Quantitative PCR (qPCR) and digital PCR (dPCR) are both methods of determining the quantity of a particular nucleic acid target in a given reaction.
To begin, let’s take a look at the foundation of both methods – the polymerase chain reaction:
What is PCR?
Polymerase chain reaction or PCR is a method which can be used to amplify or make copies of specific segments of DNA.
First, primers, or short oligonucleotides that are complementary to a specific region of DNA, are combined with a polymerase and target DNA in a PCR reaction buffer.
That buffer is heated to denature complementary strands of DNA, then cooled to allow primers to bind to the complementary region and for polymerases to incorporate free nucleotides in order to “copy” the targeted segment. This process is repeated or cycled in order to generate many millions of copies of the targeted DNA sequence.
How does qPCR work?
Quantitative PCR (qPCR), builds on the concept of basic PCR by adding measurement of the amplification process at each cycle of PCR.
Fluorescent dyes which are either attached to probes, or which intercalate into double stranded DNA are measured at each cycle. The number of cycles required to generate a signal which is significantly different from the background fluorescence is referred to as the cycle threshold (Ct). This Ct value can be used alongside a standard curve with known concentrations of the target to provide a relative quantification.
Historically qPCR has been the gold standard for measuring nucleic acids such as DNA and RNA, but the resulting data can be highly variable due to reliance on standard curves or references to perform quantification.
This variability of qPCR can be especially problematic for targets with very low abundance (single digits of copies per reaction) or scenarios where small changes in quantity need to be accurately measured – such as measuring changes in gene expression levels.
Quan et al. Sensors 18(4), 1271 
How does digital PCR work?
Digital PCR (dPCR) is an emergent technology that is poised to become the new gold standard in precise nucleic acid quantitation.
Digital PCR (dPCR) works by dividing a bulk qPCR reaction mixture into a large number of small, individual reactions. By breaking the sample into many thousands of partitions, you ensure that one or a few targeted template molecules end up in each partition. With the sample split up, PCR cycling is conducted as normal, and then the endpoint fluorescence of each partition is measured to determine if the target was present or absent.
This has two major advantages. The first is that digital PCR is not reliant on the kinetics of the PCR reaction for quantification. The second is that quantification is nearly as simple as counting positive partitions – eliminating the reliance on standard curves and enabling absolute quantification.
Digital PCR overcomes many of the limitations of qPCR, such as need for standard curves, inability to accurately quantify small numbers of target molecules and the lack of sensitivity in high background conditions.
Understanding the digital PCR difference
Digital PCR (dPCR) enables scientists to divide the sample and assay mixture into a very large number of separate small volume reactions, such that zero or one target molecule is present in any individual reaction. This solves the problem of variability, and eliminates the need for quantification with standard curves. However, many currently available technologies are time-consuming and have complicated, multi-step workflows that may contribute to sample and reagent waste.
A common digital PCR technology utilizes oil based emulsions in order to partition reagents. Droplet digital PCR (ddPCR) is performed by generating droplets in a tube using specialized reagents and a droplet generator. The droplets are then transferred to a 96-well plate, which is then placed into a thermal cycler for PCR amplification. Finally, the plate is moved to a droplet reader that scans each droplet and counts positive “hits.” This method requires many hands-on steps, each with the potential for user errors and sample contamination.
A simpler and more accurate dPCR Workflow
Combinati’s dPCR platform Absolute Q, however, consists of just one instrument, with every step of dPCR taking place in a single plate. The platform essentially turns a multi-step dPCR process into one resembling a simple qPCR workflow. All the user needs to do is to prepare the sample, load the plate, and analyze the sample, all using only one instrument. The run time is typically less than 90 minutes, and the system analyzes more than 90% of the loaded sample, minimizing sample and reagent waste.
Although the workflow is almost identical to that of qPCR, Absolute Q enables scientists to directly count positive results just like any other dPCR method. The simplicity of the system was designed to put reliable and robust digital PCR in the hands of any lab.
Multiplexing enables more data per reaction
The majority of dPCR systems are based on detection in two discrete optical channels, allowing quantification of one or two targets within a single reaction. Combinati’s platform allows for multiplexing up to four different colors, greatly expanding the possibilities for creating customized assays for different applications. These can include applications such as quantification of viral molecules shed through fecal matter into sewage as well as many other applications that require simultaneous monitoring of multiple targets. Digital PCR’s ability to detect rare targets in a high background, combined with multiplexing, simplifies workflows and drives down the cost of analysis for many research areas.1
1. T. Demeke et al., “Critical assessment of digital PCR for the detection and quantification of genetically modified organisms,” Anal Bioanal Chem, 410:4039-50, 2018.