Amber Dance is an award-winning freelance science writer based in Southern California. She is the ALS (Lou Gehrig’s disease) reporter for the Alzheimer Research Forum. She contributes to The Scientist and Nature journals, and has written about topics ranging from record-breaking rocks to bizarre new ant species.
Jana Kent and colleagues at Co-Diagnostics hope to design inexpensive tests that clinics in third-world countries can use to diagnose infectious diseases and genetic conditions. They need a PCR assay that’s cheap, quick and easy to use.
That’s why they were thrilled to discover the Magnetic Induction Cycler, or MIC, from Australia’s Bio-Molecular Systems. The PCR tubes sit in a spinning rotor, so as they spin they are all exposed to even heating from a radiofrequency source—unlike block systems that lose a bit of temperature control, unevenly, as they age. The MIC costs less than $10,000.
The latest generation of machines and methodologies shows how far PCR has come since 1983, when Kary Mullis first dreamed up the chain reaction.
The original version of the technique, endpoint PCR, still has a heavy presence in the lab. Quantitative or real-time PCR (qPCR), used to measure the approximate starting concentration of a target DNA sequence, is also popular. More recently, a form known as digital PCR (dPCR), which counts individual target molecules, has also become readily available.
Endpoint PCR
“Endpoint PCR is still a real workhorse of the lab,” says David Keys, assistant director in the applications R&D group at ThermoFisher Scientific. This kind of PCR amplifies a given target for some future use, such as DNA sequencing or microsatellite analysis. Gel electrophoresis can tell users if they’ve made a product or not, and the size of that product, but that’s about it. “It’s semi-quantitative at best,” Keys says.
Quantitative PCR
This variation on the methodology evolved as the quest for more accurate results required more detailed quantitation of the material produced. Quantitative PCR enables scientists to watch every cycle of the chain reaction, via fluorescence markers. Every cycle should about double the amount of target DNA present. So a sample that takes, say, 24 cycles to reach a predetermined threshold must have started with about half as much target as a sample that reaches that level in only 23 cycles.
One way to label the product DNA is with an intercalating dye such as SYBR Green. However, it’ll light up any product in the test tube—even nonspecific ones.
In contrast, TaqMan probes limit detection to the desired product. They are snippets of DNA that bind to the middle of the target, with a fluorophore on one end and a quencher on the other. When DNA polymerase copies a target, it chews up the probe, separating the fluorophore from its quencher so it can shine. With every cycle, more probes are destroyed and the fluorescence rises. Using TaqMan probes enables researchers to run multiple PCRs in the same tube, with different colors for each target.
Quantitative PCR is commonly used to analyze gene-expression levels. Most often, scientists simply compare their treated or mutant cells to a wild-type control, resulting in a relative quantitation—twice or half as much as the control, for example. They can also compare levels of their gene of interest to a housekeeping gene expected to remain relatively constant. Alternatively, they may use a qPCR reaction with a known quantity of target to create a standard curve of the number of cycles to reach a threshold value vs. starting amount. Then they can use the curve to infer the starting levels of their experimental samples.
Digital PCR
Still, the best qPCR can offer is that one tube started with more or less of the target than another. Digital PCR is more precise. “When you do it digitally, you’re really counting molecules,” says Prescott Deininger, director of the Tulane Cancer Center, who is interested in how mobile DNA elements replicate and move around in tumors. “You already have half a million of these things in the genome, [and] we’re looking for one or two more, so we have a classic needle-in-a-haystack thing.”
Digital PCR involves splitting a PCR reaction into thousands or millions of individual partitions, ideally such that each one contains either zero or one copy of the target DNA. Each partition is a simple endpoint reaction. It’s digital in the sense that each of the PCR reactions either has, or does not have, a product. From the number of positives reactions, researchers can determine how many targets they had to start.
Although the dPCR concept is 20 years old, in the last five years companies have made partitioning easy. Some dPCR systems, such as the QX200 from Bio-Rad Laboratories and the RainDrop from RainDance Technologies, take the vinaigrette approach. They mix the PCR master mix and target with oil and surfactant to create individual droplets of uniform size. After running this emulsion through a thermocycler, scientists use a second machine to analyze the fluorescence of each droplet, in a manner akin to flow cytometry. RainDance recently announced a tripling of fluorescence readout speed, so a set of eight 25-microliter reactions might take just an hour and 20 minutes to analyze, says Darren Link, RainDance’s cofounder and chief technology officer.
ThermoFisher’s QuantStudio 3D uses an alternative approach to partition. It has a silicon chip with 20,000 holes in it. The PCR mixture spreads across the chip and into the holes, and it is then sealed in by oil. After cycling, an imager reads the fluorescence from each well, in about 24 seconds.
The RainDrop produces about 10 million droplets; the QuantStudio 3D and QX200 each produce up to 20,000.
The high precision of dPCR makes it a good choice when scientists want to detect a rare piece of DNA, such as tumor genes within wild-type DNA in a blood sample. “Any researcher interested in detecting ‘rare targets’ or simply [in] attaining absolute quantitation would be a great candidate for using digital PCR,” says Adam Bemis, a global applications specialist with Bio-Rad.
Digital PCR also avoids issues with inhibitors, such as hemoglobin or contaminants in environmental samples, that plague qPCR.
If contaminants affect the reaction rate such that it isn’t a precise doubling, they alter the final result. But dPCR works as long as the amplification happens, even if it’s slowed.
Choosing an approach
Each approach has its advantages and challenges. Dynamic range, for example, is different for the various PCR techniques. Dynamic range refers to the ability to amplify (and detect) unknown products present at various concentrations in a solution. Quantitative PCR has a wider dynamic range than digital PCR; whether you start with 30 or 30,000 copies of the target, qPCR works the same, says Keys. In contrast, dPCR is limited by partition number, which ranges from 10,000 to 10 million, depending on the machine.
Digital PCR also suffers from throughput difficulties. It typically requires three machines—one to make the droplets, the standard thermocycler, and another to read the results. A one-box solution will likely hit the market soon, predicts John Corbett, global business manager for Bio-Molecular Systems.
Cost is also an issue. Corbett estimates endpoint thermocycler prices at about $5,000. A qPCR machine runs $15,000 to $30,000, though fancier versions cost more. A dPCR machine costs $50,000 to $100,000, he estimates; the QuantStudio 3D is a bit cheaper at $44,000. Plus, dPCR requires consumables—such as the silicon chips for the QuantStudio 3D.
“The default choice may be qPCR, unless you’re interested in those areas where digital PCR excels, such as quantification of rare SNPs or low-abundance targets,” says Rod Pennington, senior research scientist at Promega Corp.
With time, that could change, adds Randall Hayden, director of clinical and molecular microbiology at St. Jude Children’s Research Hospital. “We’re still on the first couple generations of [dPCR] instruments,” he points out. With time, Hayden predicts, the method will become easier and less expensive.
The different types of PCR work well together.
Although Hayden mostly turns to qPCR, he also uses digital. For example, he often uses dPCR to precisely quantitate the target solutions he’ll use to make his standard curve for qPCR. And Deininger often uses qPCR to make sure his probes work before he runs a dPCR reaction.
“This is not an either/or sort of comparison,” says Keys. “They work hand in hand.”
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