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Guidelines for a Successful qPCR Master Mix Comparison

Sarah Teter, PhD and Leta Steffen, PhD
Promega Corporation
Publication Date: 8/16

Abstract

Real-time quantitative PCR (qPCR) is a powerful tool to detect and quantify nucleic acids. By incorporating fluorescently labeled probes or fluorescent double-stranded DNA (dsDNA)-binding dyes into the PCR, product formation can be monitored following each PCR cycle. GoTaq® qPCR Master Mixes for dye-based or probe-based detection are optimized for fast and reproducible qPCR assays. In this guide, we outline some of the most important considerations for comparing the performance of your assay using either GoTaq® qPCR Master Mix with BRYT® Green Dye or GoTaq® Probe qPCR Master Mix to that of your current qPCR reagent. Testing previously-optimized qPCR assays with a new qPCR master mix requires careful experimental design to compare several factors of reagent performance including assay specificity, repeatability, linearity, sensitivity and efficiency. Each of these qPCR assay performance indicators will be discussed with respect to how the qPCR master mix comparison experiments should be performed as well as how to analyze the data.

Introduction

Testing previously-optimized qPCR assays with a new qPCR master mix requires careful experimental design to look at several factors of reagent performance including assay specificity, repeatability, linearity, sensitivity and efficiency. Each of these qPCR assay performance indicators will be discussed with respect to how the qPCR master mix comparison experiments should be set up and how to analyze the data to assess each of these key factors of qPCR assays.

qPCR assay plate setup

  1. Prepare serial dilutions. Start with a high-purity stock of DNA template and prepare 6 dilutions. 1:10 or 1:5 serial-dilutions are recommended. For high-complexity eukaryotic DNA templates, 100ng DNA per reaction is an appropriate starting concentration.
  2. Prepare bulk reaction mixes with your current qPCR reagent and GoTaq® qPCR Master Mix. Combine 2X qPCR master mix, primers (and probe, if applicable), and water to prepare a sufficient volume of reaction mix for 24 20µl reactions + 10% excess for pipetting error. Use your standard optimized primer/probe concentrations in the reaction mixes prepared with your current qPCR reagent and GoTaq® qPCR Master Mix. Add passive reference dye as required.
  3. Set up the assay plate. Add the reaction mix and template to the plate for a final reaction volume of 20µl (e.g. 15µl of reaction mix + 5µl template). Each template concentration should be tested in triplicate. Seal the plate and centrifuge briefly to collect the liquid at the bottom of wells.
  4. Amplify reactions with the appropriate thermal cycling conditions. Use your optimized denaturation and annealing/ extension cycling with your current qPCR reagent and with GoTaq® qPCR Master Mix. Do not change the Hot-Start Activation step for the GoTaq® qPCR Master Mix Cycling Protocol; initial activation at 95°C for 2 minutes is required. Include a dissociation/ melt curve step in assays with double-stranded DNA (dsDNA) binding dye-based amplification detection.

Analysis of qPCR amplification data quality

  1. Ensure that no amplification is observed in no template control (NTC) reactions. Check that the NTC reactions have no Cq value determined. Otherwise, very high Cq values (>38 cycles) may be acceptable, as long as they are not within 3 cycles of the reactions containing your lowest template concentrations.
  2. Analyze the data for each assay separately. For each analysis, include a subset of wells containing one assay while excluding the data from wells in which different assays were performed. Also, exclude the wells containing the NTC.
  3. Use automatic baseline and threshold settings. Select the option for automatic baseline window and threshold settings in the software, even if you use a manual threshold with typical assay use.
  4. Confirm that the appropriate baseline window was set for the data. 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.
  5. Confirm that the appropriate threshold was determined for 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.

Determining which qPCR master mix works best

Once you have confirmed the quality of the amplification data that you have generated, you can look at the specificity, repeatability, linearity, sensitivity and efficiency of the qPCR master mix and primer/probe combinations that you have tested.

  1. Reaction specificity. If using a qPCR master mix with dsDNA binding dye, a single peak in the melt curve analysis indicates specificity. For probe-based detection of amplification, the probe should impart specificity for the amplicon of interest. NTC reactions should show no amplification, late amplification, or primer artifact (low Tm) amplification only.
  2. Assay 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. A data point should be excluded if the standard deviation of Cq > 0.5.
  3. Assay linearity. The coefficient of determination, 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 which throw off the fit of the linear regression equation (r2 < 0.98 if point is included) are excluded as outside the linear range of the assay.
  4. Assay 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.
  5. Reaction 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)

Tool for comparing qPCR reagents used with your assay

This table is a rubric for assessing qPCR assay performance based on assay specificity, reproducibility, linearity, sensitivity and efficiency. Place a check mark in the appropriate column corresponding to the qPCR master mix for each criterion. The qPCR master mix that has the most checkmarks performs best for your assay of interest.

tpub_178_comparison-figureTable 1.

Reference

Bustin, S.A. et al. (2009) The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments. Clin Chem. 55, 1–12

For more detailed guidelines, see the article below.

How to Cite This Article

Scientific Style and Format, 7th edition, 2006

Teter, S. and Steffen, L. Guidelines for a Successful qPCR Master Mix Comparison. [Internet] 8/16. [cited: year, month, date]. Available from: https://www.promega.com/resources/pubhub/guidelines-for-a-comparison-of-qpcr-reagent-performance/

American Medical Association, Manual of Style, 10th edition, 2007

Teter, S. and Steffen, L. Guidelines for a Successful qPCR Master Mix Comparison. Promega Corporation Web site. https://www.promega.com/resources/pubhub/guidelines-for-a-comparison-of-qpcr-reagent-performance/ Updated 8/16. Accessed Month Day, Year.