base pairs without adding any degeneracy. In the egl-10 example, the primer pools "3T" and "3A" cover a stretch in which the last codon must encode proline or glutamine. The codons for these two amino acids all start with C, but their last two positions are degenerate. Therefore, only the nondegenerate C was included in the primer pools.5) use of codon bias Some organisms have strong biases for using particular codons to encode certain amino acids. In theory you could reduce the degeneracy of a primer pool by only including these most common codons, and taking the risk that the gene(s) you are looking for will follow the organism's general codon bias enough to allow such primer pools to work. I haven't heard of anyone actually using this codon bias strategy in a successful degenerate PCR experiment, but you might try it if you're desperate.

Other considerations in primer design

1) primer lengthIn the example below the short stretches of sequence similarity among the egl-10 family members forced me to use primers only 19-21 bases long. These are shorter than the primers I have heard of people using in other successful experiments. For example, Linda Buck's primers were 31-33mers.2) 3' endPeople I talked to emphasized the special importance of having an exact match between the primer and template near the 3' end of the primer, although I'm not aware of specific data supporting this idea. For egl-10 I tried to avoid having any inosines near the 3' ends of the primers (except for primer "5out", which in fact failed to give any products), and also anchored the primers when possible with a nondegenerate codon at their 3' ends, so that 100% of the primers in the pool would be able to pair perfectly with the correct template over these last few bases.3) nested primersIf the sequence similarity in your gene family permits, it is a good idea to make nested sets of PCR primers. That way one round of PCR can be performed using the outside primers, and individual products (or the whole mix) can then be reamplified using the inside primers. Products amplified through both rounds are more likely to be the desired new gene family members, and less likely to be spurious products from sequences that happen to contain a couple of primer annealing sites by chance.

Determining optimal reaction conditions A number of parameters can be varied to optimize reaction conditions for degenerate PCR.

These include: primer concentration, magnesium concentration, template concentration, number of cycles of amplification, and the temperatures and times of each step in the amplification cycle. If each of these parameters is to be independently varied, the number of possibilities quickly reaches mind boggling proportions. My philosophy has been to fix almost all these parameters at the standard levels that have been successful for other people, and to vary only the one parameter that I think is the most crucial: the temperature of the annealing step during amplification.My standard PCR reactions are as follows:

1. 5 l template DNA (2-300 ng)5 l 5 l 10X PCR buffer (10X buffer=100 mM Tris pH 8.3, 500 mM KCl, 15 mM MgCl2, 0.01% gelatin)8 l dNTP mix (1.25 mM each dNTP)

2. 2 l "ampliTaq" polymerase (5 U/ l)25 l dH205 l each primer pool at 20 M eachtotal volume 50 l

In practice, the reactions are set up by placing the primers and template into a 0.5 ml tube, then adding two drops of mineral oil from a blue tip, and adding on top of the oil 38.5 l of a premix containing all the other components. In this way, it is easy to set up many different primer/template combinations at once. The tubes are then briefly spun in a microfuge to combine the two aqueous phases, and the tubes are immediately placed in the PCR block preheated to 95 for a "hot start".My amplification program:

95 X 3 min. (hot start)

?? X 1 min. (this annealing temperature is varied to optimize the amplification)72 X 2 min.94 X 45 sec.40 cycles of the above 3 steps

72 X 5 min.hold at 4

This takes ~4.5 hours to run on an MJ Research machine.

To test the primers and optimize the conditions, I do a series of amplification runs starting with an annealing temperature of 25 , and increasing in 5 increments until amplification fails to occur. Typically for each primer pair being tested, at each temperature, I run 3 reactions containing different templates: 1) a positive control containing 2 ng of a cloned member of the gene family of interest as template. 2) a negative control containing no template (this is very important-- you don't want to get fooled by contaminants). 3) an experimental reaction containing a complex template such as genomic DNA or total cDNA. For total C. elegans genomic DNA I've been using 300 ng as a template. Using rat brain cDNA as a template I amplified off of only 2 ng. However, this was only because I didn't have very much cDNA. If possible, it would be better to use ~200 ng of cDNA as a template, as Linda Buck did to amplify the odorant receptors.

Choice of template Genomic DNA has the advantage that all members of your gene family are present in equimolar amounts, and genomic DNA is probably readily available. The obvious disadvantage is that introns may disrupt the primer sites, or may cause the amplification product to be so long that it is not amplified efficiently. cDNA templates, though harder to obtain, overcome this problem. A big advantage of cDNA is that the desired amplification products should be of a known size, and you can therefore easily pick them out from among spurious products of other sizes. Remember that the "correct" sized band amplified off a cDNA template may be a complex mixture of products from many gene family members, so you may have to analyze many clones generated from such a band to assess its complexity. Linda Buck used random primed cDNA for her template, presumably to avoid biasing the cDNA towards the 3' ends of transcripts. In my case, I knew that the region I was amplifying should be at the extreme 3' end of the coding sequence, so I used oligo-dT primed cDNA.For the lazy and rich, Clontech sells oligo-dT primed cDNA prepared from various tissues of many species to use as templates for PCR.

Analysis of PCR products After amplification run 20 l of each reaction out on an agarose gel. I use 2% 3:1 Nusieve:SeaKem LE agarose (you can buy this premixed from FMC) in 1X TAE. This gel is not very low melting, and thus isn't very suitable for cloning directly from the gel, but it gives very nice resolution. I use the 123 bp ladder from Gibco as a size standard. Obviously you expect to get products off the positive control, and not to get them off the negative control. Using the complex template, you will probably get a smear at the lower annealing temperatures, which will resolve into a small number of bands as the annealing temperature rises. I pick an annealing temperature that gives a modest number of bands, and then clone all these bands and sequence them.If no products are evident in the experimental samples, a good trick to try is to use 2 l of the apparently failed reaction as a template, and reamplify under the same conditions. This often gives visible products.If you want to clone products that are only barely visible, you can get more of them by just reamplifying the original reaction as described above. Another way to amplify individual products separately is to cut the bands out of the gel that was used to analyze the original reactions (actually I take a bore out of the gel with a Pasteur pipette), melt the DNA containing agarose, and use 2 l of it as a template to reamplify under the same conditions.The above described method for reamplifying specific bands can (and should) be used to test amplified products to see if they are single primer artefacts. Use 2 l of an agarose gel bore to set up each of three PCR reactions, containing either individual primer or both together. Obviously, you're only interested in products that require both primers in order to be amplified.I clone PCR products by running the PCR reaction out on a low melt agarose gel (2% Nusieve agarose in 1X TAE). I cut the desired band out, melt it at 70 , mix well by pipetting up/down, and use 5 l of the melted agarose directly in a ligation reaction with a dT tailed vector. This vector DNA is prepared as follows: cut 1 g bluescript SK with EcoRV in a 20 l reaction. Add 20 l 1X PCR buffer, and 2 l 2 mM dTTP. Add 0.5 l "ampliTaq" polymerase (2.45 U), and incubate at ~72 for 20 min. Run the DNA out on a 0.8% Seaplaque agarose gel in 1X TAE, cut

out the band, melt at 70 , mix well, and use 5 l in a ligation reaction. It turns out that only about 50% of the colonies obtained after transformation of this type of reaction may have inserts; the rest are vector reclosures. However, if blue/white selection is used, virtually all the white colonies have inserts.

Some groups use deoxyinosine (dI) at degenerate positions rather than use mixed oligos: this base-pairs with any other base, effectively giving a four-fold degeneracy at any postion in the oligo where it is present. This lessens problems to do with depletion of specific single oligos in a highly degenerate mixture, but may result in too high a degeneracy where there are 4 or more dIs in an oligo.

Elongation Temperature and Time

This is normally 70 - 72oC, for 0.5 - 3 min. Taq actually has a specific activity at 37oC which is very close to that of the Klenow fragment of E coli DNA polymerase I, which accounts for the apparent paradox which results when one tries to understand how primers which anneal at an optimum temperature can then be elongated at a considerably higher temperature - the answer is that elongation occurs from the moment of annealing, even if this is transient, which results in considerably greater stability. At around 70oC the activity is optimal, and primer extension occurs at up to 100 bases/sec. About 1 min is sufficient for reliable amplification of 2kb sequences (Innis and Gelfand, 1990). Longer products require longer times: 3 min is a good bet for 3kb and longer products. Longer times may also be helpful in later cycles when product concentration exceeds enzyme concentration (>1nM), and when dNTP and / or primer depletion may become limiting.

Reaction Buffer

Recommended buffers generally contain :

10-50mM Tris-HCl pH 8.3, up to 50mM KCl, 1.5mM or higher MgCl2, primers 0.2 - 1uM each primer, 50 - 200uM each dNTP, gelatin or BSA to 100ug/ml, and/or non-ionic detergents such as Tween-20 or Nonidet P-40 or Triton X-100 (0.05

- 0.10% v/v)

(Innis and Gelfand, 1990).  Modern formulations may differ considerably, however - they are also generally proprietary.

PCR is supposed to work well in reverse transcriptase buffer, and vice-versa, meaning 1-tube protocols (with cDNA synthesis and subsequent PCR) are possible (Krawetz et al., 19xx; Fuqua et al., 1990).

Higher than 50mM KCl or NaCl inhibits Taq, but some is necessary to facilitate primer annealing.

[Mg2+] affects primer annealing; Tm of template, product and primer-template associations; product specificity; enzyme activity and fidelity. Taq requires free Mg2+, so allowances should be made for dNTPs, primers and template, all of which chelate and sequester the cation; of these, dNTPs are the most concentrated, so [Mg2+] should be 0.5 - 2.5mM greater than [dNTP]. A titration should be performed with varying [Mg2+] with all new template-primer combinations, as these can differ markedly in their requirements, even under the same conditions of concentrations and cycling times/temperatures.

Some enzymes do not need added protein, others are dependent on it. Some enzymes work markedly better in the presence of detergent, probably because it prevents the natural tendency of the enzyme to aggregate.

Primer concentrations should not go above 1uM unless there is a high degree of degeneracy; 0.2uM is sufficient for homologous primers.

Nucleotide concentration need not be above 50uM each: long products may require more, however.

Cycle Number

The number of amplification cycles necessary to produce a band visible on a gel depends largely on the starting concentration of the target DNA: Innis and Gelfand (1990) recommend from 40 - 45 cycles to amplify 50 target molecules, and 25 - 30 to amplify 3x105 molecules to the same concentration. This non-proportionality is due to a so-called plateau

effect, which is the attenuation in the exponential rate of product accumulation in late stages of a PCR, when product reaches 0.3 - 1.0 nM. This may be caused by degradation of reactants (dNTPs, enzyme); reactant depletion (primers, dNTPs - former a problem with short products, latter for long products); end-product inhibition (pyrophosphate formation); competition for reactants by non-specific products; competition for primer binding by re-annealing of concentrated (10nM) product (Innis and Gelfand, 1990).

If desired product is not made in 30 cycles, take a small sample (1ul) of the amplified mix and re-amplify 20-30x in a new reaction mix rather than extending the run to more cycles: in some cases where template concentration is limiting, this can give good product where extension of cycling to 40x or more does not.

A variant of this is nested primer PCR: PCR amplification is performed with one set of primers, then some product is taken - with or without removal of reagents - for re-amplification with an internally-situated, "nested" set of primers.  This process adds another level of specificity, meaning that all products non-specifically amplified in the first round will not be amplified in the second.  This is illustrated below:

This gel photo shows the effect of nested PCR amplification on the detectability of Chicken anaemia virus (CAV) DNA in a dilution series: the PCR1 just detects 1000 template molecules; PCR2 amplifies 1 template molecule (Soiné C, Watson SK, Rybicki EP, Lucio B, Nordgren RM, Parrish CR, Schat KA (1993)  Avian Dis 37: 467-476).

Labelling of PCR products with digoxygenin-11-dUTP

(DIG; Roche) need be done only in 50uM each dNTP, with the dTTP substituted to 35% with DIG-11-dUTP. NOTE: that the product will have a higher MW than the native product!  This results in a very well labelled probe which can be extensively re-used, for periods up to 3 years.  See also here.

Helix Destabilisers / Additives

With NAs of high (G+C) content, it may be necessary to use harsher denaturation conditions. For example, one may incorporate up to 10% (w or v/v) :

dimethyl sulphoxide (DMSO), dimethyl formamide (DMF), urea or formamide 

in the reaction mix: these additives are presumed to lower the Tm of the target NA, although DMSO at 10% and higher is known to decrease the activity of Taq by up to 50% (Innis and Gelfand, 1990; Gelfand and White, 1990).

Additives may also be necessary in the amplification of long target sequences: DMSO often helps in amplifying products of >1kb. Formamide can apparently dramatically improve the specificity of PCR (Sarkar et al., 1990), while glycerol improves the amplification of high (G+C) templates (Smith et al., 1990). 

Polyethylene glycol (PEG) may be a useful additive when DNA template concentration is very low: it promotes macromolecular association by solvent exclusion, meaning the pol can find the DNA.  

cDNA PCR

A very useful primer for cDNA synthesis and cDNA PCR comes from a sequencing strategy described by Thweatt et al. (1990): this utilised a mixture of three 21-mer primers consisting of 20 T residues with 3'-terminal A, G or C, respectively, to sequence inside the poly(A) region of cDNA clones of mRNA from eukaryotic origin. I have used it to amplify discrete bands from a variety of poly(A)+ virus RNAs, with only a single specific degenerate primer upstream: the T-primer may anneal anywhere in the poly(A) region, but only molecules which anneal at the beginning of the poly(A) tail, and whose 3'-most base is complementary to the base next to the beginning of the tail, will be extended.

eg: 5'-TTTTTTTTTTTTTTTTTTTTTTTTT(A,G,C)-3'

works for amplification of Potyvirus RNA, and eukaryotic mRNA

A simple set of rules for primer sequence design is as follows (adapted from Innis and Gelfand, 1991):

1. primers should be 17-28 bases in length; 2. base composition should be 50-60% (G+C);

3. primers should end (3') in a G or C, or CG or GC: this prevents "breathing" of ends and increases efficiency of priming;

4. Tms between 55-80oC are preferred;

5. runs of three or more Cs or Gs at the 3'-ends of primers may promote mispriming at G or C-rich sequences (because of stability of annealing), and should be avoided;

6. 3'-ends of primers should not be complementary (ie. base pair), as otherwise primer dimers will be synthesised preferentially to any other product;

7. primer self-complementarity (ability to form 2o structures such as hairpins) should be avoided.

Examples of inter- and intra-primer complementarity which would result in problems

The identification of novel members of gene families by PCR using degenerate primers has been considered more of an art than a science, so much so that the methods books I've come across have been too timid to discuss the considerations that go into the design of this experiment, much less give a protocol for its execution. At the risk of leading my readers on wild goose chases, I'm committing my methods to paper. The following is based on my reading of the recent literature (e.g. Buck and Axel, Cell 65: 175-187, 1991; Riddle et al., Cell 75: 1401-1416, 1993; Krauss et al., Cell 75:1431-1444, 1993), discussions with several other successful practitioners of the art, and my own experience isolating vertebrate homologs of the C. elegans egl-10 gene (Koelle and Horvitz, Cell 84: 1-20, 1996).

Primer design This is the most important factor in the success of the experiment, and deserves careful deliberation. I suggest diagramming out an alignment of the existing members of your gene family, highlighting conserved residues, and labeling each important position in the alignment with the number of codons that encode the amino acid(s) at that position. An example based on the original members of the egl-10 gene family is included below, and cited in the following discussion.

Primer degeneracy

In the early days of degenerate PCR some novel genes were successfully amplified using primer pools that were over 1000-fold degenerate. However, primers of such high degeneracy appear not to have been generally successful (with some exceptions, e.g. Giovane et al., Genes Dev. 8: 1502-1513, 1994), and most recent successes have come using primer pools of 100-fold degeneracy or less. Five methods for reducing the degeneracy of the primers are discussed below: 1) judicious selection of the primer sites The positioning of the primers is a compromise between placing them at the codons for the most

conserved amino acids, and placing them at the codons for less conserved amino acids whose degeneracy may be lower. Consider the case of the 3' primers ("3T and 3A") in the example shown below for the egl-10 gene family. At first it might seem more sensible to place these primers 3 codons to the right, where there is a block of 5 out of 6 absolutely conserved amino acids:

SY(P/Q)

RFL. Unfortunately, this block of amino acids is encoded by 5184 different DNA sequences. The actual primers used were placed 3 codons to the left. At this position only 3 out of 6 amino acids are absolutely conserved:

M(E/K)(K/N)(D/N)

SY. However, this block of amino acids is encoded by only 768 different DNA sequences.2) the use of inosine as a "neutral" base Inosine is a purine (which occurs naturally in tRNAs) that can form base pairs with cytidine, thymidine, and adenosine (although the inosine:adenosine pairing presumably doesn't fit quite correctly in double stranded DNA, so there may be an energetic penalty to pay when the helix bulges out at this purine:purine pairing). Recently, most people have been using inosine in their primers at positions where any of the four bases might be required. Each use of inosine thus reduces the degeneracy of the primer pool 4-fold. However, you risk the occurrence of I:G mismatches, and therefore must assume that exact base pairing at other positions in the primer will overcome such a problem. Most oligo synthesis facilities will make inosine-containing oligos, no problem. I had excellent luck with inosine-containing primers with the egl-10 gene family, except in the case of the primer "5out", a 20mer containing 5 inosines, (including 2 near the 3' end of the primer) which failed to amplify products even from a cloned egl-10 cDNA. So, perhaps 5 out of 20 inosines is too many.Using inosine in the primers requires that the DNA polymerase used in the PCR reaction be capable of synthesizing DNA over an inosine-containing template. Taq polymerase is capable of doing this, but some others (e.g. Vent) appear not to be able to.3) using multiple separate oligo pools at a single position In an effort to use primer pools with the lowest possible degeneracy, it is sometimes useful to synthesize primers over a particular stretch of codons as two or more separate pools, each of which will have lower degeneracy than you would get by synthesizing a single pool including all of the same codons. The pools are then used separately to carry out PCR reactions. For example, the primer pools "3T" and "3A" in the egl-10 example below are identical, except at their serine codons. Sadly, serine is encoded by 6 different codons, TC(A/G/C/T) and AG(T/C). Synthesizing a single pool covering all these possibilities might require a high degeneracy and would necessarily include some non-serine codons. By splitting into two pools (one nondegenerate containing an inosine, the other 2-fold degenerate) I was able to keep the degeneracy low, and avoid all non-serine codons. Another example is shown in the case of

primers "5inE" and "5inR", which again are identical except at one codon. 4) including partial codons at the ends of the primers The various codons encoding an amino acid or a set of similar amino acids are often identical at their first (and maybe second) positions, but different at their third position. You can take advantage of this by synthesizing only the first or first and second positions of the 3' most codon covered by your primer pools, thus giving you one or two extra positions of exact match base pairs without adding any degeneracy. In the egl-10 example, the primer pools "3T" and "3A" cover a stretch in which the last codon must encode proline or glutamine. The codons for these two amino acids all start with C, but their last two positions are degenerate. Therefore, only the nondegenerate C was included in the primer pools.5) use of codon bias Some organisms have strong biases for using particular codons to encode certain amino acids. In theory you could reduce the degeneracy of a primer pool by only including these most common codons, and taking the risk that the gene(s) you are looking for will follow the organism's general codon bias enough to allow such primer pools to work. I haven't heard of anyone actually using this codon bias strategy in a successful degenerate PCR experiment, but you might try it if you're desperate.

Other considerations in primer design

1) primer lengthIn the example below the short stretches of sequence similarity among the egl-10 family members forced me to use primers only 19-21 bases long. These are shorter than the primers I have heard of people using in other successful experiments. For example, Linda Buck's primers were 31-33mers.2) 3' endPeople I talked to emphasized the special importance of having an exact match between the primer and template near the 3' end of the primer, although I'm not aware of specific data supporting this idea. For egl-10 I tried to avoid having any inosines near the 3' ends of the primers (except for primer "5out", which in fact failed to give any products), and also anchored the primers when possible with a nondegenerate codon at their 3' ends, so that 100% of the primers in the pool would be able to pair perfectly with the correct template over these last few bases.3) nested primersIf the sequence similarity in your gene family permits, it is a good idea to make nested sets of PCR primers. That way one round of PCR can be performed using the outside primers, and individual products (or the whole mix) can then be reamplified using the inside primers. Products amplified through both rounds are more likely to be the desired new gene family members, and less likely to be spurious products from sequences that happen to contain a couple of primer annealing sites by chance.

Determining optimal reaction conditions A number of parameters can be varied to optimize reaction conditions for degenerate PCR. These include: primer concentration, magnesium concentration, template concentration, number of cycles of amplification, and the temperatures and times of each step in the amplification cycle. If each of these parameters is to be independently varied, the number of possibilities quickly

reaches mind boggling proportions. My philosophy has been to fix almost all these parameters at the standard levels that have been successful for other people, and to vary only the one parameter that I think is the most crucial: the temperature of the annealing step during amplification.My standard PCR reactions are as follows:

1. 5 l template DNA (2-300 ng)5 l 5 l 10X PCR buffer (10X buffer=100 mM Tris pH 8.3, 500 mM KCl, 15 mM MgCl2, 0.01% gelatin)8 l dNTP mix (1.25 mM each dNTP)

2. 2 l "ampliTaq" polymerase (5 U/ l)25 l dH205 l each primer pool at 20 M eachtotal volume 50 l

In practice, the reactions are set up by placing the primers and template into a 0.5 ml tube, then adding two drops of mineral oil from a blue tip, and adding on top of the oil 38.5 l of a premix containing all the other components. In this way, it is easy to set up many different primer/template combinations at once. The tubes are then briefly spun in a microfuge to combine the two aqueous phases, and the tubes are immediately placed in the PCR block preheated to 95 for a "hot start".My amplification program:

95 X 3 min. (hot start)

?? X 1 min. (this annealing temperature is varied to optimize the amplification)72 X 2 min.94 X 45 sec.40 cycles of the above 3 steps

72 X 5 min.hold at 4

This takes ~4.5 hours to run on an MJ Research machine.

To test the primers and optimize the conditions, I do a series of amplification runs starting with an annealing temperature of 25 , and increasing in 5 increments until amplification fails to occur. Typically for each primer pair being tested, at each temperature, I run 3 reactions containing different templates: 1) a positive control containing 2 ng of a cloned member of the gene family of interest as template. 2) a negative control containing no template (this is very important-- you don't want to get fooled by contaminants). 3) an experimental reaction containing a complex template such as genomic DNA or total cDNA. For total C. elegans genomic DNA I've been using 300 ng as a template. Using rat brain cDNA as a template I amplified off of only 2 ng. However, this was only because I didn't have very much cDNA. If possible, it would be better to use ~200 ng of cDNA as a template, as Linda Buck did to amplify the odorant receptors.

Choice of template Genomic DNA has the advantage that all members of your gene family are present in equimolar

amounts, and genomic DNA is probably readily available. The obvious disadvantage is that introns may disrupt the primer sites, or may cause the amplification product to be so long that it is not amplified efficiently. cDNA templates, though harder to obtain, overcome this problem. A big advantage of cDNA is that the desired amplification products should be of a known size, and you can therefore easily pick them out from among spurious products of other sizes. Remember that the "correct" sized band amplified off a cDNA template may be a complex mixture of products from many gene family members, so you may have to analyze many clones generated from such a band to assess its complexity. Linda Buck used random primed cDNA for her template, presumably to avoid biasing the cDNA towards the 3' ends of transcripts. In my case, I knew that the region I was amplifying should be at the extreme 3' end of the coding sequence, so I used oligo-dT primed cDNA.For the lazy and rich, Clontech sells oligo-dT primed cDNA prepared from various tissues of many species to use as templates for PCR.

Analysis of PCR products After amplification run 20 l of each reaction out on an agarose gel. I use 2% 3:1 Nusieve:SeaKem LE agarose (you can buy this premixed from FMC) in 1X TAE. This gel is not very low melting, and thus isn't very suitable for cloning directly from the gel, but it gives very nice resolution. I use the 123 bp ladder from Gibco as a size standard. Obviously you expect to get products off the positive control, and not to get them off the negative control. Using the complex template, you will probably get a smear at the lower annealing temperatures, which will resolve into a small number of bands as the annealing temperature rises. I pick an annealing temperature that gives a modest number of bands, and then clone all these bands and sequence them.If no products are evident in the experimental samples, a good trick to try is to use 2 l of the apparently failed reaction as a template, and reamplify under the same conditions. This often gives visible products.If you want to clone products that are only barely visible, you can get more of them by just reamplifying the original reaction as described above. Another way to amplify individual products separately is to cut the bands out of the gel that was used to analyze the original reactions (actually I take a bore out of the gel with a Pasteur pipette), melt the DNA containing agarose, and use 2 l of it as a template to reamplify under the same conditions.The above described method for reamplifying specific bands can (and should) be used to test amplified products to see if they are single primer artefacts. Use 2 l of an agarose gel bore to set up each of three PCR reactions, containing either individual primer or both together. Obviously, you're only interested in products that require both primers in order to be amplified.I clone PCR products by running the PCR reaction out on a low melt agarose gel (2% Nusieve agarose in 1X TAE). I cut the desired band out, melt it at 70 , mix well by pipetting up/down, and use 5 l of the melted agarose directly in a ligation reaction with a dT tailed vector. This vector DNA is prepared as follows: cut 1 g bluescript SK with EcoRV in a 20 l reaction. Add 20 l 1X PCR buffer, and 2 l 2 mM dTTP. Add 0.5 l "ampliTaq" polymerase (2.45 U), and incubate at ~72 for 20 min. Run the DNA out on a 0.8% Seaplaque agarose gel in 1X TAE, cut out the band, melt at 70 , mix well, and use 5 l in a ligation reaction. It turns out that only about 50% of the colonies obtained after transformation of this type of reaction may have inserts; the

rest are vector reclosures. However, if blue/white selection is used, virtually all the white colonies have inserts.