Ask me anything on dPCR? 

你問我答第一集

 集結了眾多技術專家實務精華及經驗分享的你問我答精華集第一集重磅推出 : 

想要開始使用digital PCR 或是正在使用的您有下列的疑問嗎? 

• 為何要從 qPCR 轉換成dPCR?如何轉換呢? 
• 如何解讀 dPCR  scatterplot?？
• 如何優化 multiplexing assays？
• 如何在廢水分析中使用 dPCR？
• dPCR 如何區分rare events like mutations in one single nucleotide?

1. What are the advantages of dPCR over qPCR?

Digital PCR offers several advantages over qPCR, particularly in its ability to provide absolute quantification of DNA without needing reference standards or standard curves. With dPCR, you eliminate the variability introduced by reference standards that might not accurately reflect the sample’s context. By eliminating references, you also save plate space.

Another difference is that in qPCR, inhibitors can cause shifts in cycle threshold (Ct) values, affecting results. In dPCR, even if PCR efficiency drops, as long as the positive and negative partitions can be distinguished, the quantification remains unaffected. This makes dPCR more resistant to PCR inhibitors (Figure 1), which is especially important for complex samples like environmental DNA or wastewater.

In qPCR, the detection of rare mutations is often compromised by competitive binding of primers between abundant wild-type and scarce mutant DNA molecules, all within the same reaction volume. This results in a diminished signal for the mutation. In contrast, dPCR distributes DNA molecules across multiple separate reactions, typically containing only a few molecules each. This partitioning reduces competition and improves accuracy when detecting low-abundance mutations or gene expression levels (Figure 2).

Digital PCR also enhances precision, sensitivity and reproducibility due to its setup of running many reactions (up to 26,000) simultaneously. Compared to qPCR, dPCR also reduces the variability of experiments performed by different operators and labs

2. Why should I spend more on digital reagents when I have a cheaper option with real-time qPCR?

Furthermore, multiplexing is easier in dPCR than in qPCR, which can also reduce costs when you analyse more targets per well. Thus, dPCR can actually be cheaper than qPCR in some cases.

3. How can I transition from qPCR to dPCR? Is the primer and probe design different?

4. What’s the minimum quantity of DNA required for dPCR?

In digital PCR you’re measuring single molecules, so the question of minimum input is not a matter of mass, but molecules as well. In theory, a single molecule is detectable, but realistically, at such low concentrations, random chance has a substantial impact on whether you catch that one molecule or not. Usually, the limit of detection is more in the range of 5-10 molecules for most assays and sample matrices.

5. Is there any risk of cross-contamination between samples (from the same or different runs)? How can I avoid cross contamination?

The nanoplate is sealed before entering the QIAcuity^® Digital PCR System, so there is no risk of cross-contamination once the plate is inside the instrument. There is a slight risk, as would be in any PCR reaction, that moving the pipette above the plate during pipetting could lead to some template being transferred through the aerosol into the wells or some DNA from the operator ending up in the wells. Generally, the risk of contamination in the QIAcuity is extremely low if you work according to good laboratory practices.

6. How can I optimize my dPCR assay?

Use standardized and validated methods for DNA and RNA extraction, as the quality of the genetic material substantially impacts the accuracy of PCR results. Positive controls are also essential for validating assay performance and confirming the assay is functioning correctly by providing a known target that yields predictable results. To validate your assay, you should perform a dilution curve of a known sample to ensure the accuracy of future measurements. 

To ensure the reliability and reproducibility of your dPCR assay, it’s also important to establish:

• Limit of blank – the lowest detectable signal distinguishable from background noise
• Limit of detection – smallest amount of DNA that can be reliably detected
• Limit of quantification – lowest amount of DNA that can be quantitatively assessed with accuracy

Using the QIAcuity, it’s fairly easy to set up an experiment to optimize the dPCR reaction. I would suggest using the essential gradient option with 12 different temperatures, using six different primer/probe concentrations per temperature and programming the PCR to run for 30 cycles, then another five, another five and then another.

7. What are the best practices for determining the limit of detection (LOD) of an assay?

Determining the limit of detection (LOD) involves different methods and is crucial for adhering to Good Manufacturing Practices (GMP) standards. The guidelines for setting LOD are often vague, leaving organizations to define their own procedures. 

A common approach is defining LOD as LOD 95, 

meaning the target is detectable 95% of the time. For example, if a synthetic template is diluted to 0.25 copies per microliter, equating to six molecules per well, it might be detectable in 20 out of 21 tests, representing a 95% detection rate (LOD 95). If the amount is halved to three copies per well, the detection might drop to 90%, indicating an LOD of 90, not 95.

8. Should I include technical replicates in digital PCR? Are any controls necessary?

Most people run technical replicates either to improve precision or to ensure the pipetting is okay. Reason one is not applicable to dPCR, as the QIAcuity measures more accurately than most pipettes work, so using replicates does not increase precision or accuracy. Reason two is still valid since the pipetting is almost the same for qPCR and dPCR. I would recommend using more biological replicates than technical replicates. 

Controls should be used in dPCR as in any PCR. Ideally, I would always run a negative control to be sure that this is clean (measures very low or no positive partitions in the well). The negative control should have a similar matrix to your samples to account for matrix effects. For example, when working with cells, using cell lysate that doesn’t express your gene of interest is a better negative control than pure water. During the validation of an assay, a positive control, ideally of a known quantity, can be useful for determining the assay characteristics.

9. How can I standardize a reaction?

When comparing samples, it’s advisable to use similar amounts of template DNA/RNA to standardize any PCR reaction. I recommend measuring the mass of your samples and using similar amounts in your assays. Another parameter that can be standardized and allows easier multiplexing would be the PCR conditions. The microbial assays from QIAGEN, for example, all follow the same PCR conditions, such as running at the same annealing temperature, etc., making it easier to multiplex.

10. How can I interpret a digital PCR scatter plot?

In the 1D scatterplot of the QIAcuity (Figure 3), each of the partitions of the well (either 26,000 or 8500) is depicted as a single dot. These dots are ordered by the measured fluorescent signal (RFU). Generally, if one or multiple of your target template DNA molecules are present in a partition, the PCR reaction will amplify them, and this will generate a fluorescent signal (based on either probes or the DNA binding dye EvaGreen). What you normally expect is two clusters in the plot, the upper one being positive (so containing one or more molecules) and the lower one being negative (containing no molecules). A threshold is set between the two clusters either by the system’s software or manually by the user to count partitions as positive and negative. These numbers are then used in a Poisson calculation to determine the number of molecules in your sample.