If no amplification products are obtained, what parameters should be considered first when troubleshooting?
Consider the following:
- First, ensure that all PCR components were included in the reactions. A positive control should always be included to ensure that each component is present and functional.
- If there were no problems with the experimental setup, increase the number of PCR cycles (3–5 cycles at a time), up to 40 cycles. Increasing the cycle number can overcome issues with a low-abundance template or template inaccessibility due to impurities in or poor priming efficiency of the primers.
- If increasing the cycle number does not improve results, the PCR conditions might be too stringent for the particular primer set or template. Consider modifying the PCR conditions as follows:
- Lower the annealing temperature in increments of 2 degrees.
- Increase the extension time.
- Increase the template amount. Refer to the guidelines provided with the enzyme to determine the optimal amount of template.
Consider these additional possible reasons for PCR failure:
- PCR inhibitors in the template sample
If PCR inhibitors are present, using diluted template may increase PCR efficiency. Alternatively, the template may need to be purified using a kit such as the NucleoSpin Gel and PCR Clean-up kit. If purifying the template is not a possibility, an enzyme that has a higher tolerance to impurities, such as Terra PCR Direct polymerase, may improve results.
- The template has >65% GC content
When amplifying from templates with high GC content, use an enzyme formulated for this condition. Visit our PCR selection guide to find an appropriate enzyme.
- Primers are not optimal
Check your primers carefully; redesign if necessary. Also, consider re-amplifying the primary PCR product using 10-fold dilutions (1:100 to 1:10,000) using nested primers.
When using PrimeSTAR HS DNA Polymerase, consider:
- Using an appropriate amount of template. If the template is human genomic DNA or a cDNA library, use no more than ~100 ng of the template in a 50-µl reaction mixture.
- Using an extension time of at least 1 min/kb.
- Increasing the concentration of the primers.
When using PrimeSTAR Max DNA Polymerase, consider:
- Adjusting the extension time if the reaction mixture contains excess template. If the amount of template exceeds 200 ng in a 50-µl reaction mixture, set the extension time between 30 sec/kb and 1 min/kb.
- Increasing the concentration of the primers.
When using SpeedSTAR HS DNA Polymerase, consider:
- Increasing the extension time. Although the standard extension time is 10 sec/kb, the extension time can be increased to ~0.5 min/kb for complex templates such as human genomic DNA.
If there are nonspecific amplification bands, what can be done to improve specificity?
All Takara Bio PCR polymerases
Primers are not specific.
Use BLAST alignment to determine if the 3' ends of the primers are complementary to sites other than the target site(s). Redesign primers if necessary or modify PCR conditions.
PCR conditions are not sufficiently stringent.
- Increase the annealing temperature in increments of 2 degrees.
- Use touchdown PCR.
- Use a two-step PCR protocol.
- Reduce the number of PCR cycles.
Too much template was used.
Reduce the amount by 2–5 fold.
PrimeSTAR HS and PrimeSTAR Max DNA polymerases
Annealing time is too long.
To achieve specific amplification, it is essential to use a short annealing time (5–15 sec) when performing three-step PCR.
PrimeSTAR GXL DNA polymerases
Primers have suboptimal Tm values.
To amplify targets <1 kb, design primers with Tm values >55°C, and use an annealing temperature of 60°C. If the primer Tm values are <55°C, try a shorter extension time between 5 and 10 sec/kb.
Takara Ex Taq and Takara LA Taq DNA polymerases
Nonspecific primer annealing at low temperatures.
The hot-start versions of these enzymes may improve results for some primers.
SpeedSTAR HS DNA Polymerase
Smearing of the PCR product bands on a gel.
Excessively long extension times may result in smearing. The general recommendation for extension time for this enzyme is 10–20 sec/kb. If PCR yield is low, try increasing the number of cycles by 5.
If PCR generates a smear after running the products on a gel, what can be done to improve the results?
First, determine the source of the smear using positive and negative (no template) controls. This can determine if the cause of the smear is contamination or overcycling, or if the smear results from poorly designed primers or suboptimal PCR conditions.
If the negative control is blank, there is no contamination. Instead, the PCR conditions will need to be optimized; consider the following when adjusting the PCR conditions:
- Reduce the amount of template.
- Increase the annealing temperature.
- Use touchdown PCR.
- Reduce the number of PCR cycles.
- Redesign the primers.
- Use nested primers.
- Re-amplify the product. (A small plug of the gel can be removed with a micropipette tip, and the DNA can be recovered by adding the plug to 200 µl of water and then incubating at 37°C. 5 µl of this solution can be used as PCR template for re-amplification.)
If the negative control is also smeared, there is contamination. You will need to determine the source of this contamination. It may be necessary to replace PCR reagents and to decontaminate pipettes and your workstation (see questions below for more information on contamination).
What are some sources of PCR contamination?
There are four main sources of PCR contamination:
- The most common source of contamination is PCR product from previous amplifications (called "carryover contamination"). When large amounts of PCR product (1012 molecules) are generated repeatedly over a period of time, the potential for contamination increases.
- Another source of contamination is cloned DNA previously handled in the laboratory.
- Sample-to-sample contamination can also occur. This source of contamination is most likely to be found in samples that require extensive processing prior to amplification.
- Reactions can also be contaminated with exogenous DNA in the environment, including DNA present on laboratory equipment and in reagents used for DNA extraction and PCR.
How can contamination be avoided?
The sensitivity of PCR requires that samples are not contaminated with any exogenous DNA or any previously amplified products from the laboratory environment. We recommend that distinct areas are used for sample preparation, PCR setup, and post-PCR analysis.
A laminar flow cabinet equipped with a UV lamp is recommended for preparing reaction mixtures. Two stations should be established that are physically separated from each other.
- Establish a "pre-PCR area" that is for PCR reaction setup only. No items from the "post-PCR area" should be introduced into this area; this includes items such as notebooks and pens.
- Establish a "post-PCR area" that is used for PCR, purifying PCR-amplified DNA, measuring DNA concentration, running agarose gels, and analyzing PCR products.
Equipment should also be restricted to these areas. The PCR machine and electrophoresis apparatus should be located in the post-PCR area. Having pipettes and pipette tips with aerosol filters dedicated for DNA sample and reaction mixture preparation only is strongly recommended. Additional recommendations include:
- Having separate sets of pipettes and pipette tips, lab coats, glove boxes, and waste baskets for the pre-PCR and post-PCR areas.
- Labeling pre- and post-PCR items, so they are not removed from their designated work area.
- Following the golden rule of PCR: NEVER bring any reagents, equipment, or pipettes used in a post-PCR area back to the pre-PCR area.
- Preparing and storing reagents for PCR separately and using them solely for their designated purpose. Reagents should be aliquoted in small portions and stored in designated areas depending on if they are used for pre-PCR or post-PCR applications. The aliquots should be stored separately from other DNA samples.
A control reaction that omits template DNA should always be performed to confirm the absence of contamination. In addition, the number of PCR cycles should be kept to a minimum, as highly sensitive assays are more prone to the effects of contamination.
How can I decontaminate if I have PCR contamination?
- Leave pipettes under UV light in the cell culture hood overnight. UV irradiation promotes cross-linking of thymidine residues, damaging residual DNA.
- Spray workstations/equipment/pipettes with 10% bleach and then wipe clean.
- Change workstations; move the pre-PCR area to another pre-cleaned location.
- Do not use any instruments or pipettes you have used before.
What can I do if the PCR generates errors?
To avoid errors during PCR, we recommend using a high-fidelity enzyme (see selection guide). In addition, be sure to avoid the following:
Overcycling PCR reactions often:
- Changes the pH of the reaction in a manner that destabilizes DNA.
- Increases the amount of PCR product, thereby reducing the efficiency of the polymerase and promoting errors.
- Decreases the amount of dNTPs, thereby increasing the likelihood of base misincorporation due to the unbalanced dNTP concentration. (If using Takara Bio's PCR enzymes, the dNTP concentration is optimized to 200 nM; increasing dNTP concentration leads to misincorporation.)
- Causes accumulation of single-stranded and double-stranded DNA.
- High Mg2+ concentration
Mg2+ concentration ranges from 1–5 mM. Using a high Mg2+ concentration may increase yield, but it might also impact the proofreading activity of enzymes. However, the Mg2+ concentration should always be higher than the dNTP concentration.
- Template DNA damage
Limit UV exposure time when analyzing or excising PCR products from gels.
Recommended enzymes for specific applications and conditions
Which polymerases provide the highest fidelity?
PrimeSTAR polymerases provide better fidelity than Pfu DNA polymerase, which is widely considered to be a high-fidelity enzyme. PrimeSTAR Max DNA Polymerase has the highest fidelity; when this enzyme was used to amplify the entire pUC119 plasmid, sequence analysis detected only four mutations out of 370,656 total bases sequenced (an error rate of 0.0010%). In addition, compared to other enzymes, PrimeSTAR Max DNA Polymerase replicates repeat sequences with markedly better fidelity and exhibits a lower rate of template exchange (formation of chimeric molecules) with analogous sequences.
Which polymerases are optimal for amplifying GC-rich target sequences?
Consider the following enzymes:
- First choice: PrimeSTAR GXL DNA Polymerase is the most effective enzyme for GC-rich templates, such as bacterial genomic DNA. It facilitates high-fidelity amplifications with very few errors. PrimeSTAR GXL DNA Polymerase has been used successfully to amplify a region with ~70% GC content in a standard reaction using the buffer provided with the enzyme.
- Second choice: Advantage GC 2 Polymerase Mix is recommended for complex templates containing up to 90% GC content. This polymerase is suitable for fragments up to 6 kb. Advantage GC 2 polymerase used with DMSO and GC-Melt reagent allows amplification of virtually all GC-rich sequences.
- Third choice: for GC-rich targets that have rigid structures and are difficult to amplify with PrimeSTAR GXL DNA Polymerase, use TaKaRa LA Taq DNA Polymerase with GC Buffer. Try GC Buffer I first; this buffer facilitates the amplification of long products. GC Buffer II is effective for templates with complex higher-order structures, although this buffer is more effective for amplification of shorter products.
Which polymerases are suggested for long-range PCR?
PrimeSTAR GXL DNA Polymerase is recommended when both length (>6 kb) and fidelity are factors. Amplification of 30-kb products from human genomic DNA templates has been accomplished with this enzyme. Takara LA Taq DNA polymerase can also be used for long-range PCR; this enzyme is recommended when length and robust amplification are priorities.
Which polymerases are compatible with fast PCR?
We have several PCR enzymes that can be used for fast reactions.
- Speed and fidelity—PrimeSTAR Max DNA Polymerase contains a proprietary elongation factor and exhibits excellent priming efficiency, allowing extension times as short as 5 sec/kb and an annealing time of only 5 sec. This enzyme is recommended for cloning and expression studies.
- High-throughput applications and fast colony PCR—SapphireAmp Fast PCR Master Mix is an economical choice for high-throughput projects. This 2X enzyme premix includes a high-speed polymerase, optimized buffer, dNTP mixture, gel loading dye (blue), and a density reagent. Since it requires an extension time of only 10 sec/kb, colony PCR reactions can be completed in <1 hr for inserts up to 1 kb.
- SNP genotyping and fast long-range PCR—SpeedSTAR HS DNA Polymerase is highly efficient and can reliably perform PCR amplifications with extension times between 10 and 20 sec/kb.
Which polymerases are best for amplifying AT-rich target sequences?
TaKaRa Ex Taq DNA Polymerase and PrimeSTAR GXL DNA Polymerase are effective for amplifying AT-rich target sequences, such as genomic DNA containing introns or AT-rich mitochondrial DNA. We recommend PrimeSTAR GXL DNA Polymerase for amplifying AT-rich templates with high accuracy. In contrast to other PCR enzymes, PrimeSTAR GXL polymerase can amplify targets containing >60% AT content using a standard PCR protocol and the reaction buffer provided.
Note: PrimeSTAR GXL DNA Polymerase cannot be used to amplify bisulfite-treated DNA or other uracil-containing templates. For this application, try TaKaRa EpiTaq HS (for bisulfite-treated DNA).
Which polymerases can be used to amplify bisulfite-treated DNA?
Several choices are available:
- TaKaRa EpiTaq HS (for bisulfite-treated DNA) is optimized for PCR amplification using bisulfite-treated DNA templates that contain uracil. This enzyme includes a hot-start antibody and is designed for use during methylation analysis, including COBRA and bisulfite-sequencing analyses.
- The EpiScope MSP Kit is designed specifically for methylation-specific PCR (MSP) assays.
- TaKaRa Taq DNA Polymerase and TaKaRa Taq DNA Polymerase Hot Start Version, both of which lack 3'→5' exonuclease activity, may be used in some instances.
Which polymerases have the highest tolerance for PCR-inhibiting agents?
TaKaRa Ex Taq DNA Polymerase is exceptionally robust, even in the presence of PCR inhibitors such as polyphenols found in crude DNA extracts from plant tissue. PrimeSTAR GXL DNA Polymerase is recommended when high fidelity is also required. Although PrimeSTAR GXL polymerase generally produces satisfactory results with the standard protocol, doubling the enzyme concentration may improve results if very high concentrations of inhibitors are present.
Alternatively, Terra PCR Direct Polymerase Mix allows direct amplification from crude samples that may contain high levels of PCR inhibitors. Terra PCR Direct polymerase can efficiently amplify a wide range of targets, including GC- and AT-rich targets. This enzyme can also be used for direct PCR from blood samples.
In cases where amplification products cannot be produced with these enzymes, it may be necessary to purify the template DNA.
Which polymerases are suitable for amplifying DNA prepared from paraffin sections?
The Terra PCR Direct FFPE Kit can be used for crude DNA preparation and direct PCR amplification from paraffin-embedded tissue sections.
If it is necessary to extract DNA from paraffin-embedded tissue first, TaKaRa DEXPAT Easy and TaKaRa DEXPAT Reagent enable quick, efficient DNA preparation, even from slides or samples that have been stored for years.
Which polymerases are recommended when many samples will be analyzed by gel electrophoresis after PCR (e.g., genotyping screens)?
We recommend EmeraldAmp GT PCR Master Mix. This PCR master mix contains a green gel loading dye and density reagent, making it easy to prepare PCR reaction mixtures that can be directly loaded on an agarose gel for electrophoresis after PCR. EmeraldAmp GT PCR Master Mix can amplify targets up to 10 kb in length, including targets that are GC- or AT-rich. Additionally, PCR products generated with EmeraldAmp GT PCR Master Mix can be used directly for restriction enzyme digestion, sequencing, or TA-cloning without purification.
Which polymerases are recommended for colony PCR?
We recommend SapphireAmp Fast PCR Master Mix for colony PCR. This enzyme premix can tolerate a substantial presence of bacterial nucleic acids. Tracking dye and density reagent are included in the master mix, allowing the PCR reaction products to be loaded directly on an agarose gel for electrophoresis.
What precautions should be taken when using inosine-containing primers?
Inosine-containing primers should not be used with PCR enzymes that have 3'→5' exonuclease activity (e.g., PrimeSTAR HS DNA Polymerase, PrimeSTAR Max DNA Polymerase, PrimeSTAR GXL DNA Polymerase, TaKaRa Ex Taq DNA Polymerase, or Takara LA Taq DNA polymerases), nor with Terra PCR Direct polymerase. When using one of the compatible Takara Taq PCR enzymes for degenerate PCR, we recommend using a mixture of degenerate primers with A, T, G, or C at the desired position(s) rather than inosine-containing primers.
Which polymerases have the highest sensitivity?
Which polymerases do you recommend for direct amplification from tissue?
We recommend Terra PCR Direct Polymerase Mix for amplifying targets from crude extracts or directly from tissues. The recommended amounts of various tissues that should be used for direct PCR are listed below:
- Treated blood: ≤5 µl
- Mouse tail: ≤1 mm
- Mouse ear: ≤1.5 mm2
- Plant leaf: ≤1.2 mm (diameter)
- Paraffin-embedded tissue section: ≤1–1.5 cm2
Which polymerases do you recommend for multiplex PCR?
We recommend TaKaRa Taq DNA Polymerase or Titanium Taq DNA Polymerase for multiplex PCR. View our tech note for more information about using Titanium Taq DNA Polymerase in multiplex PCR for high-throughput genotyping.
For each Takara Bio polymerase, what is the structure at the ends of the amplified fragments, and what are the most suitable cloning methods?
- Enzymes in the PrimeSTAR series
These enzymes exhibit substantial 3'→5' exonuclease activity and primarily generate amplification products with blunt ends. Therefore, we recommend using the Mighty Cloning Reagent Set (Blunt End) for blunt-end cloning.
- Taq and Terra enzymes
Takara Taq, Takara Ex Taq, Takara LA Taq, SpeedSTAR HS, EmeraldAmp, SapphireAmp Fast, and Terra PCR DNA polymerases primarily yield amplification products containing 3'-dA overhangs that can be directly used for TA-cloning. Blunt-end cloning is also possible using the Mighty Cloning Reagent Set (Blunt End).
- All Takara Bio DNA polymerases
The fragment terminal structure after amplification with a particular polymerase is indicated in the table below. Some polymerases generate a mixture of blunt-end and A-overhang products; other polymerases generate only blunt-end or A-overhang products.
|Takara Taq DNA polymerases|
|Takara Ex Taq DNA polymerases|
|Takara LA Taq DNA polymerases|
|Titanium Taq DNA Polymerase|
|Advantage 2 Polymerase Mix|
|Advantage GC 2 Polymerase Mix|
|Advantage HF 2 Polymerase Mix|
|EmeraldAmp GT PCR Master Mix|
|EmeraldAmp MAX HS PCR Master Mix|
|SapphireAmp Fast PCR Master Mix|
|High Yield PCR EcoDry Premix|
|High Fidelity PCR EcoDry Premix|
|Terra PCR Direct Polymerase Mix|
|SpeedSTAR HS DNA Polymerase|
|PrimeSTAR HS DNA Polymerase|
|PrimeSTAR GXL DNA Polymerase|
|PrimeSTAR Max DNA Polymerase|
|CloneAmp HiFi PCR Premix|
|SeqAmp DNA Polymerase|
|TaKaRa Z-Taq DNA Polymerase|
|e2TAK DNA Polymerase|
|PerfectShot Ex Taq DNA Polymerase|
General PCR guidelines
What parameters do I need to consider when designing primers?
Primer design is the most important factor in determining the success or failure of PCR reactions. There are two major considerations for primer design: specificity and efficiency.
- Specificity is determined by the frequency of mispriming events. Primers with poor specificity tend to produce undesired amplicons.
- Efficiency is defined as the ability of primers to amplify a product with a two-fold increase per cycle to the theoretical optimum.
The following tables provide guidelines for primer design.
|Length||The optimal length of primers is about 24 or 25 bases. However, length can be between 21 and 28 bases if the melting temperature (Tm) needs to be adjusted. When amplifying long DNA fragments (≥10 kb), 25- to 35-mer primers may provide better results.|
|GC content||The GC content (the number of Gs and Cs in the primer as a proportion of the total number of bases) should be 40–60%.|
|3' end||Having four G and/or C bases at the 3' end might be useful if the primer length is short (the bases provide "a clamping effect"), especially for universal primers, which are typically used for amplifying all cDNA or gDNA in a sample. However, adding these bases may increase non-specific priming events for gene-specific primers.|
|Tm||If possible, design primers with a melting temperature of 68–70°C. While this is not absolutely necessary, using stringent PCR conditions (e.g., "touchdown PCR" and "two-step PCR") can enhance primer specificity.|
|Sequence specificity||Primers should be specific to your gene of interest. A BLAST search can be used to find regions of homology. Note: PCR yield often depends on the 3' hexamer of the PCR primer. Primers that form a strong stable duplex actually reduce the amplification efficiency.|
|Repeats||The maximum number of di-nucleotide repeats in a primer is four (e.g., ATATATAT).|
|Runs||Avoid long runs of a single base (more than three) as this can cause primer slippage and contribute to mispriming.|
|Complementary 3' ends||Forward and reverse primers should not anneal to each other and so should not have complementary G or C stretches (>4 contiguous bases).|
|Self-complementary 3' ends||Self-complementarity (e.g., within the forward primer) can lead to hairpin formation. A hairpin structure can form with just four G/C base pairs in the stem and three bases in the loop.|
How do I calculate the melting temperature (Tm) of primers?
The primer melting temperature (Tm) is the estimate of DNA-DNA hybrid stability. Knowing the Tm is critical for determining an appropriate annealing temperature (Ta). A Ta that is too high will result in insufficient primer-template hybridization, leading to low PCR product yield. A Ta that is too low may lead to non-specific product amplification.
Calculation of the Tm of primers shorter than 20 bases can be performed using the Wallace rule:
Tm = 2°C (A+T) + 4°C (G+C)
For accurate estimation of the Tm of primers longer than 20 bases, we recommend using free primer design software such as Primer3.
What primer concentration should be used for PCR?
The final concentration of each primer should be between 0.1 and 0.5 µM. A stock solution of each primer is typically 10–20 µM.
- For PCR amplicons less than 10 kb, 0.2 µM produces satisfactory results.
- For amplification of long targets (~17 kb) with Takara LA Taq DNA polymerases or Takara Ex Taq DNA polymerases, the primer concentration can be increased up to 1 µM.
- The recommended primer concentration for high-yield polymerases such as Advantage 2 DNA polymerases and Titanium Taq DNA polymerases is 0.4 µM.
Primer concentrations that are too high increase the chance of mispriming, which may result in nonspecific amplification. Primer concentrations that are limiting can result in extremely inefficient amplification.
How should oligos be purified for PCR?
Standard desalted primers are satisfactory for most PCR applications.
What is the optimal amount of DNA template that should be used for PCR?
The optimal amount of template required depends on the complexity of the template and the copy number of the target sequence. Approximately 104 copies of the target DNA sequence are required to detect the amplification product in 25–30 PCR cycles.
- Typically, 1 µg of human genomic DNA contains 3.04 x 105 molecules of DNA. For most PCR applications, 30–100 ng of human genomic DNA is sufficient. High-copy targets, such as housekeeping genes, require only 10 ng of template. Template amounts for higher-complexity templates range between 10 ng and 500 ng.
- Typically, 1 µg of E. coli genomic DNA contains 2 x 108 molecules of DNA; therefore, the recommended amount of template is between 100 pg and 1 ng.
- Typically, 1 µg of lambda DNA contains 1.9 x 1010 molecules of DNA; therefore, the template input can be as little as 100 pg.
- The amount of cDNA template depends on the copy number of the target. cDNA input is typically described in terms of equivalent RNA input. The amount of cDNA in a PCR reaction can be as little as 10 pg (RNA equivalent).
It is important to note that not all polymerases can tolerate excessive amounts of template. For samples containing excess template (up to 1 µg), we recommend PrimeSTAR GXL DNA Polymerase.
What are the critical factors for amplification of long genomic targets?
DNA integrity is critical for amplification of long targets. DNA damage—such as DNA breakage during DNA isolation or DNA depurination at elevated temperatures and low pH—results in a greater amount of partial products and decreased overall yield. DNA damage can also occur in acidic conditions; therefore, avoid using water for resuspending DNA templates. DNA is most stable at pH 7–8 or in buffered solutions.
- Denaturation time should be kept to a minimum to decrease depurination events.
- Use touchdown PCR; start at a higher annealing temperature and reduce by two degrees per cycle for several cycles.
- Design primers with melting temperatures (Tm) above 68°C.
We offer several PCR polymerases optimized for long-range PCR. Takara LA Taq DNA polymerase, TaKaRa LA Taq Polymerase with GC Buffer, and PrimeSTAR GXL DNA Polymerase are recommended depending on the GC content and size of the target(s).
How do I determine if a template is GC rich?
The GC ratio varies across the genome. Templates with >65% GC content are considered GC rich. GC-rich regions of the genome are mostly concentrated in regulatory regions, including promoters, enhancers, and cis-regulatory elements. GC-rich tracts tend to form inverted repeats, or hairpin structures, that may not melt during the annealing step of PCR. Therefore, amplification of GC-rich templates is hindered by inefficient separation of the two DNA strands. This results in truncated amplicons due to premature termination of polymerase extension.
What are the critical factors for amplification of GC-rich templates?
- Use higher denaturation temperatures (e.g., 98°C as opposed to 94°C or 95°C) to allow complete denaturation of the template.
- Keep annealing times for GC-rich templates as short as possible.
- Use primers with a higher Tm (>68°C), because annealing can occur at a higher temperature.
Use a polymerase optimized for amplification of GC-rich sequences. To find an enzyme, visit our selection guide.
Can DMSO be added to improve amplification of GC-rich templates?
We have heard from customers that improved amplification of GC-rich templates was obtained by adding DMSO to reactions using PrimeSTAR MAX DNA Polymerase or CloneAmp HiFi PCR Premix. The recommended concentration of DMSO is between 2.5% and 5%.
How can I optimize PCR conditions for AT-rich templates?
Some templates may have long AT-rich stretches that are hard to amplify under standard reaction conditions. The Plasmodium falciparum genome is about 80% AT, and regions flanking genes are often AT rich.
The advantage of having AT-rich templates is that a lower extension temperature can be used. For certain templates with AT content >80–85%, the extension temperature can be lowered from 72°C to 65–60°C. DNA replication at this reduced temperature appears to be reliable (Su et al. 1996).
Su, X. Z., et al. Reduced Extension Temperatures Required for PCR Amplification of Extremely A+T-rich DNA. Nucl Acids Res. 24, 1574–1575 (1996).
What is the role of magnesium in PCR, and what is the optimal concentration?
Magnesium is a required cofactor for thermostable DNA polymerases and is important for successful amplification. Without adequate free Mg2+, PCR polymerases are not active. In contrast, excess free Mg2+ reduces enzyme fidelity and may increase nonspecific amplification. A number of factors can affect the amount of free Mg2+ in a reaction, including DNA template concentration, chelating agents in the sample (e.g., EDTA or citrate), dNTP concentration, and the presence of proteins.
- Some polymerases (e.g., Takara Ex Taq DNA polymerases and Takara LA Taq DNA polymerases) are supplied with a magnesium-free reaction buffer and a separate tube of 25 mM MgCl2. For these enzymes, you can optimize the Mg2+ concentration for each reaction.
- Titanium Taq DNA polymerases and Advantage 2 DNA polymerases are magnesium-tolerant polymerases that are supplied with buffers containing 3.5 mM of MgCl2.
- The final concentration of Mg2+ for PrimeSTAR GXL DNA Polymerase and PrimeSTAR MAX DNA Polymerase reactions is 1 mM; this concentration increases fidelity for these enzymes.
What is the role of salt in PCR reactions?
Successful PCR requires that the DNA duplex separates during the denaturation step and that primers anneal to the denatured DNA. Salt neutralizes the negative charges on the phosphate backbone of DNA, stabilizing double-stranded DNA by offsetting negative charges that would otherwise repel one another. Potassium chloride (KCl) is normally used in PCR amplifications at a final concentration of 50 mM. To improve amplification of DNA fragments, especially fragments between 100 and 1,000 bp, a KCl concentration of 70–100 mM is recommended. For amplification of longer products, a lower salt concentration appears to be more effective, whereas amplification of shorter products occurs optimally with higher salt concentrations. This effect is likely because high salt concentration preferentially permits denaturation of short DNA molecules over long DNA molecules.
It is important to note that a salt concentration above 50 mM can inhibit Taq polymerases.
When optimizing PCR conditions, which conditions are particularly important?
Initial denaturation step
Preheating is sometimes required to denature complex templates (e.g., genomic DNA); 94°C for 1 min is sufficient for denaturation. Excessive heat treatment may lead to enzyme inactivation.
- For Terra PCR Direct Polymerase Mix, which is used for direct PCR amplification from tissue without DNA extraction and purification, preheating at 98°C for 2 min is required.
- For Takara LA Taq DNA polymerases and Advantage GC2 DNA polymerases, an initial denaturation step is required.
- PrimeSTAR enzymes do not require preheating for enzyme activation.
Denaturing conditions should be selected by considering the thermal cycler model that will be used. A general guideline is 94–95°C for 30 sec or 98°C for 10 sec.
If using a heat-resistant enzyme, such as one of the PrimeSTAR polymerases, we recommend a denaturation step of short duration and high temperature (i.e., 5–10 sec at 98°C).
Denaturation at an excessively high temperature or for too long may result in loss of enzyme activity and/or damage to long templates.
The annealing step should be adjusted for each primer set; the annealing temperature depends directly on the Tm of primers. Using annealing temperatures that are too low may result in mispriming and nonspecific amplification, leading to low yields of the desired product.
Amplification efficiency and specificity can be improved by adjusting the annealing temperature according to the primer's Tm or by performing two-step PCR.
- For Taq enzymes, the recommended annealing time is 30 sec.
- Enzymes in the PrimeSTAR series have excellent priming efficiency. Therefore, it is important to use a short annealing time of 5–15 sec. Excessively long annealing times may lead to mispriming-induced nonspecific amplification.
- When amplifying short sequences smaller than 1 kb, a three-step PCR protocol is recommended. For GC-rich targets or amplifications of long sequences (>10 kb), a two-step PCR protocol is recommended.
In general, an extension time of 1 min/kb is recommended. When using the high-speed enzymes SpeedSTAR HS DNA Polymerase or SapphireAmp Fast PCR Master Mix, use a reaction rate of 10 sec/kb of amplified product (i.e., 10 sec for a 1-kb product, 20 sec for a 2-kb product, etc.).
PrimeSTAR Max DNA Polymerase and PrimeSTAR GXL DNA Polymerase contain a proprietary elongation factor and allow for high-speed reactions at 5–20 sec/kb. If using these enzymes with samples containing excess template, an elongation time of 1 min/kb should be used.
Should I use a three-step or a two-step PCR protocol?
Three-step PCR includes denaturation, annealing, and extension steps. This type of protocol should be used when the Tm of the primers is lower than the extension temperature or is less than 68°C.
If the melting temperature of the primer (Tm) is close to the extension temperature (72°C) or a few degrees lower, consider using a two-step PCR protocol that includes a denaturation step and a combined annealing/extension step. With this protocol, the annealing temperature should not exceed the extension temperature.
Which extension temperature should I use, 68°C or 72°C?
A 68°C extension temperature is preferred for two-step PCR and when amplifying longer templates (>4 kb). This lower extension temperature dramatically improves yields of longer amplification products by reducing the depurination rate that influences amplification.
72°C should be used as the extension temperature when performing three-step standard PCR and for amplification of short fragments (<4 kb).
How are PCR polymerases shipped, and how should they be stored and handled after receipt?
Takara Bio's PCR polymerases are shipped on dry ice. All enzymes shipped on dry ice will be frozen, but this freezing process is gradual and one freeze-thaw cycle will not reduce enzyme activity. Before opening a new tube of enzyme, spin it briefly to collect the contents at the bottom of the tube.
Polymerases should be stored at –20°C in a non-frost-free freezer (an acceptable range is between –15°C and –23°C). Long-term storage (longer than a week) at –70°C or –80°C is not recommended. Enzymes should never be allowed to reach room temperature.
How should lyophilized PCR premixes be stored?
Lyophilized PCR premixes, sealed in the original foil pouch, should be stored at room temperature (20–22°C). Any unused product should also be stored at room temperature (20–22°C), either in the original foil pouch re-sealed with desiccant, or in a desiccator.
Are there any special considerations when handling PCR enzymes?
To avoid contamination, gloves should be worn when handling enzyme tubes, and fingers should be kept away from the tube opening. It is imperative to use a new, clean pipette tip every time you draw from a stock tube of enzyme.
When pipetting enzyme from a stock tube, place the end of the tip just far enough into the liquid to obtain the desired volume. A pipette tip should not be plunged all the way into the enzyme solution as the outside of the tip will become covered with enzyme, preventing accurate measurement and wasting enzyme.
Note: The retention of liquids to polypropylene tips varies with different types of solutions. Pipette tips lose their precision when liquid is drawn more than once. Low-retention pipette tips are recommended for use with viscous solutions, such as those containing glycerol.
What is meant by polymerase fidelity? What applications require a high-fidelity polymerase?
The fidelity of a DNA polymerase refers to its ability to accurately replicate a template, or to add the correct nucleotides starting at the 3' end of the primer. The rate of base misincorporation is known as the error rate. PCR polymerases with proofreading activity possess 3'→5' exonuclease activity that can excise incorrectly incorporated nucleotides and replace them with the correct nucleotides.
High-fidelity polymerases are recommended for gene cloning, protein expression, structure-function studies of proteins, cDNA library construction, and next-generation sequencing. To select a high-fidelity polymerase, see our PCR selection guide.
How can I compare error rates of different high-fidelity polymerases?
Error rates reported by vendors for polymerases cannot always be directly compared, as different methods are used to measure fidelity. These methods include:
- Blue-white screening
This approach is based on phenotypic changes and is widely used since it is fast, relatively simple, and cost effective. The original method for blue-white screening, known as the Kunkel method (Kunkel and Tindall 1987), is based on α-complementation of the lacZα gene that restores β-galactosidase enzyme activity and allows production of a blue color. With this method, colonies derived from lacZα PCR products containing single-nucleotide errors or frameshift mutations typically have a white color, while clones derived from error-free amplicons generate blue colonies.
- Sequencing approach
This approach utilizes Sanger sequencing of individual colonies after PCR. The blue-white screening approach can quickly measure polymerase fidelity, however it is not as accurate as the sequencing approach. The blue-white method will not detect silent mutations, single-nucleotide substitutions that do not affect translation. The sequencing method can detect all mutations, and thus is more accurate.
Kunkel, T. A. and Tindall, K. R., Fidelity of DNA synthesis by the Thermus aquaticus DNA polymerase. Biochemistry 27, 6008–6013 (1987).
PrimeSTAR Max and PrimeSTAR GXL DNA polymerases have very high fidelity; how was fidelity measured for these enzymes?
The fidelities of the PrimeSTAR polymerases were measured by Sanger sequencing of individual colonies after PCR, as described below:
- Ten arbitrarily selected GC-rich regions of Thermus thermophilus HB8 genomic DNA were amplified.
- PCR products were cloned into a plasmid vector.
- Multiple clones were selected for each respective amplification product, and the PCR insert was sequenced.
What factors are critical for multiplex PCR?
All primer pairs used in multiplex PCR should have similar priming efficiencies for their target DNA. This can be achieved by using primers with nearly identical optimum annealing temperatures.
When designing primers, pay special attention the following parameters:
- Homology with the target nucleic acid sequence
- GC content
- Primer homology (primers should not have homology either internally or with one another, especially at the 3' ends)
What is nested PCR?
Nested PCR is a method that involves re-amplification to improve PCR results. Nested PCR involves designing a new forward-nested (FN) or reverse-nested (RN) primer that is internal to the original primer and can pair with the original partner primer. A very small amount of the primary PCR product is used as a template for PCR with nested primers.
Nested PCR frequently leads to improved yield of the desired PCR product by:
- Eliminating extra bands that may have been present in the initial PCR
- Producing a robust band that may have been weak or invisible in the initial PCR
It is important to note that only a very small amount of the primary product should be used in nested PCR because this template has very low sequence complexity. To start, the primary PCR product can be diluted 1:100, and 1 µl can be used as the template for nested PCR. Also, you may need to reduce the number of cycles to 25–30. The optimal conditions for nested PCR should be determined empirically.
What is touchdown PCR (TD-PCR) and when would I need to use it?
During the PCR denaturation step, all DNA molecules will become single stranded. When the temperature decreases for annealing, three types of duplexes can be formed:
- Homoduplexes—annealing of complementary strands
- Heteroduplexes—cross-hybridization of homologous sequences that may have partial homology
- Duplexes between primers and template
To achieve higher specificity, heteroduplex formation should be minimized by increasing stringency (i.e., increasing the temperature) during the initial PCR cycles. Touchdown PCR increases specificity by using reaction conditions that gradually reduce the annealing temperature. The initial annealing temperature is set to several degrees above the estimated Tm of the primers. In subsequent cycles, the annealing temperature is slowly decreased until it reaches the calculated annealing temperature of the primers (Don 1991). By using a higher annealing temperature in the initial PCR cycles, touchdown PCR favors accumulation of amplicons for sequences with the highest primer-template complementarity, thereby enriching for the most specific amplicons. Transitioning to a lower temperature during subsequent cycles reduces stringency, improving priming conditions with the already enriched, desired template. We recommend performing an initial 5–10 cycles with the higher annealing temperature, and then gradually decreasing the temperature until the optimal annealing temperature, or "touchdown temperature," is reached. For example, if the Tm of your primers is 68°C, the recommended TD-PCR conditions for the annealing temperature are:
- 5 cycles at 72°C, then
- 5 cycles at 70°C, then
- >25 cycles at 68°C
Don, R. H., et al. 'Touchdown' PCR to circumvent spurious priming during gene amplification. Nucl Acids Res. 19(14):4008 (1991).
How can I clone a blunt-end PCR product into a TA-cloning vector?
If a PCR product is amplified with a high-fidelity polymerase that generates blunt ends, you can perform A-tailing using Taq polymerase. A brief protocol for adding 3' A-overhangs to PCR products is provided below.
- Purify the PCR product. Before adding overhangs, it is very important to remove all of the polymerase in the reaction by purifying the PCR product using a PCR purification kit or by phenol extraction and DNA precipitation. This step is critical, since the proofreading activity of any residual DNA polymerase would degrade the A overhangs, thus recreating blunt ends.
- Prepare the Taq DNA polymerase reaction mix:
|Final concentration||Volume (µl)|
|Purified PCR product||0.15–1.5 pmol||Varies*|
|dATP (10 mM)||0.2 mM||1|
|PCR buffer with Mg2+ (10X)||1X (1.5 mM MgCl2)||5|
|Taq DNA polymerase (5 U/µl)||1 U||0.2|
|ddH2O||to 50 µl|
*The A-addition reaction works best when a specific amount of the PCR product is used. The recommended amount is 10–100 ng per 100 bp of the PCR product. This corresponds to 0.15–1.5 pmol of PCR product (see table below).
|PCR product size||Amount of PCR product to use|
|100 bp||10–100 ng|
|250 bp||25–250 ng|
|1,000 bp||100–1,000 ng|
3. Incubate for 20 min at 72°C.
Proceed to TA cloning. For optimal ligation efficiency, it is best to use fresh PCR products, since 3' A-overhangs will gradually be lost during storage.
What are PCR inhibitors?
Impurities that interfere with PCR amplification are known as PCR inhibitors. PCR inhibitors are present in a large variety of sample types and may lead to decreased PCR sensitivity or even false-negative PCR results. PCR inhibitors may have both inorganic and organic origins (Schrader 2012).
Inorganic PCR inhibitors include:
- Calcium or other metal ions that compete with magnesium
- EDTA that binds to magnesium, reducing its concentration
Some organic PCR inhibitors include:
- Polysaccharides and glycolipids that mimic the structure of nucleic acids, interfering with primers binding to the template
- Melanin and collagen that form a reversible complex with DNA polymerase
- Humic acids that interact with template DNA and polymerase, preventing the enzymatic reaction, even at low concentrations
- Urea that may lead to degradation of the polymerase
Other organic compounds that can inhibit PCR include:
- Hemoglobin, lactoferrin, and IgG in blood, serum, or plasma samples
- Anticoagulants such as heparin
- Polyphenols, pectin, and xylane from plants
- Ethanol, isopropyl alcohol, phenol, or detergents such as SDS
If inhibitors are present in the template preparation, a 100-fold dilution of the starting template may sufficiently dilute the inhibitor and allow amplification. Alternatively, ethanol precipitation of the template may be needed to resolve the problem.
Schrader, C., et al. PCR inhibitors—occurrence, properties and removal. J Appl Microbiol. 113:1014–1026 (2012).
What is PCR overcycling? How do I know if my product is overcycled?
PCR overcycling is when cycling goes beyond the exponential phase of amplification. Overcycling occurs when the following events take place during PCR:
- Depletion of substrates (dNTPs or primers).
- The reagents (dNTPs or enzymes) are no longer stable at the denaturation temperature.
- The PCR polymerase is inhibited by the product (pyrophosphate, duplex DNA).
- Competition for reagents (dNTPs and primers) by nonspecific products.
- Lowering of the pH of the reaction.
- Incomplete denaturation/strand separation of products at high product concentrations.
The indicator of PCR overcycling is an intense background smear with indistinguishable bands when the reaction is resolved on an agarose gel. It is always recommended to perform a preliminary test to determine the minimal number of PCR cycles needed to yield a sufficient product. The PCR product remains in the linear phase of amplification if the product yield is noticeably increased every 3–5 cycles. We find that overcycled cDNA does not produce suitable template for any downstream application.
What types of mutations can be caused by PCR?
PCR polymerases can introduce different types of mutations, including single-base substitutions, deletions, and insertions. Base substitutions are typically caused by misincorporation of an incorrect dNTP during DNA synthesis.
Polymerases may generate mutations at locations where one or more nucleotides are lost or gained. The frequency of this type of mutation can be sequence dependent, and might be higher in highly repetitive sequences. The most common mutation is a loss of a single nucleotide, which could be a result of template-primer misalignment within a repetitive homopolymeric sequence. DNA rearrangements can also occur when the polymerase terminates synthesis on one DNA strand and continues synthesis after priming occurs on a complementary strand (i.e., strand-switching or jumping PCR). This type of mutation takes place when there is high homology between different regions of DNA. Excessive DNA template in the reaction may also promote this type of mutation.
What factors contribute to PCR-introduced mutations?
The following factors can contribute to PCR-introduced mutations:
- Unbalanced dNTP concentrations
Unequal amounts of the four dNTPs can increase base substitution by as high as eight-fold. Using equal concentrations of the four dNTPs is critical for reducing the error rate of the polymerase.
- High enzyme concentration
- Long incubation times
- Lack of 3'→5' exonuclease activity
- Magnesium concentration
Fidelity is highest when the concentration of Mg2+ is equimolar to the total concentration of the dNTPs. Fidelity decreases when the concentration of free divalent cations increases.
- pH of the reaction
Lowering the pH of the reaction by three units can increase base substitutions up to 60-fold. Low pH (<6.0) may lead to spontaneous purine loss.
- DNA damage
DNA damage can occur at high temperatures, possibly increasing the rate of mutation. One frequent mutation is deamination of cytosine to produce uracil.
- The presence of A stretches in primer sequences
Error rate is increased when the DNA concentration is increased during the final PCR cycles. The total number of cycles should be kept to a minimum to produce the desired PCR product without errors.
What are PCR artifacts?
The following are common PCR artifacts:
- Primer dimers
Primer dimers are formed through self-complementarity at the 3' end of the amplification primers. Primer dimers are suspected if product is produced in a template-free reaction (negative control). To avoid primer dimers, primers shouldn't have complementarity at their 3' ends.
- Chimeric PCR products
Chimeric PCR products can be caused by incompletely extended template. In other words, single-stranded template that was not completely replicated due to premature polymerase termination can anneal to partially homologous template. This creates chimeric PCR products. To minimize chimeras, use the fewest possible PCR amplification cycles.
- PCR bias
PCR bias occurs when some sequences are amplified more efficiently than others due to preferential binding by PCR primers. If one sequence is amplified 10% more than another in one cycle, it will be 17.4-times more abundant after 30 cycles. To reduce PCR bias, use a high ramp rate between the denaturation and annealing steps and use low annealing temperatures. Long extension times (>180 sec) should be avoided.
- PCR drift
PCR drift is due to stochastic fluctuation in the interactions of PCR reagents, particularly in the early cycles when a very low template concentration exists. This artifact is observed in multiplex assays, where a loss of sensitivity is caused by the interactions between different sets of primers. It is important to carefully design primers for these types of assays.
- PCR generates high-molecular-weight products that barely migrate through the agarose gel
There is no good explanation for this artifact. Most researchers assume that this is caused by overcycling, since in the later stages of PCR, both single- and double-stranded molecules accumulate. Accumulation of such single-stranded molecules can create heteroduplexes by competing with the primers. Incomplete denaturation in later stages, when there is a high concentration of PCR products, prevents DNA strand separation, and thus a newly formed amplicon may remain bound to the previously made template. This process could repeat, trapping PCR products in a network of molecules.
Another explanation for the origin of high-molecular-weight smears is the partial extension of templates during initial PCR cycles. Partial extensions could be generated by jumping artifacts—when a primer or single-stranded DNA anneals and extends from one priming site, then anneals partially to a homologous segment elsewhere (see Chimeric PCR products, above). Partially extended molecules can act as new primers, since they contain a free 3'-OH, and could generate chimeric molecules that combine the initial priming site and the "jump" site.
Finally, this type of artifact can also be generated when a crude template is used for PCR. Products amplified directly from animal or plant tissues can become trapped in cell debris, which prevents them from migrating in the gel. This problem can be solved by Proteinase K digestion of the amplified PCR product:
- Add 15 µl of loading buffer containing Proteinase K to the entire 50-µl PCR reaction.
- Before loading your samples onto a gel, add 1 µl of the loading buffer containing Proteinase K to 4 µl of the PCR reaction.
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