Almost from the moment the science recognized the value of PCR amplification, it has been a bit of a love-hate relationship. One of the latest additions to the PCR portfolio, real-time or quantitative PCR (qPCR), can be an amazingly powerful tool. However, just like traditional PCR, qPCR can be frustrating. There are a number of parameters that can influence the success of your qPCR assay. Below I have highlighted ten things to consider when trying to improve your qPCR results.
- Sample purity is crucial. Use precautions to minimize the potential for sample cross-contamination. Some practices to help with contamination problems include using aerosol resistant tips, wearing gloves and changing them frequently. Establish dedicated pre- and post-amplification areas so that you are never opening reaction tubes after amplification in the area where you will be setting up new reactions.
- Internal standards will help you have confidence in your results. Incorporating an internal standard control such as a second primer pair that amplifies a housekeeping gene with a consistent expression level in the target samples will allow you to verify the quality of the target DNA or RNA as well as the performance of the reaction components.
- Optimize! I am sorry to say that there is really no way around the fact that you will need to optimize your amplification conditions. The parameters for optimization include all the usual suspects: magnesium concentration, primer design (include the reverse transcription primers if you are doing RT-qPCR), template quality (including the possible presence of inhibitors), cycling parameters, buffer composition, enzyme concentration and including PCR additives or enhancers.
- Decide in advance what results you are after. With qPCR you can determine either relative or absolute quantitation of your target. Although both use the Cq (the cycle number where the amplification curve crosses the amplification threshold), there are differences. Absolute quantification you will need to create a standard curve with known amounts of the template. You do this by plotting the Cq versus the concentration (log concentration). Using a linear regression analysis formula, you can use the Cq value of your unknown sample and to determine the value for that sample. If you are looking for relative quantitation values, you can simply compare Cq values using the rule that for a reaction with 100% efficiency, a single template molecule will result in 2n molecules (where n is the number of cycles).
- Specificity is important, especially with fluorescent DNA-binding dyes. In addition to the optimization information described in number 3, if you are using fluorescent DNA-binding dyes you will need to test your PCR conditions to ensure that the results are producing specific product. Fluorescent dyes will not distinguish between specific and nonspecific amplification products, so you will want to have confidence in your amplification product. Moving forward you can use melting curves to verify the specificity of your PCR products.
- Primer and probe concentrations should be optimized. If you will be performing probe-based qPCR or RT-qPCR, you should optimize the primer and probe concentrations. A good place to start is 900nM for the PCR primer and 250nM for the hydrolysis probe. These are only starting points, so play around with the concentrations to see what gets the best results for your assay.
- Template quality is always important. For RT-qPCR the RNA quality and purity is particularly crucial as it can affect the efficiency of first-strand cDNA synthesis. Whether you are using total RNA or poly(A)+ RNA, you must be sure that your RNA template is free from any genomic DNA (gDNA) contamination. You can test for gDNA in RNA samples by performing a no-reverse transcription qPCR reaction (you should get no amplification product). An alternative approach is to design your primers so that they span and intron, ensuring that no gDNA can be used as a template in the amplification reaction.
- Choose the RT-qPCR method that will work best for your experiment. For RT-qPCR, you will have to decide if one-step or two-step RT-qPCR fits your purposes. In one-step reactions, the reverse transcription (RT) and PCR steps occur in the same tube, so that the all product from the cDNA synthesis reaction acts as a PCR template. One-step RT-qPCR can be more sensitive and requires less pipetting. In two-step RT-qPCR, the RT step is performed in a separate tube and then only a portion of the cDNA is used as a PCR template. Two-step RT-qPCR allows you to perform more than on PCR assay from a single RT reaction.
- The amount of RNA template you will need varies by the target. When you don’t know the expression level of your target, it can be challenging to figure out how much RNA to start with. The amount of RNA template needed will depend upon the abundance of the target. A high-copy-number transcript might only need a few picograms of starting material for detection, whereas a low-copy-number transcript might require well over 100ng of starting material. If you are looking for a place to start, try 100ng of the total RNA template per RT reaction and then adjust accordingly.
- Finally, review the guidelines located in the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE; 1,2). Once you have done all the optimizing and reaction preparation and you are ready to report your results, following these common sense guidelines ensures that those reading your results have the information they need to fully understand, and possibly replicate, your experiments.
So there you have it: Ten things to think about as you start a new qPCR or RT-qPCR assay. This is not an exhaustive list of things that can influence the success of a qPCR assay. If you have things that this list missed and you think are important, please add them as a comment below.
- Bustin, S.A., et al. (2009) The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin. Chem. 55, 611–22.
- Bustin, S.A., et al. (2011) Primer Sequence Disclosure: A Clarification of the MIQE Guidelines, Clin. Chem. 57, 919–21.