dPCR vs. qPCR vs. end-point PCR

When comparing digital PCR vs qPCR, we should first be aware of the principle of digital PCR (dPCR). In dPCR, the sample is divided into thousands of independent partitions, so that each partition contains a few, one or no target sequences. Each partition acts as an individual PCR microreactor. At the end of the amplification, partitions containing amplified target sequences are detected by fluorescence. The distribution of the target sequences in the partitions is determined with Poisson statistics. The ratio of positive partitions, which display a fluorescent signal, over the total number of partitions is used to calculate the concentration of the target in the sample. 

For more details on the dPCR method, refer to our beginners’ dPCR guide.

End point PCR is a method for nucleic acid amplification where the maximum number of copies of the DNA or RNA target is produced. This type of PCR offers qualitative answers on whether or not a particular target sequence is present, but no quantitative information.

In more detail, during end point PCR, also known as traditional or conventional PCR, a tube containing the PCR mixture undergoes amplification in a thermocycler until all desired cycles are completed. A portion of the tube’s content is typically analyzed by gel electrophoresis and stained with ethidium bromide dye. Agarose gel electrophoresis separates the DNA fragments in the mixture according to their sizes. The smaller fragments move father through the agarose gel than the larger fragments. Ethidium bromide binds DNA and fluoresces when excited under a UV light. If the amplification was successful, then a product called an amplicon, can be detected as a band on the agarose gel.

For more information, including primer design, method development, qPCR vs PCR, and more, visit our bench guide on PCR.

Quantitative PCR (qPCR) enables the measurement of DNA amplification in real time through monitoring of fluorescence. How does qPCR work? In qPCR analysis, fluorescence is measured after each PCR cycle. The intensity of the fluorescent signal reflects the current amount of DNA amplicons in the sample at that specific time. 

In initial cycles, the fluorescence signal cannot be distinguished from the background. The point where the fluorescence intensity increases above detection levels corresponds proportionally to the initial number of template DNA molecules in the sample. This point, referred to as the quantification cycle (Cq) or threshold cycle (Ct), enables the determination of quantity of target DNA in the sample according to a calibration curve constructed from serial dilutions of standard samples with known concentrations or copy numbers. The most important difference in qPCR vs PCR is that PCR is only semi-quantitative in nature, whereas qPCR analysis is considered a quantitative method.

A qPCR system can also provide semi-quantitative results without standards, but with controls used as reference material. In this case, the observed qPCR results can be expressed as higher or lower multiples with refence to the control. Gene expression studies are common qPCR applications that makes use of semi-quantitative results. 


In qPCR vs PCR, the key differences are quantification, speed and resolution. End point PCR enables qualitative or semi-quantitative analysis at the end of all PCR cycles via an agarose gel or microchip. qPCR relies on fluorescent dyes or probes and calibration curves to deliver quantitative data in real-time. In PCR vs qPCR, qPCR tends to be faster and with higher resolution.


The main difference between RT PCR vs PCR is the starting material. RT PCR uses purified RNA as a template to generate complementary DNA (cDNA). This cDNA is then amplified. Following normalization to standards, RT PCR can be used to determine the relative quantity of starting target gene expression. In contrast, in PCR, the starting template is purified DNA, which can be used for cloning, genotyping or sequencing.

A point of frequent confusion is RT-PCR vs qPCR. RT-PCR, or reverse transcriptase PCR, is a method used to detect and amplify cDNA. In RT-PCR, an RNA sample is reverse transcribed into cDNA, which serves as a template amplified in a PCR reaction. 

qPCR analysis quantifies nucleic acids based on real-time amplification and fluorescence detection. In fact, rather than pitting qPCR vs RT PCR against each other, a combination of RT-qPCR or RT-dPCR is very much possible. RT-qPCR and RT-dPCR methods can be used for quantitative analysis of RNA and gene expression studies.

Real-time PCR vs qPCR

There is no difference between real-time PCR vs qPCR. qPCR is simply another term for real time PCR. qPCR is also known as real time PCR because qPCR systems monitor the amplification of a DNA target in real-time during PCR amplification. Most of the confusion of real time PCR vs qPCR arises from the incorrect abbreviation of RT-PCR as real-time, rather than reverse transcriptase PCR.

More detailed answers to questions on what is qPCR, how does qPCR work, the qPCR workflow and qPCR vs PCR can be found in our PCR bench guide.

The decision of whether to select qPCR or dPCR depends on several factors, such as sample type, presence of PCR inhibitors, economic factors, speed, throughput and others. qPCR is suitable for screening a large number of samples with higher throughput and larger dynamic range. dPCR systems offer more precisions and are ideal for measurement of fractional abundance, or the mutant vs wild type ratio.
Let us explore in more detail the factors that could affect our selection of digital PCR vs qPCR.

In qPCR, the relationship between Ct and a target concentration can vary by a factor of 1000. These variations are caused by many factors including target sequence, background, PCR chemistry, primer efficiency, probe intensity and the qPCR instrument. Further, a qPCR workflow requires calibration with standard curves. To generate a standard curve, sample dilutions over a 5 log range with multiple replicates at each concentration are needed. These calibrations are time-consuming and require additional sample. Further, calibration results may vary from lab to lab.

dPCR relies on statistical analysis to provide an absolute quantification of a target number without influence by the characteristics of targets, primers, probes, background or PCR chemistry. dPCR does not require calibration and standard curves. This makes the dPCR assay more consistent across laboratories and varying reaction conditions. One pitfall of dPCR is that the sample concentration must fall within the dynamic range to minimize partitioning error. Hence, dPCR may require a titration step when dealing with unknown concentrations.


Precision is the ability to reproducibly resolve small differences in copy number, a condition that is especially important for studies in copy number variation (CNV) and gene expression. In qPCR, fluorescence doubles at each cycle, so a singlex assay can normally resolve only a twofold difference in copy number. Better precision or tighter confidence intervals can be achieved by increasing replicates. But higher number of replicates comes at a cost of additional sample with fixed concentrations.

In dPCR, the desired precision can be achieved by increasing the number of partitions to reduce partitioning error. The dPCR method has been shown to have higher resolution and much lower coefficient of variation compared to qPCR.

Detection of rare targets

In qPCR, inhibitors or excess background DNA can impact amplification efficiency and the relationship between Ct and target number. In some applications, such as rare mutation detection, the DNA within a sample may contain mutant and wild-type alleles during thermal cycling. The wild-type molecule could overwhelm and negatively impact amplification of the lower-concentration target by using up the polymerase, nucleotides and probes.

In dPCR, partitioning enriches the target from the background, which improves amplification efficiency and tolerance to inhibitors. More partitions can be used to increase the dynamic range, so that high-concentration wild-type and low-concentration mutant can be detected in the same run.

Dynamic range

The dynamic range, or the range of sample concentration within which a PCR system can perform quantification, is higher for qPCR than dPCR. This is because in dPCR, the maximum number of quantifiable targets is limited by the number of partitions. If the sample concentration is above the dynamic range, the sample must be first diluted, which adds an extra step into the dPCR workflow. qPCR might be more suitable for measuring large differences in expression between two genes.

Large volume samples

If a PCR instrument’s sample volume is smaller, subsampling errors might be introduced, as a minimum number of rare targets necessary for quantification might not be collected. qPCR can accommodate larger-volume samples than dPCR, so that qPCR is better suited for detection of low concentrations in larger sample volumes. 


Sample throughput is defined by the number of unique samples that can be processed per day. During qPCR, samples are not partitioned and are read during thermal cycling. The dPCR workflow requires pre-PCR partitioning and post PCR-reads of each partition, adversely affecting throughput. However, to match the precision of dPCR, qPCR requires running replicates, also reducing sample throughput. 
To summarize, when it comes to dPCR vs qPCR for high-throughput applications, qPCR is more suitable for cases where similar samples are processed with the same protocol and a single calibration. dPCR might be a better choice for absolution quantification of dissimilar samples.

Operational considerations

In terms of ease of use, qPCR is a more familiar technique, but dPCR does not require calibration. Hence, digital PCR is more suited to automation and handling by personnel without specialized training. The dPCR method tends to generate more reproducible data across laboratories.


dPCR instrument cost is currently higher than qPCR instrument cost. However, the per sample cost in qPCR is impacted by added expenses for calibration standards, which are not needed in dPCR. With the advancement of dPCR instruments, the cost of dPCR machines is expected to keep dropping.

Lively debate on digital PCR vs qPCR

qPCR is an established technique for gene analysis used in a broad range of applications. qPCR measures amplification as it occurs, whereas endpoint, conventional or traditional PCR collects results after each reaction is complete. End-point PCR is a qualitative or semi-quantitative assay. qPCR can provide relative or absolute quantification based on the number of amplification cycles and standard curves to determine the initial amount of template nucleic acid in each sample. dPCR uses statistical methods to obtain absolute number of molecules in the starting reaction without the need for references and standard curves.

The difference between qPCR and PCR, and comparison of qPCR vs PCR and qPCR vs dPCR can be found in the table below.

  End-point PCR  Quantitative PCR
Digital PCR
Measures final amount
of PCR product
following completion
of a desired number
of PCR cycles 
signal of a bulk
reaction mix
after each
PCR cycle 
Involves partitioning
of sample into many
small compartments,
endpoint PCR
and fluorescence
of positive/negative
signal for each
Quantification  Qualitative
to semi-quantitative 
Quantitative based
on standard curves 
Quantitative based
on Poisson statistics 
Precision  ++  +++ 
+++  ++ 
Cost  ++  +++ 
Multiplexing  +++ 
Ease of use  Established method,
but PCR workflow
with many manual
steps, including
staining of gels
with hazardous
ethidium bromide dye 
Established method,
but requires trained
Little training
is required;
data is highly
reproducible across
different laboratories 

Because of their features, conventional PCR, qPCR and digital PCR are suited to different applications.

End-point PCR applications

As one of the most established types of PCR in the lab, traditional PCR is regularly used to determine the presence or absence of target in a sample. Downstream applications of end-point PCR include cloning, genotyping, colony screening, sequencing and others. 

qPCR applications

Due to its quantitative nature, real-time PCR is routinely used to quantify gene expression and detect siRNA, lncRNA, miRNA. The qPCR method is also suitable for analysis of copy number variation, SNP genotyping, microarray verification, assay validation, pathogen detection and analysis of environmental samples.

Digital PCR applications

Digital PCR enables absolute quantification of nucleic acids without the need for reference material. The characteristics of the dPCR method enable detection of SNPs, DNA methylation, chromosomal translocations, alternatively spliced mRNA, rare alleles and copy number variations. dPCR is frequently used in cfDNA analysis, as well as in quantification of viral and bacterial loads. Digital PCR assays are also complementary to next generation sequencing (NGS), as the dPCR method is ideal for quantification of NGS libraries. Digital PCR can also be used to validate nucleic acid standards, to detect pathogens, identify species, investigate GMOs and analyze challenging environmental samples, such as plant, soil and waste.

In summary, the benefits and limitations of end-point PCR, qPCR and dPCR are shown in the table below.

  End-point PCR  qPCR  dPCR 
  • Poor precision
    and low sensitivity
  • Limited dynamic
  • Low resolution
  • Non - automated
  • Size-based
    discrimination only
  • Qualitative data
  • Cumbersome,
    workflow with post
    PCR processing
  • Relies on standard
    curves, which add time,
    cost and adversely
    affect PCR efficiency
  • Less precise than dPCR
  • Sensitive to PCR
    inhibitors, contaminants,
    nontarget DNA
  • Poor interlaboratory
    data reproducibility
  • Assay development
    requires training
  • Low dynamic range
  • Not suitable for very
    large sample volumes
  • Could have lower
    than qPCR
  • Experimental design
    is more challenging
    than with endpoint
  • Easy to plan
  • Simple operation
    without extra
  • Cost-efficient use
    of consumables
    and reagent
  • Wide dynamic range
  • Two-fold changes
    in target concentration
    can be detected
  • Fast turnaround time
  • Low chance
    of contamination
  • No post-PCR
  • No calibration
    curves, so PCR
    efficiency is
    unaffected by
    differences between
    calibrant and sample
  • Very high precision
  • Suitable for detection
    of rare targets in high
    background of
    non-target DNA
  • Less affected
    by PCR inhibitors

If you are interested in converting your qPCR workflow to dPCR, the process might not be as painful as you would think. Much of the fundamental chemistry is similar between qPCR and dPCR. For example, primers, probes, DNA binding dyes, primer and probe design, one-step or two-step reactions are highly similar between qPCR and dPCR. But there are several factors to take into account for a trouble-free transfer from qPCR to dPCR. Consider using:

  • Pre-designed and conditionally validated assays from commercial sources – these are usually dMIQE-compliant and tailored to unique master mixes and kinetics of thermocyclers specific to a dPCR system
  • Published, peer-reviewed designs – make sure the assays in the literature you are reading meet dMIQE guidance; double-check the specificity of the published primers
  • In-house designed assays – follow the recommended conditions of your dPCR system. Your primers should ideally:
    • Be designed with specialized software (Primer3Plus, Primer Express)
    • Generate amplicons ≥150 bp
    • Be 18–30 nucleotides in length with 30–70% GC content
    • Have a melting temperature between 58 and 62°C and within 2°C of each other
    • Be void of highly repetitive sequences, 3’-end cross-complementarity, within-or-across primer complementarity, 3’template mismatch, ≥3 Gs or Cs at 3' end, regions with secondary structure specifically at the binding sites of the primers
    • Be unique for the template sequence (verified with a BLAST search)

    And your probes should be:

    • Designed with specialized software (Primer3Plus or PrimerExpress)
    • The melting temperature of the probes should be 8-10°C higher than the melting temperature of the primers
    • Free of a G at the 5’-end of the probes and free of runs of ≥4 G nucleotides
    • With a binding strand so that the probe has more C than G bases
    • Not complementary to the primers
    • Designed under the same settings as the primers, so that they work optimally under the same cycling conditions (60°C annealing/extension)

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Hindson CM et al. Absolute quantification by droplet digital PCR versus analog real-time PCR. Nature Methods. 2013; 10:1003–1005.

Kralik P and Ricchi M. A Basic Guide to Real Time PCR in Microbial Diagnostics: Definitions, Parameters, and Everything. Frontiers in Microbiology. 2017; 8.

Pecoraro S et al. Overview and recommendations for the application of digital PCR. EUR 29673 EN, Publications Office of the European Union, Luxembourg, 2019. https://gmo-crl.jrc.ec.europa.eu/doc/WG-dPCR-Report.pdf

Seidman LA. Basic Laboratory Calculations for Biotechnology. 2nd ed. CRC Press; 2021.