End-point PCR and PCR Primers Support - Troubleshooting

Here are some suggestions:

• Make sure primers don’t have complementary sequences at the 3′ ends.
• Try using a hot-start polymerase.
• Optimize the annealing step by increasing the temperature in 2–5 degrees C increments, and minimizing the annealing time. You can try higher annealing temperatures in the first few cycles, and lower annealing temperatures in the subsequent cycles.
• Optimize the magnesium concentration for each template and primer combination.
• Try nested PCR.

Please see some reasons below for seeing smearing:

• The enzyme, primer, Mg2+, and/or dNTP concentration was too high.
• The annealing temperature was too low for the primers being used.
• Too many cycles were used.
• The annealing and extension times were too long.
• Bad or old primers.
• Too much template was used initially, try to start with 10⁴–10⁶ molecules
• Consider using additives or PCR Optimizer™ Kit (Cat. No. K122001), especially if you feel strongly that the primers should work/have worked before and are using Taq.

Please see our suggestions below to increase yield:

• Do not use a wooden toothpick to pick colonies or scoop out DNA from a gel prior to PCR. It has been reported that this technique can inhibit PCR. [Lee (1995) BioTechniques 18:225].
• Not enough enzyme was used.
• Denaturation/extension temperature was too high and enzyme died prematurely.
• Too much DMSO (>10%).
• Incorrect annealing temperature: run a series of reactions using different annealing temperatures, starting 5 degrees below the calculated Tm.
• Too few cycles.
• Insufficient or too much Mg2+.
• Poorly designed primers: double check primer sequence against template sequence, primers should have similar melting temperatures, avoid complementary sequences at the 3’ end of primers.
• Carryover inhibitors (e.g., blood, serum).
• Denaturation time was too short. Genomic and viral DNA can require denaturation times of 10 minutes.
• Not a long enough extension time was used depending on the size of product being amplified.
• Use of super-irradiated (treated with >2500 mj/cm²) mineral oil will either inhibit or decrease yield of PCR product [Dohner (1995) Biotechniques 18:964].
• Template had long runs of GC's [Woodford et al. (1995) Nucleic Acids Res 23:539 show that by eliminating all potassium from the amplification reactions, GC-rich regions in templates are sufficiently destabilized to allow PCR]. Alternatively, a combination of 1.0 M betaine with 6–8% DMSO or 5% DMSO with 1.2–1.8 M betaine can be used to amplify GC-rich templates [Baskaran (1996) Genome Res 6:633].
• Other inhibitors of Taq DNA polymerase were present (e.g., indigo dyes, heme, melanin, etc.). Add BSA to the PCR (~160–600 μg/mL), increase the amount of Taq, and/or increase the volume of the PCR to dilute out the inhibitor. The concentration of BSA to add may be dependent on the amount and type of inhibitor present. Additionally, fatty acid–free, alcohol-precipitated BSA, or Fraction V BSA all should be effective.

Here are some reasons why your PCR experiment may be failing:

• NaCl at 50 mM will inhibit the enzyme.
• Too much KCl in the reaction. Do not exceed 50 mM.
• Incorrect annealing temperature was used.
• Incomplete denaturation (time and temperature must be long and high enough).
• Template had long runs of GC's [Woodford et al. (1995) Nucleic Acids Res 23:539 show that by eliminating all potassium from the amplification reactions, GC-rich regions in templates are sufficiently destabilized to allow PCR].
• 10% DMSO partially inhibits Taq.
• Hemin (in blood samples) inhibits Taq.
• Use of super-irradiated (treated with >2500 mJ/cm2) mineral oil will either inhibit or decrease yield of PCR product [Dohner (1995) Biotechniques 18:964].
• Do not use a wooden toothpick to pick colonies or scoop out DNA from a gel prior to PCR. It has been reported that this technique can inhibit PCR [Lee (1995) BioTechniques 18:225].
• Other inhibitors of Taq DNA polymerase were present (e.g., indigo dyes, heme). Add BSA to the PCR, increase the amount of Taq, and/or increase the volume of the PCR to dilute out them inhibitor.

Please see the following possibilities and suggestions we have:

• Primer design: try longer primers to avoid binding at alternative sites, avoid 3 consecutive G or C nucleotides at the 3′ end.
• Annealing temperature: increase annealing temperature to increase specificity.
• Mg2+ concentration: try a lower concentration.
• DNA contamination: use aerosol tips and separate work area to avoid contamination, use UNG/UDG technique to prevent carryover.

This artifact occurs when either too many cycles were performed or too much DNA is added to the reaction. Try heating to 65 degrees C and putting sample on ice before loading.

We have the following recommendations:

- Increase the extension time from 1 min/kb to 1.5 min/kb.
- Vary total PCR cycles from 20-40; 35 cycles is typical.
- Try denaturing at 95 degrees C for 45 seconds.
- GC-rich or problematic targets work better with MgSO4 instead of MgCl2.
- For GC-rich templates, test higher annealing temperatures starting with the temperature that is equal to your primer Tm and in increments of 2 degrees C for up to six different temperatures.

Note: Addition of Platinum GC Enhancer solution enhances the amplification of GC-rich and problematic sequences.

We have the following recommendations:

- Increase the extension time from 1 min/kb to 1.5 min/kb.
- Vary total PCR cycles from 20-40; 35 cycles is typical.
- Try denaturing at 95 degrees C for 45 seconds.
- Increase the amount of template for targets >5 kb.
- Use 2.5 U of Platinum™ Taq for each 50 µL reaction.
- GC-rich or problematic targets work better with MgSO4 instead of MgCl2.
- For GC-rich templates, test higher annealing temperatures starting with the temperature that is equal to your primer Tm and in increments of 2 degrees C for up to six different temperatures.
- Vary the KB extender solution from 1.5 µL to 4.5 µL per 50 µL reaction.

EquiPhi29 DNA Polymerase is a thermostable enzyme with a fast reaction speed. Amplification at 45 degrees C for 1-2 hr is sufficient to obtain specific product amplification without background amplification. The reaction is saturated after 3 hr of incubation.

The oligo may not have been fully solubilized. After addition of TE buffer, make sure the oligo was vortexed for a full 30 seconds and/or pipette up and down more than 10 times. Primers may be present along the sides of the tubs, so when resuspending the oligo, the sides of the tubes should be “rinsed” too.

It is important to differentiate naturally occurring mutations linked to the chemical nature of the oligo manufacturing process from the perceived mutations that occur when desalted oligos are used in certain applications.

Naturally occurring mutations are inherent to the chemical synthesis of oligos and the chances of having one single insertion or deletion in a given oligo of about 30 bases is around 2%. Invitrogen will be happy to replace any oligo that falls into this category.

With regards to the perceived mutations, following DNA synthesis, the completed DNA chain is released from the solid support by incubation in basic solutions such as ammonium hydroxide. This solution contains the required full-length oligo but also contains all of the DNA chains that were aborted during synthesis (failure sequences). If a 30-mer was synthesized, the solution would also contain 29-mer failures, 28-mer failures, 27-mer failures, etc. The amount of failure sequences present is influenced by the coupling efficiency. For an oligo of this type, the percentage of full-length oligo would be between 74 and 54%, assuming a 99 or 98% coupling efficiency. This percentage is even lower when you consider oligos that are longer.

Because the oligos are synthesized from 3′ to 5′ end, the primers that are desalted and not purified for length will have missing bases at the 5′ end. Hence, oligos that are desalted are only recommended for diagnostic PCR, microarray, or sequencing. Invitrogen recommends purification of the oligos if they will be used in certain demanding applications such as mutagenesis or cloning, especially if restriction sites are added to the 5′ end.

Other sources of perceived mutations for both desalted and purified oligos are sequencing artifacts, point mutations introduced during PCR, unstable stem-loop structures in the primers, propagation of the plasmid DNA after cloning in an E. coli strain, i.e., muS, mutD, or mutT or a silent mutation selected by the bacterial strain because of codon usage in that strain.

Better purification of the oligos is recommended to provide you with full-length oligo sequence. Adding restriction sites adds on 10 or more bases to the basic 20–25-mer, making primers longer than 30 bases with a relatively low percentage of full-length sequences after desalting. Additionally, failure sequences occur at the 5′ end of the sequence as oligos are generated from 3′ to 5′ end. Therefore, restriction sites introduced at the 5′ end of primers can be compromised, resulting in missing bases.

There are two possibilities that could occur in any round of extension when creating your primer:

1. The added base is not detritylated correctly, missing one base addition but allowing possible extension in the next round.
2. The trityl group was removed, but not coupled or capped correctly before addition of the next base, allowing the chain to continue.

If detritylation occurs inappropriately and/or if the synthesizer has an error and delivers the wrong base, an extra inserted base can occur in your primer. Please contact techsupport@thermofisher.com for assistance.

Most of the time the color should not affect PCR or any other experimental application since typically it is caused by the iodine used in the synthesis. There are some exceptions, however. Brown oligos can also be caused by the primer being overdried, and if this is the case, the primer may not work.

If an oligo appears green in color, this is most likely due to ink falling into the tube. The oligo should still be fully functional. The color can be removed by doing an ethanol precipitation.

If the oligo was overheated, it will appear as a “ball”-shaped pellet attached to the bottom of the tube. This should not affect the quality of the oligo, and the oligo should be readily soluble in water.

The drying method dries the primer in a thin layer along the sidewalls of the tube instead of the bottom, therefore a pellet is not always visible and should still be ready to use.

Primers should be aliquoted for single use before PCR set-up. Heat just the aliquoted primers to 94 degrees for 1 min. Quick chill the primer on ice before adding to the PCR reaction. Some primers may anneal to themselves or curl up on themselves.

Oligos should be run on a polyacrylamide gel containing 7 M urea and loaded with a 50% formamide solution to avoid compressions and secondary structures. Oligos of the same length and different compositions can electrophorese differently. dC’s migrate fastest, followed by dA’s, dT’s, and then dG’s. Oligos containing N’s tend to run as a blurry band and generally have a problem with secondary structure.