Detection of low-level pathogens is important in situations where bacterial or viral pathogens cause human illnesses, often through contaminated food or water supplies. In this context, it is important in research to detect pathogens in the food chain at the earliest opportunity to reduce their impact. While real-time PCR (qPCR) can be a useful tool for pathogen detection in general, it is sensitive to PCR inhibitors present in the crude samples common to food and water testing.
By contrast, digital PCR is less sensitive to inhibitors and offers a more robust testing platform. It works by compartmentalizing each bulk prepared PCR sample into thousands of independent microchambers. In contrast to real-time PCR, digital PCR is not as easily affected by most common PCR inhibitors. This is largely due to the natural dilution of these inhibitors across the tens of thousands of microreactions as well as because it is an endpoint reaction. As such it is possible to measure pathogen concentration more accurately and precisely in what would be an otherwise highly inhibited sample. In addition, digital PCR can provide an absolute count of individual pathogenic target molecules, which eliminates the need for reference material or a standard curve.
1. Why does digital PCR tolerate more inhibitors than quantitative or real-time PCR?
dPCR works by digitizing samples into >20,000 microchambers and taking positive/negative readings rather than measuring the rate of amplification. By diluting inhibitory substances that impact the DNA polymerase enzyme and by the nature of endpoint quantification, dPCR enables enhanced robustness against PCR inhibitors often found in more challenging sample types.
2. Can I measure more than one pathogen per digital PCR reaction?
Yes. Using multiple assays in a single dPCR reaction, known as multiplexing, can increase the number of targets detected per reaction. Multiplexing different amplicons in a single dPCR reaction is often as simple as mixing assays with different fluorophores without primer limitation. However, every assay should be tested individually in a simplex format with appropriate positive controls. This can help ensure concordance between the results of both simplex and multiplex dPCR reactions.
Setting the threshold for a multiplex dPCR assay, or an assay with more than one fluorescently labeled probe, should be performed by visualizing the data in a two-dimensional (2D) plot. When thresholding for two targets in 2D, four clusters can be identified in the plot. A 2D plot of multiplex data with thresholds of 5,000 for FAM dye and 1,000 for VIC dye is shown in Figure 1. The black dots represent microchambers that are negative for both targets. The purple dots represent microchambers that are positive only for the target labeled with FAM dye, while the orange dots represent microchambers are that are positive only for the target labeled with VIC dye.
When performing multiplex dPCR, it is typical to detect microchambers that contain more than one target. As the target concentrations increase, the number of microchambers that test positive for multiple targets will also increase. In Figure 1, microchambers that are positive for both targets are represented by the green dots, and the quantity of double-positive microchambers can be easily counted with the software. Thresholding for multiplex assays in 2D will help you best identify an appropriate place for the fluorescence threshold for each channel.
Figure 1. dPCR plot of data from a multiplex d PCR assay with the two targets and two probes labeled with FAM dye (y-axis) and VIC dye (x-axis)
The fluorescence intensities of both dyes are plotted for each analyzed microchamber. The purple dots above the horizontal threshold represent microchambers that are counted as positive for the probe labeled with FAM dye. The orange dots beyond the vertical threshold represent microchambers that are counted as positive for the probe labeled with VIC dye. The green dots represent microchambers that are counted positive for both probes, which indicates both targets are present.
3. Why is dPCR used for routine pathogen quantification?
Sensitive and accurate quantification is necessary to detect changes in research analyzing viral quantities at extremely low concentrations and complements existing qPCR methods for detection. dPCR technology has significant advantages over common PCR platforms, including high sensitivity, high precision, and reliable performance.
Digital PCR does not rely on a reference sample or assay standard; it can be used for absolute quantification, in research measuring the exact copy number of a nucleic acid target of interest. The bulk reaction mix is distributed or digitized into thousands of small independent reactions so that each micro-chamber contains either one or zero copies of the target. Statistical methods are then used to calculate the original concentrations based on the number of positive and negative micro-chambers.