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PCR amplification and error propagation

PCR amplification and error propagation


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Say I have one dsDNA that undergoes normal PCR (where amplification is exponential). If there is a mistake, say a G is swapped for an A during the 2nd round of replication, what percentage of the final DNA will have the mistake if there are 12 more rounds?

I was talking to my teacher about this today. I say 33%, my teacher said 12.5%.


I will format it to .ppt as soon as I have more time! If something is not readable, please let me know!

I have to point out several things:

  1. The fraction of incorrect DNA molecules does not depend on the number of cycles (as long as the number of cycles is higher than 2).

  2. There is one exception: if the mutation occurs outside of the region to be amplified (for example at the 4th molecule, downstream of the region of interest), the total number of wrong DNA molecules will be 0.


Your teacher is indeed correct.

In the first round you would get two identical molecules of the dsDNA.

In the second round you would get 3 identical molecules and one molecular with anAsubstituted for aGin one of the strands. ie.

No error (3 of the 4 molecules):

------G------- ------C-------

One mismatch (1 of the 4 molecules):

------A------- ------C-------

So there are a total of 8 strands of DNA after the second round and one of those strands has the mismatch.1 / 8= 0.125 = 12.5%

In round 3 you would have 8 dsDNA molecules and only one of those 8 dsDNA molecules would have the mismatch.


What Is Polymerase Chain Reaction (PCR)?

PCR stands for polymerase chain reaction, a molecular biology technique for amplifying segments of DNA, by generating multiple copies using DNA polymerase enzymes under controlled conditions. As little as a single copy of a DNA segment or gene can be cloned into millions of copies, allowing detection using dyes and other visualization techniques.

Developed in 1983, the process of PCR has made it possible to perform DNA sequencing and identify the order of nucleotides in individual genes. The method uses thermal cycling or the repeated heating and cooling of the reaction for DNA melting and replication. As PCR continues, the “new” DNA is used as a template for replication and a chain reaction ensues, exponentially amplifying the DNA template.

PCR techniques are applied in many areas of biotechnology including protein engineering, cloning, forensics (DNA fingerprinting), paternity testing, the diagnosis of hereditary and/or infectious diseases, and for the analysis of environmental samples.

In forensics, in particular, PCR is especially useful because it amplifies even the smallest amount of DNA evidence. PCR can also be used to analyze DNA that is thousands of years old, and these techniques have been used to identify everything from an 800,000-year-old mammoth to mummies from around the world.


PCR Cloning Method

PCR cloning differs from traditional cloning in that the DNA fragment of interest, and even the vector, can be amplified by the Polymerase Chain Reaction (PCR) and ligated together, without the use of restriction enzymes. PCR cloning is a rapid method for cloning genes, and is often used for projects that require higher throughput than traditional cloning methods can accommodate. It allows for the cloning of DNA fragments that are not available in large amounts.

Typically, a PCR reaction is performed to amplify the sequence of interest, and then it is joined to the vector via a blunt or single-base overhang ligation prior to transformation. Early PCR cloning often used Taq DNA Polymerase to amplify the gene. This results in a PCR product with a single template-independent base addition of an adenine (A) residue to the 3' end of the PCR product, through the normal action of the polymerase. These "A-tailed" products are then ligated to a complementary T-tailed vector using T4 DNA ligase, followed by transformation.

High-fidelity DNA polymerases are also now routinely used to amplify sequences with the PCR product containing no 3' extensions. The blunt-end fragments are joined to a plasmid vector through a typical ligation reaction or by the action of an "activated" vector that contains a covalently attached enzyme, typically Topoisomerse I, which facilitates the vector:insert joining. Some PCR cloning systems contain engineered "suicide" vectors that include a toxic gene into which the PCR product must be successfully ligated to allow propagation of the strain that takes up the recombinant molecule during transformation.

A typical drawback common to many PCR cloning methods is a dedicated vector that must be used. These vectors are typically sold by suppliers, like NEB, in a ready-to-use linearized format and can add significant expense to the total cost of cloning. Also, the use of specific vectors restricts the researcher's choice of antibiotic resistance, promoter identity, fusion partners, and other regulatory elements.

  • High efficiency, with dedicated vectors
  • Amenable to high throughput
  • Limited vector choices
  • Higher cost
  • Lack of sequence control at junction
  • Multi-fragment cloning is not straight forward
  • Directional cloning is difficult

PCR Cloning

Note that times are based on estimates for moving a gene from one plasmid to another. If the source for gene transfer is gDNA, add 2 hours to calculation for the traditional cloning method. Total time does not include transformation, isolation or analysis.

Analysis of the PCR product

There are two main methods of visualizing the PCR products: (1) staining of the amplified DNA product with a chemical dye such as ethidium bromide, which intercalates between the two strands of the duplex or (2) labeling the PCR primers or nucleotides with fluorescent dyes (fluorophores) prior to PCR amplification. The latter method allows the labels to be directly incorporated in the PCR product. The most widely used method for analyzing the PCR product is the use of agarose gel electrophoresis, which separates DNA products on the basis of size and charge. Agarose gel electrophoresis is the easiest method of visualizing and analyzing the PCR product. It allows for the determination of the presence and the size of the PCR product ( Figure 2 ). A predetermined set of DNA products with known sizes are run simultaneously on the gel as standardized molecular markers to help determine the size of the product.

From Riedl et al, 2004: Identification of an alternatively spliced mouse Langerin transcript. (a) Ethidium bromide-stained agarose gel showing PCR products from full-length mouse Langerin obtained from C57BL/6 fresh LC (fLC) and from fetal skin-derived dendritic cells (FSDDC).

Typically, when PCR is used to detect the presence or absence of a specific DNA product, it is called qualitative PCR. Qualitative PCR is a good technique to use when PCR is performed for cloning purposes or to identify a pathogen. For example, in the report by Dworkin et al, qualitative PCR was used to detect the presence of Merkel cell polyomavirus in cutaneous squamous cell carcinoma (SCC) in immunocompetent individuals (2009). Using genomic DNA isolated from SCCs excised from immunocompetent individuals and primers specific to virus genes, the investigators were able to demonstrate the presence of a 351base pair (bp) viral gene in 6 out of 16 samples tested, by the presence of a PCR-product band about 351 bp long, as seen on a 2% agarose gel with ethidium bromide ( Figure 3 ). The experiment also included template DNA from a polyomavirus containing plasmid as a positive control (P) and a negative water control (W). The first lane marked by (M) is the molecular marker, which is used to identify the size of the detected PCR product. The presence of a viral specific gene detected by PCR is marked by (+) absence of viral gene is marked by (−).

From Drowkin et al, 2009: MCPyV detection. (a) The presence of MCPyV in SCCs, genomic normal DNA, and adjacent skin DNA was determined by PCR using VP1 primers. A representational result is shown with 6 of 16 samples tested showing a PCR product at 351 bp. All experiments included DNA from an MCPyV plasmid as a positive control (P) and a negative water control (W). M, molecular weight marker +, positive for virus −, negative for virus.


PCR Troubleshooting Guide

Common issues in PCR are mainly associated with reaction conditions, sequence accuracy, and amplification yield and specificity. On this page, learn about their possible causes and our recommendations on how to resolve these issues.

On this page:

  • Minimize shearing and nicking of DNA during isolation. Evaluate template DNA integrity by gel electrophoresis, if necessary.
  • Store DNA in molecular-grade water or TE buffer (pH 8.0) to prevent degradation by nucleases.
  • Follow manufacturer recommendations stringently when using purification kits to isolate template DNA. Consult the user manual and troubleshooting guides to mitigate poor DNA quality.
  • Ensure that no residual PCR inhibitors such as phenol, EDTA, and proteinase K are present if following chemical or enzymatic DNA purification protocols.
  • Re-purify, or precipitate and wash DNA with 70% ethanol, to remove residual salts or ions (e.g., K + , Na + , etc.) that may inhibit DNA polymerases.
  • Choose DNA polymerases with high processivity, which display high tolerance to common PCR inhibitors carried over from soil, blood, plant tissues, etc.
  • Examine the quantity of input DNA and increase the amount if necessary.
  • Choose DNA polymerases with high sensitivity for amplification.
  • If appropriate, increase the number of PCR cycles.
  • Choose DNA polymerases with high processivity, which display high affinity for DNA templates and are more suitable to amplify difficult targets.
  • Use a PCR additive or co-solvent to help denature GC-rich DNA and sequences with secondary structures.
  • Increase denaturation time and/or temperature to efficiently separate double-stranded DNA templates.
  • Check amplification length capability of the selected DNA polymerases. Use DNA polymerases specially designed for long PCR.
  • Choose DNA polymerases with high processivity, which can amplify long targets in a shorter time.
  • Reduce the annealing and extension temperatures to help primer binding and enzyme thermostability.
  • Prolong the extension time according to amplicon lengths.
  • Review primer design. Use online primer design tools when appropriate.
  • Ensure that the primers are specific to the target of interest.
  • Verify that the primers are complementary to the correct strands of the target DNA.
  • Aliquot primers after resuspension and store properly.
  • Reconstitute fresh primer aliquots, or obtain new primers if necessary.
  • Optimize primer concentrations (usually in the range of 0.1–1 μM).
  • For long PCR and PCR with degenerate primers, start with a minimum concentration of 0.5 μM.
  • Use hot-start DNA polymerases to prevent degradation of primers by the 3’→5’ exonuclease activity of proofreading DNA polymerases. Hot-start DNA polymerases also increase yields of the desired PCR products by eliminating nonspecific amplification.
  • Alternatively, set up PCR on ice, or add DNA polymerase last to the reaction mixture.
  • Choose DNA polymerases with high sensitivity for amplification.
  • Review recommendations on the amount of DNA polymerase to use in PCR, and optimize as necessary.
  • Increase the amount of DNA polymerase if the reaction mixture contains a high concentration of an additive (e.g., DMSO, formamide) or inhibitors from the sample sources.
  • Optimize Mg 2+ concentration for maximum PCR yields. The presence of EDTA, other metal chelators, or atypically high concentrations of dNTPs may require a higher Mg 2+ concentration.
  • Check the DNA polymerase’s preference for magnesium salt solutions. For example, Pfu DNA polymerase works better with MgSO4 than with MgCl2.
  • Review the recommended concentrations of PCR additives or co-solvents. Use the lowest possible concentration when appropriate.
  • Adjust the annealing temperatures, as high concentrations of PCR additives or co-solvents weaken primer binding to the target.
  • Increase the amount of DNA polymerase, or use DNA polymerases with high processivity.
  • Consider using an additive or co-solvent specifically formulated for a given DNA polymerase (e.g., GC Enhancer supplied with Invitrogen Platinum DNA polymerases).
  • Ensure that the selected DNA polymerases are able to incorporate the modified nucleotides.
  • Optimize the ratio of the modified nucleotide to dNTP to increase PCR efficiency.
  • Mix the reagent stocks and prepared reactions thoroughly to eliminate density gradients that may have formed during storage and setup.
  • Optimize the DNA denaturation time and temperature. Short denaturing times and low temperatures may not separate double-stranded DNA templates well. On the other hand, long denaturation times and high temperatures may reduce enzyme activity.
  • Optimize the annealing temperature stepwise in 1–2°C increments, using a gradient cycler when available. The optimal annealing temperature is usually 3–5°C below the lowest primer Tm.
  • Adjust the annealing temperature when a PCR additive or co-solvent is used.
  • Use the annealing temperature recommended for a specific DNA polymerase in its optimal buffer. Annealing temperature rules for primer sets can vary between different DNA polymerases.
  • Select an extension time suitable for the amplicon length.
  • Reduce the extension temperature (e.g., to 68°C) to keep the enzyme active during amplification of long targets (e.g., >10 kb).
  • Use DNA polymerases with high processivity for robust amplification even with short extension times.
  • Adjust the number of cycles (generally to 25–35 cycles) to produce an adequate yield of PCR products. Extend the number of cycles to 40 if DNA input is fewer than 10 copies.
  • Review the optimal amounts of DNA input. Lower the quantity to reduce the generation of nonspecific PCR products.
  • Degraded DNA may appear as smears or lead to high background in gel electrophoresis. Minimize shearing and nicking of DNA during isolation.
  • Evaluate the integrity of the template DNA prior to PCR by gel electrophoresis, if necessary. Store DNA in molecular-grade water or TE buffer (pH 8.0) to prevent degradation by nucleases.
  • Choose DNA polymerases with high processivity, which display high affinity for DNA templates and are more suitable to amplify difficult targets.
  • Use a PCR additive or co-solvent to help denature GC-rich DNA and sequences with secondary structures. Increase denaturation time and/or temperature to efficiently separate double-stranded DNA templates.
  • Check amplification length capability of the selected DNA polymerases. Use DNA polymerases specially designed for long PCR.
  • Choose DNA polymerases with high processivity, which can amplify long targets in a shorter time.
  • Reduce the annealing and extension temperatures to help primer binding and enzyme thermostability. Prolong the extension time according to amplicon lengths.
  • Review primer design. Use online primer design tools when appropriate. Verify that the primers are specific to the target, with minimal homology to other regions in the template.
  • Ensure that the primers do not contain complementary sequences or consecutive G or C nucleotides at the 3′ ends, to prevent primer-dimer formation.
  • Avoid direct repeats in the primers to prevent misalignment in binding to the target. Consider longer primers to enhance specificity.
  • Consider nested PCR to improve specificity.
  • Optimize primer concentrations (usually in the range of 0.1–1 μM). High primer concentrations promote primer-dimer formation.
  • Review recommendations on the amount of DNA polymerase to use in PCR, and decrease as necessary.
  • Use hot-start DNA polymerases that have no activity at room temperature but are functional only after a high-temperature activation step, to enhance specificity. With non–hot-start DNA polymerases, set up PCR on ice to keep enzyme activity low.
  • Review Mg 2+ concentrations and lower as appropriate to prevent nonspecific PCR products. Optimize Mg 2+ concentrations for each primer set and target DNA.
  • Increase denaturation time and/or temperature to efficiently separate DNA when working with GC-rich templates and sequences with secondary structures.
  • Use the annealing temperature recommended for a specific DNA polymerase in its optimal buffer. Annealing temperature rules for primer sets can vary between different DNA polymerases.
  • Increase the annealing temperature to improve specificity. The optimal annealing temperature is usually no less than 3–5°C below the lowest primer Tm.
  • Optimize the annealing temperature stepwise in 1–2°C increments, using a gradient cycler when available.
  • Consider touchdown PCR to enhance specificity.
  • Shorten the annealing time to minimize primer binding to nonspecific sequences.
  • Reduce the extension temperature 3–4°C to help the DNA polymerase’s thermostability, especially for long PCR.
  • Prolong the extension time when amplifying long DNA targets.
  • Include a final extension step with sufficient time (5–15 minutes) to extend the whole target.
  • Reduce the number of cycles, without drastically lowering the yield of the desired PCR products, to prevent accumulation of nonspecific amplicons.
  • Use DNA polymerases with exceptionally high fidelity to generate PCR fragments for downstream applications such as cloning, sequencing, and site-directed mutagenesis.
  • Review Mg 2+ concentrations and reduce as necessary. Excessive concentrations favor misincorporation of nucleotides by DNA polymerases.
  • Ensure equimolar concentrations of dATP, dCTP, dGTP, and dTTP in the reaction. Unbalanced nucleotide concentrations increase the PCR error rate.
  • Reduce the number of cycles without drastically lowering the yield of the desired PCR products. High numbers of cycles increase the incorporation of mismatched nucleotides.
  • Increase the amount of input DNA when appropriate to avoid running an excessive number of cycles.
  • Use a long-wavelength UV (360 nm) light box to visualize fragments in gels, and limit the illumination time as much as possible.
  • If using a short-wavelength (254–312 nm) light box, limit the UV illumination to a few seconds and keep the gel on a glass or plastic plate.
  • Alternatively, use dyes with longer-wavelength (less damaging) excitation to visualize the DNA.
  • Sequence both DNA strands to verify the reliability of sequencing results. Use duplicate samples when appropriate.
  • Avoid direct repeats within primer sequences, as multiple repeats may appear from sequence misalignment at the ends of the PCR products.
  • Order PCR primers with purification to remove non–full-length DNA oligos, which are truncated at their 5′ ends.
  • Use molecular-grade, nuclease-free reagents in PCR setup. Set up reactions on ice to keep the activity of possible contaminating exonucleases low.
  • Use a long-wavelength UV (360 nm) light box to visualize fragments in gels, and limit the illumination time as much as possible.
  • If using a short-wavelength (254–312 nm) light box, limit the UV illumination to a few seconds and keep the gel on a glass or plastic plate.
  • Alternatively, use dyes with longer-wavelength (less damaging) excitation to visualize the DNA.
  • Sequence both DNA strands to verify the reliability of sequencing results. Use duplicate samples when appropriate.
  • Avoid primers containing complementary and self-complementary sequences, which favor primer-dimer formation and self-oligomerization and their subsequent amplification.
  • Avoid contamination of DNA in the work environment by following general recommendations for PCR setup.
  • Use pipette tips with aerosol barriers. Dedicate a separate work area and decontaminate the area after each use.
  • Follow PCR carryover control techniques such as dUTP incorporation with UDG treatment.

For more troubleshooting assistance, please visit our End-Point PCR and PCR Primers Support Center or contact our technical support team.


PCR amplification and error propagation - Biology

qPCR is more complex than perceived by many scientists.

The production of an amplification curve and an associated quantitative cycle value does not necessarily mean interpretable data.

The MIQE guidelines and associated methodology articles published thereafter, underline the ongoing drive to help scientists produce reproducible data from qPCR, culminating in a simple, stepwise methodology to ensure high-quality, reproducible data from qPCR experiments.

The concept of data normalization has led to the ongoing publication of articles solely focused on this subject for various sample types and experimental parameters.

The analysis of qPCR data can be challenging, especially as experiments grow in sample number and complexity of biological groups. A defined approach to qPCR data analysis is necessary to clarify gene expression analysis.

Quantitative PCR (qPCR) is one of the most common techniques for quantification of nucleic acid molecules in biological and environmental samples. Although the methodology is perceived to be relatively simple, there are a number of steps and reagents that require optimization and validation to ensure reproducible data that accurately reflect the biological question(s) being posed. This review article describes and illustrates the critical pitfalls and sources of error in qPCR experiments, along with a rigorous, stepwise process to minimize variability, time, and cost in generating reproducible, publication quality data every time. Finally, an approach to make an informed choice between qPCR and digital PCR technologies is described.


Whole genome amplification (WGA)

Most cells contain only one or a few copies of their genome, constituting picograms of DNA, which is not enough for direct analysis with current sequencing technologies. Assaying single cell DNA for diagnostics, which can require multiple experiments with limited sample, also demands rapid unbiased DNA amplification. The research on ancient DNA, which is often fragmented, poses yet another type of challenge. These are areas where isothermal amplification technology is used to increase DNA material needed for downstream analysis.

To increase the amount of limited DNA targets, isothermal whole genome amplification (WGA) is the most efficient technique.[3] This is particularly useful in genetic disease research, where many repetitions are required. DNA amplified by WGA is used in downstream next-generation sequencing, Sanger sequencing, genotyping with microarrays, and single nucleotide polymorphism (SNP) genotyping.[4] Various WGA techniques have been developed that differ both in their protocols and ease of use.

Phi29 DNA polymerase is the main enzyme of choice for WGA. Thermo Scientific also offers an improved EquiPhi29 DNA polymerase, which is a proprietary Phi29 DNA polymerase mutant developed through in vitro protein evolution.[5] This enzyme is significantly superior over Phi29 in protein thermostability, reaction speed, product yield, and amplification bias. Moreover, it retains all the benefits of the wild-type enzyme, including high processivity (up to 70 kb), strong strand displacement activity, and 3'–5' exonuclease (proofreading) activity. For this reason, exo-resistant random primers are recommended. Table 3 compares classical Phi29 and EquiPhi29 DNA polymerases.

Table 3. Comparison of Phi29 and EquiPhi29 DNA polymerases with supporting data.

Phi29-type DNA PolymerasesEquiPhi29 DNA Polymerase
Processivity/strand displacementHigh (up to 70 kb)High (up to 70 kb)
Optimal amplification temperature30-37°C42-45°C
Reaction timeSlow – up to 12hSlow – up to 3h
Proofreading3'–5' (low error rates)3'–5' (low error rates)
AccuracyHigh (low error rates)High (low error rates)
YieldHighVery high
Sequence bias (preference)Low biased, uniform amplification of long fragments (whole genome)Very low bias, including GC and AT rich (data valid for 0.5 ng starting material)

Error Rate Comparison during Polymerase Chain Reaction by DNA Polymerase

As larger-scale cloning projects become more prevalent, there is an increasing need for comparisons among high fidelity DNA polymerases used for PCR amplification. All polymerases marketed for PCR applications are tested for fidelity properties (i.e., error rate determination) by vendors, and numerous literature reports have addressed PCR enzyme fidelity. Nonetheless, it is often difficult to make direct comparisons among different enzymes due to numerous methodological and analytical differences from study to study. We have measured the error rates for 6 DNA polymerases commonly used in PCR applications, including 3 polymerases typically used for cloning applications requiring high fidelity. Error rate measurement values reported here were obtained by direct sequencing of cloned PCR products. The strategy employed here allows interrogation of error rate across a very large DNA sequence space, since 94 unique DNA targets were used as templates for PCR cloning. The six enzymes included in the study, Taq polymerase, AccuPrime-Taq High Fidelity, KOD Hot Start, cloned Pfu polymerase, Phusion Hot Start, and Pwo polymerase, we find the lowest error rates with Pfu, Phusion, and Pwo polymerases. Error rates are comparable for these 3 enzymes and are >10x lower than the error rate observed with Taq polymerase. Mutation spectra are reported, with the 3 high fidelity enzymes displaying broadly similar types of mutations. For these enzymes, transition mutations predominate, with little bias observed for type of transition.

1. Introduction

With the rapid pace of developments in systems biology-based research, for example, genomics, proteomics, and metabolomics, larger-scale biological discovery projects are becoming more common. Put differently, the scope of many projects has changed from the study of one/few targets to the study of hundreds, thousands, or more. An example of research that has been transformed by developments in systems biology is the cloning of expressed open reading frames (ORFs) from cDNA substrates. The traditional path for ORF cloning has usually started with experimental observations driving the identification of one or several genes of interest to a particular pathway. Cloning of target(s) then typically resulted in further refinements of pathway details and often identification of new cloning targets. With the creation and continual refinements of databases of genomic sequences, cloning now often takes place on a much larger scale. Microarray technology and DNA sequencing breakthroughs have led to a vast increase in the number of ORFs present in biological databases. Furthermore, biological observations no longer necessarily precede target identification, which now is often driven in large part by bioinformatics-based predictions and analyses. Examples of large-scale cloning efforts include structural genomics projects to systematically determine protein structures [1], pathogen ORF cloning to understand disease and therapeutic mechanisms [2], and creation of the entire human ORFeome which will further developments in basic and applied biomedical sciences [3].

DNA polymerases used to amplify targets during PCR cloning are high fidelity enzymes with error frequencies typically in the range of

mutations/bp amplified [4]. Minimizing PCR-generated errors is especially important for larger-scale cloning projects because, given a sufficiently large pool of target DNA sequence, even high fidelity enzymes will produce clones with mutations. There are a variety of methods to assay the fidelity of a DNA polymerase. However, error frequencies for PCR enzymes are almost always assayed using one (or a few) defined DNA target that samples a limited portion of DNA sequence space. Early studies using the relatively low-fidelity Taq DNA polymerase relied on the sequencing of cloned PCR products (e.g., [5, 6]). Direct sequencing of clones was a practical approach at the time due to the low fidelity of the polymerase that is, most clones that were sequenced would contain at least one mutation.

With the introduction of higher fidelity polymerases, new screening methods were developed to rapidly interrogate large numbers of PCR products for the presence of mutations. These assays were based on a forward mutation fidelity assay developed by Kunkel and colleagues, which used a gap-filling reaction with a DNA polymerase on a lacZ template sequence, followed by ligation and transformation into E. coli. Colorimetric screening based on a functional lacZ gene allowed rapid identification of mutations, which were subsequently sequenced to determine the nature of the DNA alteration [7]. A similar approach was used to screen PCR products for mutations, by cloning a lacZ fragment amplified by PCR as opposed to simple gap filling by DNA polymerases. This method, sometimes using a different reporter gene, has been used to screen a variety of high fidelity PCR enzymes and to optimize PCR reaction conditions to minimize mutations [4, 8]. Finally, methods that rely on assaying PCR mutations based on differing chemical properties (i.e., melting temperature) of reaction products with mismatches relative to perfect duplexes have been developed and applied to a variety of enzyme systems [9, 10]. While reported fidelity values differ among research groups and assay methods, there is a general consensus that a relatively low-fidelity enzyme such as Taq has a fidelity value in the

range and higher fidelity enzymes have values that are in the range (usually reported as mutations per bp per template doubling).

A tradeoff involved in using screening methods like those described above is that generally only one DNA sequence is interrogated during the assay. Additionally, limitations built into the assays further restrict the possible mutations that can be detected. For example, the assay based on screening lacZ gene amplification products uses a single 1.9 kb target, of which only 349 bases will produce a color change when mutated [11]. Likewise, assaying mutations based on differential duplex melting profiles is restricted to unique target sequences that are short enough, typically in the 100–300 bp range, and have thermal melting profiles that allow resolution of single mismatches [9, 10].

Because polymerase errors are known to be strongly dependent on DNA sequence context (reviewed in [12]), ideally one would use a large set of DNA sequences when measuring enzyme fidelity. This becomes especially relevant in the context of large-scale cloning projects, which involve hundreds or thousands of targets and thus contain an almost infinite DNA sequence space. To this end, we have designed and executed a study that measures enzyme fidelity by direct sequencing of cloned PCR products. Falling costs for DNA sequencing have made this method of fidelity determination practical, even for enzymes that make few mistakes. Our goals are to compare fidelity values derived from direct clone sequencing to those derived from screening-based methods, as well as to evaluate these results in the context of choosing an enzyme for a high-throughput cloning project.

2. Results and Discussion

To determine error rates and observe mutational spectra for a variety of DNA polymerases used in PCR cloning, we directly sequenced clones produced from 94 different plasmid templates. These plasmids, each with a unique target DNA sequence, are a subset of a larger group of glycosyltransferase clones that we have prepared from Arabidopsis thaliana cDNA (manuscript in preparation). The 94 plasmids have inserts with size ranging from 360 bp to 3.1 kb (median 1.4 kb) and GC content ranging from 35% to 52% (median 44%). A summary of the 6 DNA polymerases used in this study is presented in Table 1. We included Taq polymerase in our study because of the extensive body of literature that exists on the fidelity properties of this enzyme. The other enzymes included are all typically classified as “high fidelity” and therefore are potential candidates for large-scale cloning projects. And while comparison of fidelity values is difficult due to differences in assay and quantitation methods among different studies, a general ranking of the enzymes studied here (lowest fidelity to highest) appears to be Taq < AccuPrime-Taq < KOD

Pfu Pwo < Phusion.

Our cloning pipeline uses recombinational insertion of purified PCR products into a plasmid vector using the Gateway cloning system, a method widely used for high-throughput cloning studies (reviewed in [17]). Since our input plasmid DNA templates were prepared using the Gateway system, the target genes of interest are all flanked by att recombination sequences. This allowed the use of common primers for all PCR reactions, thus eliminating the need for target-specific optimizations. Purified plasmid DNA was used as template for PCR, and in all cases vendor-recommended buffers were used. We used small amounts of plasmid template (25 pg/rxn), in order to maximize the number of doublings in the PCR reaction, and the size of insert relative to total plasmid size was taken into account to determine the amount of target fragment present in the template. The PCR protocol used 30 cycles of amplification, with an extension time of 2 minutes/cycle for targets ≤2 kb (82 of 94 targets) and 4 minutes/cycle for targets >2 kb (12 of 94 targets). Figure 1 shows gel images for a representative set of PCR reactions for each enzyme. In all cases, a single major product band migrating at the expected size was observed. Amplification efficiency was measured by quantitation of PCR product using a dsDNA-specific dye and calculating the fold-amplification based on a known quantity of input DNA template. The fold-amplification is used to determine the number of template doublings that occurred during PCR. As reported in Table 2, amplification efficiency values were fairly uniform for all samples within a plate. We observe similar amplification efficiencies between different enzymes, with the exception that we routinely observed fewer template doublings in reactions with Pfu polymerase. We have kept thermocycling protocols constant for all enzymes, and thus it is possible that some parameters were not optimal for amplification by Pfu.

) is the average of doubling values for each of the 94 PCR reactions in one plate, where doublings are calculated from the formula

= (ng DNA after PCR/ng DNA input). Error rate (f) is calculated as f =

is the number of mutations observed for all clones that were sequenced and the (target size ×

for each target that was cloned.


Representative agarose gel electrophoresis images of products of PCR amplification of 24 unique DNA targets, using six different enzymes. Each lane contains 1/25 of the entire PCR reaction. Expected product sizes range from 1.4 to 1.7 kb in size.

Following amplification, PCR products were purified by precipitation with PEG/MgCl2, which is known to selectively fractionate DNA on the basis of size [18], to remove short products <300 bp in size. This precipitation step can be performed in 96-well plate format, which is a requirement when the number of samples becomes large. We have adopted this protocol for routine use and have observed a higher efficiency for insertion of correct-size DNA into the vector compared to purification using kit-based PCR purifications, which typically have size cutoffs of

100 bp (data not shown). In the case where off-target PCR products of >300 bp are present, gel extraction is used to isolate the desired product. Purified PCR products were incubated with vector DNA and BP Clonase II and transformed into competent cells. Three colonies per plate were picked and grown up in 96-well plates, and cultures were screened for correct-size insert by colony PCR. Insertion efficiency values for BP Clonase II, expressed as the average number of clones having an insert at/near the expected size (out of 3 colonies screened per transformation), were typically 80–90% (data not shown). For each target, one or more clones for each target containing a correct-size insert (if obtained) were cultured and used for DNA sequencing.

For method validation purposes, we used Taq DNA polymerase, a Family A DNA polymerase and the enzyme used in the earliest PCR experiments [6]. As an early workhorse in PCR technology, Taq polymerase has been studied extensively for purposes of fidelity determination. Taq DNA polymerase lacks a 3′→5′ exonuclease activity and thus is unable to correct misincorporated nucleotides that occur during DNA synthesis. Various assays have been used to assay Taq fidelity, and, depending on the method used, error rate values (expressed as mutations per base pair per template duplication) for Taq polymerase range from

(e.g., [7, 20]). Furthermore, the mutational spectrum of Taq polymerase has been characterized, with A•T → G•C transitions predominating due to the propensity for the enzyme to misincorporate incoming dCTP with a template thymine nucleotide [6, 9, 21].

By direct sequencing of clones from two independent PCR experiments with Taq polymerase, we observed 99 unique mutations out of >100 kbp of target DNA sequence. The type and number of individual mutations are listed in Table 3. Given the amplification efficiency of each PCR reaction, the error rate (average of 2 experiments) for Taq polymerase is

mutations/bp per template duplication. This value is in excellent agreement with other published values for this enzyme, and the relatively high variance suggests that calculated error values differing by up to 2-fold are probably not significant relative to the experimental noise. The majority of the mutations (67 of 99) are A•T → G•C transitions, which could result from either incoming dCTP mispairing with template A or incoming dGTP mispairing with template T. Transitions of the G•C→A•T type, resulting from either incoming TTP mispairing with template G or incoming dATP mispairing with template C, are the second most prevalent mutation (28 of 99). There were 3 transversion mutations, with 1 A•T→T•A and 2 A•T→C•G changes. Overall, the spectrum of the base substitution mutations agrees well with previous observations on Taq polymerase reported in the literature [7]. There was only one insertion or deletion (indel) mutation observed in our data set, a single T deletion in a

template sequence. Taq polymerase has been reported to produce indel mutations with a significant frequency, as much as approximately 25% of total mutations, with all occurring in homopolymeric runs [7]. Since our target pool contains 1481 instances of homopolymer runs of at least 4 bp, we suspect that other differences between the earlier assay conditions and those used here explain the discrepancy. Specifically, the earlier experiments were performed with elevated magnesium (10 mM versus 1.5 mM used here) and elevated dNTP levels (1 mM versus 0.2 mM used here). Both elevated magnesium and dNTP levels were subsequently shown to elevate frameshift (indel) mutations preferentially relative to base substitution mutations [21].

An important control for these experiments is necessitated by the method used to generate template for DNA sequencing. For larger-scale cloning projects, DNA sequencing using cell culture is advantageous because of the saving in time and resources relative to purifying plasmid DNA. However, sequencing using cell culture requires a PCR or another amplification step, and this step could in principle be a source of “additional mutation.” To address this directly, we sequenced miniprep DNA prepared from a subset of clones produced with Taq polymerase. Every one of the fourteen mutations detected in the subset using cell culture as the source for sequencing template was also observed when sequencing from plasmid DNA template (data not shown). We conclude that our method has a false positive rate of <7% (1/14) and is acceptable for assaying PCR-induced mutations. Furthermore, based on our results with Taq polymerase, we conclude that our method for fidelity determination gives results in excellent agreement with other studies and is thus an accurate measure of polymerase accuracy.

Our results indicate that 3 of the enzymes included in the study, Pfu polymerase, Phusion Hot Start, and Pwo polymerase, have error rates that are significantly lower than the others. This is consistent with previous findings demonstrating very high fidelity PCR amplification for these enzymes. Interestingly, error frequency values for these three enzymes are extremely similar to each other, approximately 2-3 × mutations/bp/template doubling. The slight error frequency value differences are probably not significant, given that the small number of mutations is produced by these high fidelity polymerases in addition to the experimental variability discussed above for the results with Taq. Given the costs of cloning and sequencing and finite research budgets, mutation detection by DNA sequencing of clones generates a relatively small data set of mutations when the enzyme fidelity is high. This is a drawback to our assay, and despite the fact that DNA sequencing costs continue to drop screening bacteria is still a far more economical method of interrogating a large number of clones. For all mutant clones produced by Pfu, Phusion Hot Start, and Pwo polymerases, samples were resequenced to rule out sample processing or DNA sequencing as a source of error. In all cases, the original mutation was present, confirming the PCR reaction as the most likely source of the mutation. From the standpoint of use in a large-scale cloning project, any one of these enzymes would be acceptable, judged on the criteria of minimizing error rate. Other factors need to be considered of course, such as amplification efficiency, mutation spectra, performance with high GC content templates, and cost, to name a few. As far as mutation spectra, the 3 high fidelity polymerases all produced predominantly (>75%) transition mutations, with no significant template bias. With Phusion enzyme, we observed 15% (2/13) indel mutations, which are problematic for cloning applications where translation reading frame should be maintained. Both indel mutations occurred in repeat regions, with one being an A insertion into an

template sequence and the other being a (TCT) deletion within a

template sequence. This result was unexpected in light of the high processivity of Phusion polymerase relative to other commonly used PCR enzymes (vendor website). Because multiple studies have found that increased polymerase processivity reduces the frequency of slippage mutations that result in indel mutations [22, 23], we expected Phusion to produce the fewest of this class of errors. It should be noted, however, that this conclusion is based on a small sample size and a larger number of mutations should be analyzed for confirmation.

It was interesting to us that none of the enzymes tested here was found to have an error rate below

2 × . Other studies in the literature have reported sub-10 −6 error frequencies for PCR enzymes,

[10] for Pfu polymerase assayed by differential duplex Tm measurement and 4.2 × for Phusion, using HF buffer assayed with a method called BEAMING [16]. For the study of Phusion fidelity, the PCR used a different buffer than the one employed here, which according to the vendor does result in a 2-3-fold lower error rate. In addition, that study uses the BEAMING method, an extremely sensitive flow cytometric protocol that screens large numbers of beads that contain PCR products for the presence of nucleotide variations. However, only one specific mutation, a G•C → A•T mutation at a single position, was interrogated in that study. Thus, while the assay is extremely sensitive for detection of defined mutations, results obtained with the BEAMING method for mutation frequency at a single position may not necessarily reflect the fidelity properties of an enzyme for much larger sequence spaces. For the study on Pfu error rate, several fundamental methodological differences are present: in the earlier study, the PCR was performed under “almost anaerobic” conditions with significantly shorter cycling times, the target size was limited to 93 bp, and mutation detection relied on a physiochemical method: separation and isolation of PCR products containing mismatches by capillary electrophoresis [10]. And while this method has been successfully used in the detection of rare mutations in mitochondrial DNA samples from normal and cancer tissues [24], the requirement for a mutation to result in a molecule with an altered melting profile may bias the number of mutations that can be detected. A major discrepancy between our results and those from this earlier report on Pfu fidelity, which may be connected to the differing mutation detection methodologies, can be seen in the mutation spectra results in Table 3. We observed

90% (8 of 9) transition mutations, with a slight bias for G•C → A•T alterations. In contrast, the study using capillary electrophoresis for detection resulted in predominantly (3/5) transversion mutations, with a single A•T → G•C transition and a single 1 bp deletion mutation. Transversion mutations require the polymerase to synthesize either a purine•purine or a pyrimidine•pyrimidine mismatch, both of which are significantly disfavored relative to the different purine•pyrimidine mismatches in Family B polymerases, including Pfu polymerase [25, 26]. Because the types of mutations we observe are consistent with previously reported mutational spectra for other Family B polymerases, we believe our method has detected polymerase errors in a bias-free fashion.

The other two enzymes included in our study, KOD polymerase and AccuPrime-Taq High Fidelity, have fidelity values intermediate between Taq polymerase and the higher fidelity enzymes. The error rate observed for KOD polymerase was only

4-fold lower than that of Taq polymerase and

2.5-fold higher than for Pfu polymerase. The initial report on fidelity of KOD polymerase, a Family B/pola-like polymerase from Thermococcus kodakaraensis KOD1, reported an error rate very slightly lower than Pfu polymerase and

4-fold lower than for Taq polymerase [13]. That study used a forward mutation assay (not PCR), expressed fidelity simply as the ratio of white colonies to blue with no accounting for PCR amplification efficiency, and used experimental conditions (Mg 2+ concentration) that differ significantly from typical PCR conditions. A subsequent study measuring fidelity under PCR conditions, using a different reporter gene but still a simple ratio of mutant to wild-type colonies, reported error rates

50x lower than those with Taq and marginally lower than those for Pfu polymerase [14]. In neither of those studies was there a report of the molecular changes leading to mutant colonies. The large difference between these two results, which are from the same research group, serves to highlight the difficulties in making comparisons between studies where there are significant methodological differences. In the present study, we find that the mutation spectrum for KOD polymerase is similar to the other B-family polymerases (Pfu, Pwo, and Phusion) assayed here. As shown in Table 3, transitions predominate (14 of 16 mutations), with a slight bias (64%) for A•T → G•C mutations.

For the PCR performed with AccuPrime-Taq High Fidelity system, we observed a 3-fold improvement in fidelity relative to Taq polymerase. According to the vendor, AccuPrime-Taq High Fidelity is an enzyme blend that contains Taq polymerase, a processivity-enhancing protein, and a higher fidelity proofreading polymerase from Pyrococcus species GB-D. The lower error rate seen with AccuPrime-Taq most likely arises from the GB-D polymerase editing mistakes introduced by Taq polymerase as opposed to enhanced processivity since increased processivity has been shown to have no significant effect on base substitution errors [22, 27]. The mutation spectrum of the blend is almost identical to that seen with Taq polymerase alone, with transitions predominant and a significant bias for A•T → G•C changes (71% for AccuPrime-Taq versus 73% for Taq). However, it should be noted that a study on the mutation spectra of GB-D DNA polymerase (commercially available as Deep Vent) found A•T → G•C transitions to be the predominant mutation [28]. Detailed analysis on the contribution of each enzyme to the overall mutation spectrum is also precluded by the proprietary enzyme formulation used by the vendor.

In summary, we have used direct DNA sequencing of cloned PCR products to assay polymerase fidelity and evaluate other aspects of enzyme suitability for large-scale cloning projects. Based on minimizing PCR errors, Pfu polymerase, Pwo polymerase, and Phusion all produce acceptably low levels of mutations. Phusion was observed to produce more indel mutations than Pfu or Pwo polymerases, although the total number of mutations was limited. This type of mutation is particularly problematic for ORF cloning projects and should be taken into account in the process of enzyme selection. Aside from fidelity considerations, amplification efficiency values were significantly higher for Phusion and Pwo compared to Pfu, although further optimization of the PCR reaction for Pfu would likely improve efficiency values. Likewise, for cloning projects where targets are either very long or very highly GC-rich fidelity may be of lesser importance relative to the ability to amplify “difficult” target DNA. And finally, since the application space for PCR technology is huge, with cloning representing only a small fraction, enzymes other than those studied here need to be compared and evaluated based on project-specific needs and challenges.

3. Materials and Methods

3.1. PCR Reactions

All enzymes and reaction buffers were from commercial sources: Fermentas (Taq polymerase), Invitrogen/Life Technologies (AccuPrime-Taq), EMD Chemicals/Novagen (KOD Hot Start), Agilent (cloned Pfu polymerase), Finnzymes (Phusion Hot Start), and Roche (Pwo polymerase). PCR reactions were carried out in a final volume of 50 μL using buffer conditions and enzyme amounts recommended by the vendor. For reactions with Phusion, the GC buffer was used. In all cases, reactions included 0.2 mM each dNTP (Fermentas) and 0.2 mM each primer (IDT) with the sequences (5′ to 3′) GGGGACAAGTTTGTACAAAAAAGCAGGCTTCACC for the forward primer and GGGGACCACTTTGTACAAGAAAGCTGGGTC for the reverse primer. Template for PCR reactions was miniprep plasmid DNA, with each plasmid template containing a unique target sequence of known sequence and size, ranging from 0.3 to 3 kb. The target insert was cloned in between the att sites of a pDONR vector, allowing the use of a common primer set for all plasmids. Each PCR reaction contained 0.025 ng plasmid DNA, quantitated using the PicoGreen DNA quantitation reagent (Invitrogen/Life Technologies), and thus the amount of input target (i) was calculated as

ng (size of target ÷ (size of target + size of plasmid)). The thermocycling protocol for all reactions with target length ≤2 kb was 5 minutes, 95°C, then 30 cycles of 15 seconds, 95°C → 30 seconds, 55°C → 2 minutes, 72°C, and finally 7 minutes at 72°C. For the targets >2 kb in size, the 2-minute extension step was extended to 4 minutes. For analysis of PCR products by gel, 2 μL of each PCR reaction was run on a 2% agarose eGel (Invitrogen/Life Technologies) run according to vendor recommendations.

3.2. Quantitation of PCR Reactions

Efficiency of PCR amplification was determined by measuring the amount of product using a modified PicoGreen dsDNA quantitation assay. This method was facilitated by optimizing the PCR reaction to produce a single product band (Figure 1). Using a Biomek FX-P (Beckman) automated liquid handing system, 5 μL of each PCR reaction was diluted 50-fold in TE buffer (pH 8) into a new 96-well plate. From this plate, 5 μL from each well was mixed with 195 μL of PicoGreen solution, a 500-fold dilution of dye in TE (pH 8). Fluorescence measurements were taken with a Paradigm (Beckman) plate reader. Background fluorescence was determined from a PCR reaction that contained no template DNA. Following background subtraction, DNA concentration was determined by comparing fluorescence readings to those obtained with a standard curve using DNA of known concentration supplied with the dye. Extent of target amplification (e) is calculated as e = (ng DNA after PCR) ÷ (ng of target DNA input), and the number of template doublings during PCR (d) can be calculated as

3.3. Cloning of PCR Products

PCR reactions were purified in 96-well plate format by the addition of PEG 8000 and MgCl2 to final concentrations of 10% and 10 mM, respectively, directly to each well of the PCR plate using a multichannel pipettor. The plate was spun at 4000 rpm for 60 minutes at room temperature, and the supernatant was discarded. Pellets were washed two times with cold isopropanol, air-dried, and resuspended in 25 μL TE (pH 8). This protocol resulted in excellent yields (50–75%) of PCR products, with no products <300 bp, as judged by gel electrophoresis. Purified PCR products were cloned into a pDONR223 vector (a generous gift of Drs. Dominic Esposito and Jim Hartley, NCI, Frederick, MD) using BP Clonase II (Invitrogen/Life Technologies). Clonase reactions were assembled using a multichannel pipettor in 96-well PCR plates in a 5 μL volume and contained 75 ng pDONR223, 1 μL purified PCR product (typically 50–150 ng DNA), and 1 μL BP Clonase II. Sealed plates were incubated at least 16 hours at 25°C, and 1 μL of each reaction was immediately (no proteinase K treatment) used to transform either 25 μL or 50 μL of competent TOP10 cells (Invitrogen/Life Technologies). Following heat shock and recovery, following addition of 250 μL of SOC media, 100 μL of cells was plated on LB plates containing 50 mg/mL spectinomycin. Equivalent numbers of colonies were observed in transformations using 25 μL or 50 μL of frozen competent cells, and control BP reactions lacking BP Clonase II or PCR product resulted in no transformants.

3.4. Screening of Transformants

Three colonies from each transformation plate were picked and cultured in 96-well plates (Costar 3788) sealed with gas-permeable membrane, with each colony incubated in 150 mL of LB media with 50 mg/mL spectinomycin and 10% glycerol. After overnight incubation at 37°C (no shaking), 1 μL of each culture was used to screen by colony PCR for the presence of insert with expected size. Colony PCR reactions (25 mL) used the same primers used for cloning at a final concentration of 0.1 mM each, with 30 amplification cycles as described above, with GoTaq polymerase (Promega). Reactions were analyzed by agarose gel electrophoresis, and the presence of a band at or near the expected size was scored as a “hit.” The number of hits (0–3) for each target was determined, and an average number of hits per target for each plate were determined and used as a measure of Clonase reaction efficiency.

3.5. Clone Sequencing

In cases where Clonase efficiency values were >66%, average of at least 2 hits out of 3 colonies screened, the entire liquid culture plate was replicated with a 96-pin replicator onto an agar plate with the same dimensions as a 96-well plate. The plate was immediately submitted to an outside vendor (Quintarabio, Berkeley, CA), and after growth overnight sequencing was performed on amplified DNA from each clone. If Clonase efficiency values were <66% (Taq and Pfu polymerase reactions), a rearray step was added, using a Qpix2 colony picking robot (Genetix) to maximize the number of clones with correct-size insert on one plate. For comparing sequencing results using cells versus miniprep DNA, one plate of colonies picked from a Taq cloning reaction was replicated into a 96-well deep well plate with 800 mL media per well and grown overnight with shaking at 300 rpm. Cells were pelleted, and DNA was prepared using a Qiaprep 96 Turbo Miniprep Kit (Qiagen). Eluted DNA was submitted directly for sequencing.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgments

This work was part of the DOE Joint BioEnergy Institute (http://www.jbei.org) supported by the U.S. Department of Energy, Office of Science, Office of Biological and Environmental Research, through Contract DE-AC02-05CH11231 between Lawrence Berkeley National Laboratory and the U.S. Department of Energy. The authors would like to thank Drs. Dominic Esposito and Jim Hartley (NCI, Frederick, MD) for the gift of pDONR223 DNA, Huu M. Tran (JBEI/Sandia National Laboratories) for assistance with the laboratory automation, Drs. Richard Shan and Sue Zhao (Quintara Biosciences, Berkeley, CA) for DNA sequencing and analysis, and Dr. Nathan J. Hillson for critical reading of the paper.

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Copyright

Copyright © 2014 Peter McInerney et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


MATERIALS AND METHODS

Replication-cycle reaction (RCR)

A 5 × replication-cycle reaction (RCR) buffer (100 mM Tris-HCl pH8.0, 750 mM potassium glutamate, 50 mM ammonium sulfate, 50 mM Mg(OAc)2, 40 mM dithiothreitol (DTT), 20 mM creatine phosphate, 5 mM each NTP, 0.5 mM each dNTP, 0.25 mg/ml yeast tRNA, and 1.25 mM NAD + ) and a 4 × Enzyme mix (2 mg/ml bovine serum albumin, 0.08 mg/ml creatine kinase, 0.1 mM adenosine triphosphate (ATP), 1.6 μM SSB4, 160 nM IHF2, 1.6 μM DnaG, 160 nM DnaN2, 20 nM Pol III*, 80 nM DnaB6C6, 400 nM DnaA, 40 nM RNaseH, 200 nM ligase, 200 nM Pol I, 200 nM gyrase (GyrA2B2), 20 nM Topo IV (ParC2E2), 200 nM Topo III and 200 nM RecQ) were prepared for the standard reaction, unless otherwise noted. The reaction mixture (final 10 μl, unless otherwise noted) included 5 × RCR buffer (2 μl) and 4 × Enzyme mix (2.5 μl) and was assembled on ice. After addition of oriC-containing circular DNA, the reaction was incubated at 30°C for the indicated times. [α- 32 P]dATP was included at 40–100 cpm/pmol of total deoxynucleotides when indicated. The reaction was stopped by addition of an equal volume of 2 × Stop buffer (50 mM Tris–HCl pH8.0, 50 mM ethylenediaminetetraacetic acid, 0.2% sodium dodecyl sulphate, 0.1 mg/ml proteinase K, 10% glycerol and 0.2% bromophenol blue), and further incubated at 37°C for 30 min followed by phenol–chloroform extraction. An aliquot (2 μl) was analyzed by 0.5% agarose-gel electrophoresis followed by SYBR Green I staining (Molecular Probes) or by phosphor imaging. The images were acquired with a Typhoon FLA 9500 (GE Healthcare). The 4 × Enzyme mix can be stored at −80°C after rapid freezing with liquid nitrogen and its activity is retained even after five freeze-thaw cycles. Protein purification is described in Supplementary Methods .

Measurement of [α- 32 P]dATP-incorporation

An aliquot of the RCR sample was treated with 10% trichloroacetic acid. Nucleotide incorporation into DNA was measured by liquid-scintillation counting of acid-insoluble material retained on Whatman GF/C glass microfiber filters.

Quantification of the number of circular DNA molecules by transformation into E. coli

RCR was performed with oriC circular DNA templates containing a kanamycin-resistance cassette. An aliquot (1 μl) of the RCR sample was directly subjected to chemical transformation into E. coli strain DH5α. The RCR sample was diluted if necessary. Kanamycin (25 μg/ml)-resistant colonies were grown at 37°C overnight on LB-agar plates and counted. To deduce the number of circular DNA molecules from the colony number, 1 ng (10 8 molecules) of pOri8 circular DNA was used for each transformation experiment as a quantitative standard, which typically gave ∼10 000 colonies.

Measurement of the rate of replication errors

In this equation, F is the fraction of colonies that are blue and m is the number of population doublings. The number of error-detectable sites (i.e. non-silent target sites) in the 3 kb lacZ gene was estimated to be 1000.

DNA assembly for cell-free cloning

A 2 × modified Gibson Assembly (mGA) mix (200 mM Tris–HCl pH7.5, 20 mM MgCl2, 0.4 mM each dNTP, 20 mM DTT, 10% PEG8000, 2 mM NAD + , 16 mU/μl T5 exonuclease, 50 mU/μl Phusion DNA polymerase) was prepared in-house, with some modifications as described previously ( 29). The reaction (final volume 5 μl) consisted of 2 × mGA mix (2.5 μl) and the indicated amounts of DNA fragments, and was assembled on ice and incubated at 50°C for 1 h. DNA fragments were PCR-amplified with PrimeSTAR HS DNA polymerase (Takara Bio) and purified with the Wizard SV PCR Clean Up System (Promega). A 1 kb oriC fragment was amplified with primers SUE654/SUE656 using the E. coli K-12 MG1655 genome as a template. A 3.3 kb lacZ fragment including the lac promoter was amplified with primers SUE594/SUE655 using the MG1655 genome as a template. The 4.6 kb parABC-Km cassette was amplified with primers SUE635/SUE637 using pETcocoKm as a template. Primer sequences are listed in Supplementary Table S1 . The assembly products before or after RCR were analyzed following transformation of E. coli strain NM554.

‘Pop-out’ preparation of oriC-containing circular DNA

pOri8 (9.5 kb), pOri80 (85 kb), pOri200 (205 kb), pMSR227 (205 kb) and pOriDif (12 kb) were prepared by a pop-out method. λ Red recombination was carried out in E. coli K-12 MG1655 strain containing pKD46, as described previously ( 30), except that MH005, instead of MG1655, was used for pMSR227. The Km cassette (1.2 kb) was PCR-amplified from pUC4K with primers SUE351/SUE352 for pOri8. The parABC-Km cassettes (4.8 kb) were PCR-amplified from pETcocoKm with primers SUE409/SUE410 for pOri80 and SUE411/SUE412 for pOri200 and pMSR227. The Km-oriC cassette (2.1 kb) was PCR-amplified from pETcocoKmOri with primers SUE507/SUE509 for pOriDif. Primer sequences are listed in Supplementary Table S1 . These cassettes contain 40 bp homologous arms targeting sites 4 kb upstream and downstream of the chromosomal oriC site (for pOri8), 40 kb upstream and downstream of oriC (for pOri80), 100 kb upstream and downstream of oriC (for pOri200 and pMSR227) and 4.2 kb upstream and 6 kb downstream of the chromosomal dif site (for pOriDif). After electroporation, kanamycin-resistant colonies were selected and successful generation of the oriC-containing plasmids was verified through colony PCR. The plasmids were then partially purified and used for transformation of E. coli strains DH5α (for pOri8), HST08 (Takara Bio) (for pOri80, pOri200 and pMSR227) or DH10B (Life Technologies) (for pOriDif). Transformants were selected for large-scale purification of the plasmids using the QIAfilter plasmid kit (Qiagen) (for pOri8 and pOriDif) or NucleoBond Xtra BAC Kit (Takara Bio) (for pOri80, pOri200 and pMSR227). In the MH005 strain, a MG1655 derivative, the dnaN gene was replaced with a mCherry-dnaN fusion at its endogenous locus.

Other DNA constructs

To construct M13ms9 (8 kb), the 0.42 kb oriC fragment was PCR-amplified from the MG1655 genome using primers SUE260/SUE261, followed by NcoI–NsiI digestion and cloning into a vector prepared from M13mp18 by PCR with primers SUE226/SUE227, followed by NcoI–NsiI digestion. To construct M13ms10 (8 kb), a 0.25 kb fragment containing the chromosomal terB region and a second inverted terB sequence was PCR-amplified from the MG1655 genome using primers SUE236/SUE238, followed by NheI–NsiI digestion and cloning into the XbaI–PstI sites of M13ms9. Duplex, replicative forms of phage were prepared in E. coli JM109 strain and purified using the QIAfilter plasmid kit (Qiagen).

pPKOZ (8.9 kb) was constructed using the cell-free cloning method described in Figure 6. After transformation of DH5α, the plasmid was purified with the QIAfilter plasmid kit. To construct pETcocoKm (11.3 kb), the 1.2 bp Km fragment was PCR-amplified from pUC4k using primers SUE296/SUE375, digested with PciI–PstI, and cloned into the PciI–NsiI sites of pETcoco-2 (Novagen). To construct pETcocoKmOri, a 1 kb oriC fragment was PCR-amplified from the MG1655 genome using primers SUE505/SUE506 and cloned into the NheI–AatII sites of pETcocoKm. Primer sequences are listed in Supplementary Table S1 .


PCR Cloning Method

PCR cloning differs from traditional cloning in that the DNA fragment of interest, and even the vector, can be amplified by the Polymerase Chain Reaction (PCR) and ligated together, without the use of restriction enzymes. PCR cloning is a rapid method for cloning genes, and is often used for projects that require higher throughput than traditional cloning methods can accommodate. It allows for the cloning of DNA fragments that are not available in large amounts.

Typically, a PCR reaction is performed to amplify the sequence of interest, and then it is joined to the vector via a blunt or single-base overhang ligation prior to transformation. Early PCR cloning often used Taq DNA Polymerase to amplify the gene. This results in a PCR product with a single template-independent base addition of an adenine (A) residue to the 3' end of the PCR product, through the normal action of the polymerase. These "A-tailed" products are then ligated to a complementary T-tailed vector using T4 DNA ligase, followed by transformation.

High-fidelity DNA polymerases are also now routinely used to amplify sequences with the PCR product containing no 3' extensions. The blunt-end fragments are joined to a plasmid vector through a typical ligation reaction or by the action of an "activated" vector that contains a covalently attached enzyme, typically Topoisomerse I, which facilitates the vector:insert joining. Some PCR cloning systems contain engineered "suicide" vectors that include a toxic gene into which the PCR product must be successfully ligated to allow propagation of the strain that takes up the recombinant molecule during transformation.

A typical drawback common to many PCR cloning methods is a dedicated vector that must be used. These vectors are typically sold by suppliers, like NEB, in a ready-to-use linearized format and can add significant expense to the total cost of cloning. Also, the use of specific vectors restricts the researcher's choice of antibiotic resistance, promoter identity, fusion partners, and other regulatory elements.

  • High efficiency, with dedicated vectors
  • Amenable to high throughput
  • Limited vector choices
  • Higher cost
  • Lack of sequence control at junction
  • Multi-fragment cloning is not straight forward
  • Directional cloning is difficult

PCR Cloning

Note that times are based on estimates for moving a gene from one plasmid to another. If the source for gene transfer is gDNA, add 2 hours to calculation for the traditional cloning method. Total time does not include transformation, isolation or analysis.

The Science Behind the Test for the COVID-19 Virus

Mayo Clinic's new test for the virus that causes COVID-19 is described in a recent news release as a PCR test. While most won't know what that means, PCR is a well-used tool in the laboratory and medical testing. Larry Pease, Ph.D., a Mayo Clinic immunologist and the Gordon H. and Violet Bartels Professor of Cellular Biology and Kyle Rodino, Ph.D., a clinical microbiologist, explain how this test works.

To start, PCR stands for a laboratory technique known as polymerase chain reaction. In this test, the goal is to selectively amplify trace amounts of genetic material, identifying specific parts of DNA. Just as a reminder, DNA is the genetic code that is present in every cell in the body. When a cell divides, it copies DNA, separating the two strands and then creating a new strand of DNA by copying the template. PCR mimics what normally happens in cells.

Mayo Clinic announces on March 12, 2020 that it has developed a test that can detect the SARS-CoV-2 virus in clinical samples. The SARS-CoV-2 virus causes COVID-19.

DNA is used because at the most discriminating level, the structure of DNA can tell you what organism is being looked at. In the case of humans, PCR can identify a person using his or her genetic signature. In the case of COVID-19, researchers have published more than 100 genomes collected from patients to identify key features of the virus that causes that disease, SARS-CoV-2.

PCR only works on DNA, and the COVID-19 virus uses RNA as its genetic code. RNA is similar to DNA, but only has a single strand. Fortunately, viral enzymes to convert RNA into DNA were discovered decades ago, and have been harnessed, along with PCR, to find unique signatures in RNA, too. In this case, PCR is referred to as reverse transcription PCR, or RT-PCR.

First a person with symptoms of COVID-19 calls his or her local health care provider and asks how to be evaluated. Remember: Call first. Don't go to your clinic or hospital without calling to find out the safest way to get tested. Once a patient arrives at a safe testing site, a sample is taken by the health care team. Usually that means a narrow swab is placed in a person's nose or mouth to collect cells from the back of the throat.

"An upper respiratory specimen, particularly a nasopharyngeal swab, is the most common sample collected to reliably detect the virus," says Dr. Rodino. "Some lower respiratory samples like sputum are also acceptable in some settings."

At the lab, the sample is processed so RNA is isolated and collected. Everything else is removed. The RNA is mixed with other ingredients: enzymes (DNA polymerase and reverse transcriptase), DNA building blocks, cofactors, probes and primers that recognize and bind to SARS-CoV-2.

Then the viral RNA is converted to a DNA copy, and that single copy is then converted into millions of copies using PCR that can be easily detected.

The process is as follows: Using heat and enzymes, converted viral DNA strands are forced apart. Short primers of DNA matching the complementary strand of the viral DNA template stick together, functioning as an artificial start site for DNA synthesis.

Chemical building blocks of DNA are added and joined together, extending the synthetic DNA primer to form a copy of the viral DNA template. A second primer made in the opposite orientation downstream of the first primer is also present in the reaction. This makes a copy that is complementary to the first synthesized strand.

After one round of DNA synthesis, the reaction mixture is heated to melt the two strands apart, generating two templates that can be amplified further in the next round. The copies accumulate, round by round, exponentially, so that millions and millions of copies are generated to be studied using conventional approaches.

Because the enzymes and chemicals added to the reaction tube are relatively heat-resistant — "The heat sensitive enzymes are isolated from thermal resistant bacteria from hot springs," says Dr. Pease — the reaction can proceed in an automated fashion of heating, cooling and DNA synthesis. It only takes hours to complete the assay and get the results.

If SARS-CoV-2 complementary DNA is present in the sample, the primers can copy the targeted regions. As they copy these regions, probes stuck to these new fragments release a visual signal that can be read by the instrument used in this process.

"The millions of copies amplify this signal so it can be easily detected as a positive result. If the virus is not present, the probes do not stick, there is no signal release and it is a negative result," explains Dr. Rodino.

This type of analysis is used for research and clinical lab testing. PCR can detect all types of bacteria, parasites, viruses and fungi, starting with DNA or RNA. While the principle and ingredients are similar, each use requires specific primers or probes to detect different organisms. That's why something for SARS-CoV-2 had to be developed from scratch. During development, these sorts of tests are tweaked to make sure they are very good at detecting the organism of interest (sensitive) and making sure the test does not show a positive result when the organism is not there (specific).

"The importance of the steps involved in PCR has been recognized by a series of Nobel prizes over decades," says Dr. Pease. "Medical science advances as a result of basic discoveries about the molecular basis of living systems, and this is one example of how these discoveries come together to solve an important problem in our lives."

Working as a research and clinical team, Mayo experts were able to roll out a PCR test for SARS-CoV-2 in a matter of weeks ― shaving down what typically takes months to years.