Tips for successful PCR and Troubleshooting PCR/qPCR

20 min Read

Polymerase Chain Reaction (PCR) is one of the most commonly used molecular techniques in research, diagnostics and forensics as it allows for fast and inexpensive amplification of target DNA. It is a very sensitive assay and hence can be easily biased by contamination which can disrupt the overall quality of your experiment or research. Furthermore, it requires pipetting minuscule volumes of reagents, making the setup error-prone.

Life scientists across the board have bonded over the relatable experience of “Pipette, Cry, Repeat” - the dreaded feeling of a PCR failure. But, you’re not alone and not without resources! Put those Kimwipe® tissues aside because we have written a comprehensive guide on how to set-up a successful PCR, including how to avoid contamination, designing the best primers, selecting appropriate reagents, and optimizing cycling conditions. A troubleshooting tool is also included that showcases commonly observed problems, an explanation for their potential causes, and our recommended solutions.

PCR Troubleshooting Guide

Best practices for setting up a successful PCR

1. Reduce environmental cross-contamination

Due to PCR’s ability to generate many copies from only small amounts of DNA/cDNA, even the smallest bit of contamination in the starting material can quickly amplify into a big problem! Eliminating the risk of cross-contamination from your work space is essential in order to avoid false positives/negatives and achieve high-quality results. To accomplish this, ensure you have the following work space set up:

  • A dedicated space and equipment for pre-PCR setup vs. post-PCR work: Ideally, dedicate separate spaces for processing and preparing samples (e.g. DNA/RNA isolation, master mix preparation, and template addition) vs. post-PCR analysis. The most common type of contamination is from aerosolized PCR products lingering on benches/equipment that can be unknowingly added to other PCR reactions. At the minimum, assign a dedicated room or lab bench as well as dedicated equipment for your pre-PCR setup that is far away from any post-PCR work. Clean the work area and equipment thoroughly before and after each experiment using bleach. If a laminar flow hood or biosafety cabinet is available, set up your pre-PCR work there as the positive air pressure and the UV light sterilisation will be helpful for keeping contaminants out.
  • Wear clean gloves and a dedicated pre-PCR lab coat before you enter the work area and change them regularly: If you need to leave the dedicated space, get a fresh pair of gloves before re-entering and touching your pre-PCR setup or dedicated equipment.
  • Use sterile filter tips: This will prevent contamination from entering your samples from your pipette.
  • Keep your tubes closed and avoid overloading wells with product: This prevents aerosolization and reduces risk of contaminants sneaking into your samples.
  • Prevent carryover contamination using UNG: To avoid contamination of PCR reactions from previously amplified nucleic acids, dUTP and uracil N -glycosylase (UNG) can be used in real-time PCR mastermixes, rendering previously amplified DNA non-amplifiable in future reactions.

Pre-PCR and Post-PCR Lab Set-Up

Figure 1 – Example of a pre-PCR and post-PCR lab set-up that is well separated to prevent risk of environmental contamination from aerosolized PCR amplicons.

2. Verify that your PCR reaction is actually clean

Even if you have set up a clean dedicated space for your pre-PCR setup, it is important to include a verification step by including controls. We recommend the following:

  • Include a “No Template” Negative Control (NTC): This is a sample that includes all PCR reagents with the DNA template substituted with ddH2O. If any DNA is detected and amplified in this control sample, you know that there is a high chance that there is contamination in your setup. In qPCR, contamination would appear as a fluorescence signal in the NTC reaction, whereas for PCR, you may observe the presence of incorrect or multiple bands in your gel analysis.

3. Design optimal primer sequences

Correct primer design is essential for efficient and precise primer-template annealing! Sub-optimal primer design may contribute to non-specific amplification and formation of primer-dimers (also known as self-dimers) that will compete with template-primer annealing and affect product yields. It is critical to test the efficiency of a new set of primers by:

  • Running amplified products on an agarose gel (for PCR): Primer-dimers will appear as a smear of bands in the 30-80 low molecular weight range
  • Performing a melt curve analysis of amplified products (for qPCR): Because they are a mixture of low molecular weight products, primer dimers will melt at lower temperatures and create a broad peak.

To learn more about factors to consider for optimal primer design please read our article on primer design tips.

PCR Primer Dimers on gel electrophoresis and melt curve analysis

Figure 1 – Primer dimers will appear as a smear of low molecular weight bands on an agarose gel or a broad low temperature peak in a melting curve analysis.

abm offers a one-stop custom qPCR primer design service that saves you time, effort, and money by preventing complications or even failures in qPCR experiments due to poor primer design. Using our proprietary software we are able to run in silico PCR to select and ensure quality primer designs.

4. Ensure PCR reagents are high quality (e.g. Primers/Probes and DNA template)

PCR is only as good as the quality of your reagents! Before beginning PCR, ensure you do your due diligence to make sure your reagents are contaminant-free and high quality. Here are a few tips:

  • Determine the purity and concentration of your template: Use a UV-Vis nano-spectrophotometer, fluorometer, or agarose gel electrophoresis to assess your sample. Using a nano-spectrophotometer, ensure that the 260/280 nm absorbance ratio is within the range of 1.8-2.0 and that the concentration is within the effective range of the instrument.
  • Store your reagents properly: Maintain the quality of the primers, probes and templates you are using by storing them correctly. DNA should be stored at -20°C or -70°C under slightly basic conditions to prevent depurination. Primers/Probes should be stored in appropriate buffers in order to generate high-quality data.
  • Aliquot your PCR reagents: This avoids multiple freeze/thaw cycles that can degrade your reagents. It can also help reduce contamination caused by excessive handling and prevents the headache of having to throw out all your reagents if anything gets contaminated.
  • Use reagents that can prevent gDNA and RNase contamination in your template: If you are performing RT-PCR, it is good practice to use reagents such as our 5X RT All-In-One MasterMix (Cat. No. G592) which effectively removes gDNA and inhibits RNase contaminants so that you can generate a high quality cDNA template.
  • Use autoclaved and filtered ddH2O dedicated for pre-PCR setup use: ddH2O used in PCR reactions should be autoclaved separately from any post-PCR materials before being filtered through a 0.45 micron nitrocellulose filter to reduce contamination.

5. Determine optimal cycling conditions

In PCR, the cycling times and temperatures can be changed to promote amplification under different experimental conditions. Here are some best practices for optimising your PCR cycling conditions:

  • Lower the temperature or shorten the denaturation cycle: This reduces the chances of nucleotide depurination which can result in truncated products or mutations in the PCR products.
  • Adjust annealing temperatures and extension times: These parameters can be adjusted to reduce non-specific amplification and primer-dimer formation. Analyse the melt curve to determine whether the proper annealing temperature was chosen and optimise accordingly.
  • Follow the manufacturer’s recommended PCR cycling protocol: It is also important to ensure the recommended PCR protocol is followed when switching from one PCR enzyme provider to another; this is because each enzyme was developed to be used for different optimal cycling conditions.
  • Verify that the correct PCR program is in use: Once your PCR samples are ready to run, double-check that the program you are about to run was correctly entered. Most thermal cycler instruments are communal and another user may have changed your program without your knowledge.

For a more in-depth discussion of thermal cycling conditions to use for your PCR please read our article on how to optimize thermal cycling conditions.

6. Stay organised

  • Keep track of the reagents: Make a checklist of what reagents need to be added to each sample. After adding each reagent, tick it off your checklist to avoid missing or accidentally double-adding reagents.
  • Use a master mix to reduce the need to repetitively pipette small amounts: This also aids in the reduction of sample-to-sample and well-to-well variation, as well as improves reproducibility. The use of a master mix reduces the number of pipetting steps and makes the entire process less exhausting and error-prone.
abm’s BlasTaq™ 2X PCR MasterMix (Cat. No. G895) is specially engineered to reliably run PCR 3X faster than regular Taq. Starting at an affordable $0.30/reaction, this ready-to-use MasterMix contains gel loading dye and all you need for your PCR reaction. Free samples are available!

Best practices for setting up a successful PCR

PCR and qPCR Troubleshooting Guide

Given that many factors may influence the success of a PCR reaction, many issues can also arise. Not to worry! We’ve categorised all the common issues and provided some solutions you can try. Simply select your problem or observation and our tool will generate some recommended solutions: