Real-time quantitative PCR (qPCR) is a powerful tool for the detection and quantification of nucleic acids. Over multiple PCR cycles, DNA amplification is monitored by the detection of fluorescent probes or double-stranded DNA intercalating dyes. Quantification is achieved by plotting and analyzing fluorescence signal versus cycle number. qPCR requires a target-specific assay and qPCR master mix, both of which should be carefully chosen and optimized for performance. A master mix is a premixed solution containing all of the components for qPCR that are not target-specific: a thermostable DNA polymerase, dNTPs, MgCl2, and proprietary additives in a buffer optimized for qPCR.
When choosing among master mixes or considering a switch to a new master mix, careful experimental design and analysis is required to investigate reagent performance. Key qPCR performance criteria such as specificity, repeatability, linearity, sensitivity, and efficiency should be considered. This article outlines user considerations and experimental setup for evaluating and comparing qPCR master mixes.
1. qPCR assay plate setup and amplification
- Prepare serial dilutions. Performance of qPCR reagents must be compared over a range of template concentrations, as a single-point comparison of Cq with different qPCR reagents can be misleading. Use 1:10 or 1:5 serial dilutions starting with 100 ng DNA and prepare bulk reaction mixes per manufacturer guidelines. Always include a no-template control (NTC) where the DNA template is replaced with diluent, and test each template concentration in triplicate.
- Set up assay plates. Each qPCR master mix should be compared on individual plates and in separate instrument runs. This allows setup of run parameters for each master mix that follow the manufacturers’ recommended protocols. It also ensures that the data from each qPCR master mix are collected independently, avoiding influence of other qPCR master mixes on signal gain adjustments applied plate-wide by the instrument software.
- Determine amplification conditions. The timing and temperatures for each step of the qPCR thermal cycle are critical for efficient DNA amplification and are influenced by the salts and additives present in the master mix. Start with the manufacturer’s suggested thermal cycling conditions for each master mix, as protocols will vary. Include a final melt curve step when using intercalating dyes. Low amplification efficiencies may indicate a need for optimization of cycling conditions, in which case a new plate with varied denaturation and primer annealing temperatures should be tested by varying temperatures by 1-2oC.
2. Analysis of qPCR amplification data quality
- Analyze data with appropriate settings. If you are evaluating multiple assays, analyze the data for each assay separately. Exclude NTC wells from analysis. Automatic baseline and threshold settings should be used, even if you use a manual threshold with typical assay use, as these settings are affected by both assay and master mix.
- Confirm that the appropriate baseline window was set. On a linear plot of fluorescence signal versus cycle number, confirm that the baseline signal looks flat within the window defined by the analysis software.
- Confirm that the appropriate threshold was determined for the data. The line defining the threshold must fall within the exponential phase of amplification across all reactions for a specific assay. On the semi-log plot (log(fluorescence signal) versus cycle number) the threshold should cross within the portion of each amplification curve that appears straight.
A.
B.
Figure 1. Check the baseline and threshold settings for the amplification data. The baseline window can be checked when fluorescence signal is plotted on a linear scale versus amplification cycle number, (Panel A): the baseline should appear flat within the range of cycles used to establish the baseline. The setting for the threshold for quantitation (Cq) can be checked when fluorescence signal is plotted on a log scale versus amplification cycle number (Panel B): the threshold should fall within the exponential phase of amplification and should be significantly above the background signal.
3. Determining which qPCR master mix works best
- Determine specificity. The Cq value of the NTC wells should be undetermined or very high (>38 cycles) but should not be within three cycles of the reactions containing the lowest template concentrations. Additionally, for dye-based detection, a single peak in the melt curve analysis indicates specificity.
- Assess repeatability. Repeatability can be assessed from the standard deviation around the Cq for the technical replicates. ±0.5 Cq is a reasonable cutoff for high standard deviation. The fewer points excluded for high standard deviation, the better.
- Calculate linearity. The coefficient of determination, or r2, for the fit of the linear regression equation fitted to the semi-log plot, Cq vs. log(input concentration), should be ≥ 0.98. Points at the extreme ends of the concentration range that throw off the fit of the linear regression equations (r2<0.98 if point is included) are excluded as outside the linear range of the assay.
- Determine sensitivity. The sensitivity is the lowest template concentration at which a Cq can be reliably determined. Information about the assay sensitivity can be inferred from the lowest data point included in the standard curve with a good fit of the linear regression equation.
- Assess efficiency. The efficiency for the qPCR assay can be determined from the slope of the linear regression line according to the following equation and is automatically calculated in most software: Efficiency = -1 + 10(-1/slope). Efficiency should be between 90–110%.
Sarah Teter, Ph.D., Leta Steffen, Ph.D., and Samantha Lewis, Ph.D., work at Promega
Images: Promega & Dreamstime Images
Additional qPCR resources by Promega