In this section, you will find audience questions and thorough answers from our application scientists grouped by common real-time PCR topics.
qPCR Primer and Assay Design
What makes for a good qPCR assay?
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Key things to look for when designing and validating a qPCR assay are efficiency and specificity.
When we run qPCR, in theory, the PCR amplicon should be doubling with each cycle as we take it through the cycling protocol. So an assay that is 100% efficient would be one that is perfectly doubling. We know that nothing ever works out perfectly in biology. So an efficient assay can be between 90% and 110%. That can be determined by validating the assay, running a standard curve, making a serial ten-fold dilution, and using the slope of that best fit line to determine what the PCR efficiency is.
The other thing we want to look for is specificity. We want to make sure that our PCR primers are amplifying only one specific target, the one we're looking for. Anyone who has run a PCR gel enough times knows that sometimes you would run that lane on the gel and you might see extra. When running a gel, that may not be a big deal because you can just ignore them. But SYBRgreen is going to bind to those extra products if you have them. If you have a primer dimer that could be confounding your data.
There are different ways to check the specificity. You can run a melt curve if using SYBRgreen to ensure that your primers amplify only one specific product. Some people run post PCR gels after the PCR to make sure that they only have one band. Some even sequence their PCR products to ensure that they're getting what they're looking for.
What are the factors that I need to consider when designing PCR primers?
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In the case of the reaction itself, first ensure that you do not have primer dimers; that your primers do not bind to themselves, that they have a TM (melting temperature) that's roughly comparable to what you're going to be running the reaction at. So most reactions are run in the 55 to 60 Celsius range. What we're hoping for there is that it's just warm enough for the primers to bind, but not so much that if you say I had a high TM for it, they would bind to sites that were non-cognate, the off-target effect of binding to something non-discriminately.
So above all else, the condition that matters is the TM at which the primers bind and the amplification occurs. I would say the GC content and hairpinning are the other concerns.
I've heard hydrolysis probes are more specific, should I be using those instead of SYBR Green?
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One perception is that because SYBR Green is a double-stranded DNA-binding dye, using hydrolysis probes are going to give you a more specific answer. However, it's important to keep in mind that the most important thing in determining the specificity of an assay is the primer design. I can get an assay that's perfectly specific with SYBR Green if I've designed the primers appropriately.
Regardless of what detection chemistry you're using, it has to be validated for specificity. So it's true that having a probe, a third oligo is going to give you another layer of binding that can enhance specificity. But if your assay design is done properly, and you validate that you can use either chemistry and be perfectly comfortable with your results.
Even though probes can add an element of specificity, they also are sometimes difficult to design. And in particular, when targeting small areas, trying to fit in a 25-base probe within a 60 or 70-base amplicon can be difficult. If you have to compromise efficiency for that added level of specificity, SYBR Green with the right controls, with the right validation is a perfectly specific technology.
There are a number of factors that can be altered to obtain optimum assay performance, which will lead to higher molecular sensitivity, specificity, and precision. A key point is that, while assays can be purchased from a number of skilled commercial providers to determine sensitivity and specificity, how can this process be automated?
PrimePCR PCR Primers, Assays, and Arrays are expertly designed to conform to MIQE guidelines and give optimal performance using algorithms developed in collobaration with BioGazelle. PrimePCR assays are wet-lab validated, and validation data is available for each assay. Choose preconfigured assay for for tens of thousands of gene targets and complete pathways to save design time and assure the best results.
qPCR Sample Preparation
Can I use synthetic DNA as a control for SARS-CoV2?
Yes. Synthetic DNA is an appropriate substitute if inactive or recombinant RNA control material is not available. The benefit of using inactivated virus or recombinant RNA is that it controls for both the reverse transcription and amplification part of the reaction. You can also use it to control as an extraction control.
Bio-Rad provides recombinant RNA SARS-CoV-2 Standards:
How do I amplify RNA? Should I use one step or two step?
This is dependent upon your experimental design and how many targets you are interested in assessing. For pathogen detection, such as an assay used for detection of SARS-CoV-2, time to results and minimal hand-on time is preferable so a one-step protocol is used.
If you need assess many targets or needs to run multiple technical replicates to account for variability in the qPCR assay, then use a cDNA synthesis kit to convert RNA to cDNA then use this pool of cDNA as template for your qPCR reactions.
Real-Time PCR Methods and Validation
How and how often should a lab create standard curves?
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If you want to calibrate your samples to a standard curve, you want to run a standard curve on each plate each time you run qPCR to account for any inter-run variability. Where you may run into problems is when you run a standard curve and samples on plate A and samples but no standard curve on plate B and trying to calibrate plate B Cq back to the standard curve on plate A.
I might run into some problems because that plate B may have come up a half cycle later and there may have been some inter-run variability. If you need to run standard curves, and calculate your quantities in an absolute manner, then you want to make sure that they're on each plate that you're running the samples for.
Are new qPCR instruments coming from Bio-Rad?
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The CFX Opus is the newest real-time PCR system in our robust CFX Family. CFX Opus offers a number of new features:
- More uniform thermal performance
- Expanded connectivity — Wi-Fi, ethernet, and USB
- Cloud connectivity: use CFX Opus on our BR.io cloud platform or with desktop CFX Maestro software
- Network storage drive access for excellent data management
- Shuttle optical system yields consistent optical measurements across your sample plate
CFX Opus Real-Time PCR System
Why is qPCR less widely used when testing targets in cell-free DNA, e.g., rare somatic mutation detection? Is digital PCR better for detecting a rare targets?
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The purpose of that type of study is really to determine, not just whether certain mutations are present, but also at what level they are present. Therefore, you're analyzing the percentage of a certain mutation over the entire amount of wild type DNA in circulating cell free DNA. This is particularly relevant in several types of cancers, and this is used in the field of liquid biopsy. Among the methods that can be used, qPCR, digital PCR and also now NGS are three methods that are being published on. The advantage that digital PCR presents over other methods is that because every reaction is partitioned, you minimize the risks of competition and basically overpowering of the wild type DNA over the fraction of mutated DNA.
In a qPCR reaction, you have one set of primers, two sets of probes, one probe for the mutated allele, one probe for the wild type allele. If you run the qPCR reaction, you can detect a fractional abundance down to about 1% of that mix. Now clinical researchers would like to get as low as possible in order to detect cancers as early as possible. That's when methods like droplet digital PCR provide an advantage.
With the increased demand for qPCR supplies, what can I do to increase my throughput and use fewer reagents? Are there any downsides or caveats to doing this?
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With the COVID-19 pandemic, there has been a lot of extra demand for qPCR reagents. If you are finding limits in the plastics, using 384-well formats instead of 96-well plates can help you get more out of your plates. Regarding reagents, yes, you can do reactions with a less volume. If you are used to running a 20 microliter reaction in a 96-well plate, you can translate that to a 10 microliter reaction in a 384-well plate.
If you are performing pathogen detection, in particular COVID-19 detection, you still need to validate your LOD (limit of detection) with said volumes because you are now putting in less volume and less DNA than before.
How do I ensure that my qPCR follows literature and industy scientific rigor?
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The universal guidelines for this already exist. Within the scientific literature in 2009, there was a set of guidelines, the MIQE guidelines, that were created to ensure qPCR is done correctly, that is replicable, that is transparent, and can be conveyed to anyone else within a publication so scientists can go and reproduce the same outcomes.
See our Learning Center page for more information: MIQE and RDML Guidelines
Our PrimePCR Assays are expertly designed to conform to MIQE guidelines and lab-validated to give optimal performance.
How exactly does the real-time monitoring of amplification in qPCR make it better the traditional PCR when we still have to run the end mixture on a gel to make sure that there isn't any non-specific amplification?
Actually, you can still run a gel on the product of a qPCR reaction. However, there are better ways to assess the reaction specificity. If you run the reaction using an intercalant chemistry (using SYBR Green or Eva Green). you can perform a melting curve analysis, which allows you to determine whether there is a single or multiple products in the PCR reaction. Finally, the use of fluorescent probes is also recognized as a good way to ensure reaction specificity.
How exactly is the target quantified in a sample? Can you explain the complete procedure in detail?
Please visit our Learning Center web page for a comprehensive introduction to real-time PCR technology and methods:
I have a ton of samples, do I have to run my reference genes, genes of interest, and controls on the same plate?
To minimize batch effect, it is recommended to spread samples across multiple plates and include all targets on a single plate. Be sure to include an interplate calibrator (this can be your reference genes!!) on every plate so that you an compare data across plates.
What is the difference between conventional PCR vs. qPCR?
Conventional PCR (or end-point PCR, or just PCR) is the original form of this technique, and it is used for preparative purposes only (cloning, sequencing, preparation of libraries). qPCR (quantitative PCR or real-time PCR) is used for detection and quantification of certain DNA targets.
What kind of controls should I be running on my plate? every plate? every day?
Always include positive, negative, and internal controls on every plate. If performing absolute quantificaton, a standard curve is recommended on every plate.
Real-Time PCR Data Analysis
What is the smallest amount of target that qPCR can detect?
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In theory, qPCR can detect a single molecule. Publications have shown that you could detect a single nucleic acid molecule in a reaction. Most reactions though don't determine the limit of detection of an assay that way. Instead, they basically establish that limit of detection by the lowest amount that can be detected with 95% certainty.
And that also is based on the amount of false positives that could arise from their no template reactions. So long story short, in theory, single molecule, but in reality, closer to five to ten molecules per reaction. That has been the standard that has been claimed at least by most pathogen detection assays. It is very well accepted that if you can detect 10 molecules per reaction, you are at the lowest limit of detection by this method.
If you run a RT-PCR for 50 cycles, for example, would you accept as positive a sample with a CT value of 49? How would you suggest to set thresholds and baselines?
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Any PCR reaction given enough cycles will amplify something. Every PCR is always amplifying some other target at exceedingly low efficiency, and it appears not to occur within the 40 cycles we typically run a PCR because it hasn't had enough time to become a fully manifest species. But anything after about cycles 38 for an efficient PCR is exceedingly spurious. We don't want to put a lot of faith in that, because if we were to back-calculate from there to the starting concentration, after cycle 38 it's less than a single integer copy of DNA. And that can't happen. The minimum piece of DNA you can have is one.
As far as the thresholds and the baselines, they don't really feel any different after cycle 49, you would simply take the samples that you had seen before cycle 30 or so, and extrapolate those out at the inflection point of the amplification plot. As long as you have a sigmoidal amplification plot, and you see an inflection point between the exponential and the linear phases, it wouldn't matter how late it came up, but it would be very suspect without a lot of controls.
How frequently do I need to perform fluorophore calibration and thermal validation for my qPCR instrument?
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General mainentance procedures will vary between instruments and the frequency at which maintenance is performed is dependent upon the laboratory conditions. Generally speaking, it is a good idea to perform an annual a preventative maintenance on any real-time PCR system.
Thermal validation could be part of this PM but is not necessary unless validation is part of the lab GMP procedures. CFX Real-Time PCR systems uptilize an LED-based optical shuttle that does not flucuate in lumen intensity over time so a fluorophore calibration is not required.
Is it always best practice to let the qPCR machine set the threshold/baseline?
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The vast majority of the time it is. However, an algorithm is an algorithm. It can be a good one, but nothing is ever perfect. I've seen a few instances where the software set a threshold, but maybe one sample whose fluorescent level jumped up really briefly at an early cycle and pass that threshold before it came back down. And you really saw that it was amplifying cycle 20 or something like that.
Generally speaking, you can trust where the threshold are set, but you want to look at each data point and make sure that what you're seeing is in conjunction with the data that are being represented.
What is the purpose of the calibrator sample when running a qPCR?
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It depends on what is meant by calibrator sample. If you are running multiple plates, and you're looking at the same gene on multiple different plates that have to be run at the same time, and you want to compare the expression levels or the quantities of different samples for the same gene on those different plates, then you need an inter-run calibrator. You need at least one sample, the same sample, same assay on both plates to account for any inter-run variability. In that case, the calibrator sample is there to account for any differences that may have occurred between this plate and that plate.
If by calibration you mean instrument calibration, some instruments require routine calibration because of the way they read the plate. Some older instruments have a single bulb in a fixed position that is exciting the fluorescence for every well on that 96 or 384-well plate; each well is getting light at a different angle, a different amount of light, and that needs to be accounted for.
Can I use the data from two machines and combine their Cts into one plot to analyze the data?
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You would need to have some type of calibrator in the form of a single sample, or just simply comparing to a standard curve on both plates. If you've taken a sample that's raw Ct or Cq value on a plate, compare that to a known standard curve, then extrapolate that out to a concentration, that can now be compared across any given samples. However, the raw Ct or Cq values as a whole should not be compared directly plate to plate, particularly from machine to machine, where detection differences are going to very much be real.
Do I need to run a standard curve on every plate if I am doing gene expression analysis?
No, it is not necessary to run a standard curve on every plate if you are performing gene expression analysis. A standard curve is useful when first optimizing and validating new assays but once the thermal cycling conditions are dialed in, you do not need a standard curve.
Be sure to run positive, negative, and no template controls on every plate though! And if you experiment requires multiple plates to be run, be sure to include a well that can be used as an interplate calibrator.
How many replicates do I need and how to I know if a well fails?
In qPCR experiments, there are technical replicates and biological replicates. I'll presume we are asking about technical replicates, which are repeated measurements of the same sample. Technial replicates are necessary to account for the variability that is naturally inherent in the chemistry and address the reproducibility of the assay or technique. (Biological replicates, on the other hand, are are independent samples that address the reproducibility of the effect or event that you are studying.).
How many is enough is dependent upon the assay – the more variability there is in the assay, the more technical replicates are required to account for this variability. At minimum, consider running 3 technical replicates if an assay demonstrates 100% efficiency and an R2 close to one (1.0). Data from technical replicates can be excluded if they are a statistical outlier.
Is the ddCT method better than standard curves, or vice versa? Or are both acceptable?
Both methods are acceptable for gene expression experiments, as long as they are performed correctly. In my experience, the vast majority of gene expression experiments are done using the ∆∆Cq (or ddCT) method. Because gene expression experiments are looking for relative differences in gene expression, this allows the user to calculate those differences without having to run a standard curve on every plate and for every assay. It is only necessary to run a standard curve when you are 1. Validating an assay to confirm good efficiency and R-squared or 2. Performing absolute quantification (e.g. viral titer assays).
When performing ∆∆Cq calculations, it's a good idea to use the Pfaffl Method to account for assay efficiency. See chapter 4 of our qPCR Applications Guide. Also bear in mind that, if you're doing gene expression experiments with standard curves, you will still need to normalize to at least one reference gene by dividing the quantity of your gene of interest by that of the housekeeping gene.
Real-Time PCR Troubleshooting
Why are the CT values for all samples not exactly the same-when the starting inputs were the same?
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There is always an underlying assumption that we can truly measure the amount of DNA within a sample using a fluorometer or a spectrometer. That measurement is reliant on either how much light is bounced off of a target that is fluorescing or how much passes through a sample that is occluding because of the DNA there. That is an incredibly rough estimate, and it is assuming a fixed amount of every single equal amount of those genomes or different DNA that you might be working with.
You can get within a very close range, but expecting to be able to extrapolate a spectrophotometric concentration directly to copies of DNA is unfortunately not going to carry over. So it can be off by something on the order of say plus and minus five-fold. If you end up within one or two cycles with this kind of data, that's a testament to you doing everything right.
Are there any common inhibitors that prevent the expression of qPCR runs? Some of my samples are from the wastewater, which does not amplify well.
There are a variety of inhibitors which can affect PCR efficiency. Some of the most common ones can be derived from the sample preparation method (ethanol, guanidinium salts), other are linked to the type of sample (formalin, heparin, hematin).
In your case, the most likely culprit would be humic acid (and humic-like acids). Guanidium-based sample preparation methods typically are very efficient at removing these type of inhibitors, but if it isn't the case, you may want to test different mixes for your qPCR (not all enzymes and formulations were created equal).
How do we ensure that our results are accurate, removing the chances of false positives or false negatives?
Always include controls on your plate and do not deviate from the protocol. Behind every instruction manual is LOTS of R&D work that dials in the parameters to provide results that are as accurate as possible. Of course, if there is every any doubt as to the accuracy of the results, technical replicates can help provide confidence in your data.
qPCR for COVID-19 & Disease Research
What is qPCR, and why is it useful in the detection and research of infectious disease?
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qPCR is a technique that allows us to monitor the same classic PCR reaction we've done for years, but in real time, by using a fluorophore that increases in brightness as we accumulate additional amounts of DNA amplification. So it allows us to take the number of cycles that are necessary to amplify a target and understand the starting amount of material. And this is very useful in studying diseases and infectious diseases, because it's highly specific and can identify things with remarkably low amounts of starting material.
For example, using qPCR, we can detect the SARS-CoV-2 genome, and we can that it is in fact that genome alone, and not another target, at exceedingly low levels that might be found during parts of the infection.
How does qPCR compare against dPCR in terms of utility in pathogen detection applications?
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qPCR and droplet digital PCR are both great technologies, and each of them kind of have their own set of advantages. droplet digital PCR has excellent sensitivity and precision because ddPCR partitions the sample and counts the number of DNA molecules that you have in a sample. Compared to qPCR, one big difference is going to be the throughput and time to answer. With qPCR, you can run a plate in an hour and a half. If you're looking for a positive or negative signal, qPCR quickly detects whether or not you have something. The workflow for droplet digital PCR takes about five to six hours.
If you're looking for absolute quantification, great precision, then ddPCR will help you with that, but it is going to take more time and have a lower throughput. ddPCR offers a great advantage in that the quantification is much less sensitive to the presence of any PCR inhibitors, which is particularly useful when testing raw samples for environmental studies where you start with very crude material.
What are some of the most common applications of qPCR for pathogen testing in food safety, water quality, and environmental areas?
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Here are three examples of qPCR applications in infectious diseases:
- Testing for Legionella in air filters
- Testing for foodborne pathogens
- Testing for norovirus in water
Of course, one timely application of qPCR is to test COVID-19 levels in sewage water to estimate whether the levels are increasing or decreasing compared to previous periods. qPCR is used very heavily for clinical applications. qPCR can be employed to determine the pathogen in certain infections, as in the diagnosis of sepsis to choose which antibiotics to apply to the patient. The rapidity of qPCR and its sensitivity allow clinicians to give a fast answer for those type of questions.
What is the reference method for COVID-19 diagnosis?
While several methods can be used for diagnosing COVID-19 infection, RT-qPCR remains the gold standard, due to its extreme sensitivity, rapidity and low cost.
