In case the name doesn’t give it away, real-time PCR is a PCR application that monitors DNA amplification in real time. This means that amplification is monitored during the PCR reaction, and not at the end of the reaction as with end-point PCR, where PCR products are typically analyzed post-run on agarose gels.
This article is the first in a two-part series. In part 1, we will go through the basics of real-time PCR, including its advantages over end-point PCR, the workflow, the typical data output, the choice of fluorescent labeling systems available and the pros and cons of each, while part 2 will cover the different quantification methods available, setup tips, primer design and quality control.
Real-Time PCR vs. qPCR vs. qRT-PCR, and so on…
Because DNA amplification is monitored in real time in a specialized PCR instrument, real-time PCR is a quantitative method, and for this reason it is also commonly referred to as qPCR (quantitative PCR). When real-time PCR is used to monitor gene expression by quantitative analysis of cDNA (i.e., reverse-transcribed RNA), it may be referred to as real-time RT-PCR, qRT-PCR, or modifications of these. We will use ‘real-time PCR’ throughout this article to avoid confusion.
For standard guidelines on real-time PCR nomenclature and how to report results, check out the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE)1.
Real-Time PCR Workflow
The real-time PCR workflow is quite straightforward, and the PCR reaction setup is similar to end-point PCR.
During sample preparation, the sample (i.e., your template) is prepared by isolation of high-quality DNA or RNA. For gene expression analyses, RNA is subjected to reverse-transcription, providing cDNA as the template for the real-time PCR reaction.
The set up of a real-time PCR reaction is much like an end-point PCR reaction – you prepare (or purchase a ready-made) PCR master mix containing all of the standard PCR reaction components e.g., dNTPS, primers, buffer, and polymerase. However, there is one major difference between an end-point and real-time PCR reaction mix. This is the addition of a fluorescent reporter molecule to real-time PCR reactions, as described below.
Where Does the ‘Real-Time’ Part Come into It?
Real-time monitoring of template amplification is made possible by the presence of a fluorescent reporter molecule in each PCR reaction that produces a fluorescent signal with increasing intensity as the amount of PCR amplicon increases. Broadly speaking, there are two choices of fluorescent reporters for real-time PCR:
1. DNA-binding dyes e.g., SYBR green.
2. DNA-specific fluorescence quencher probes e.g., SensiFAST™ probes, TaqMan® probes, molecular beacons, and Scorpions® probes.
Real-time thermal cyclers are equipped with fluorescence detectors that can monitor the fluorescence signal emitted during amplification. The amount of fluorescence measured is proportional to the amount of PCR amplicon generated, and the change in the fluorescence signal over time is used to calculate the amount of amplicon produced in each cycle. In this way, data acquisition happens in real-time.
Most modern real-time PCR instruments include analysis software that can graph the cycle number vs. fluorescence intensity to provide quantitative information about the amplified DNA. Otherwise, the raw data can be extracted from the instrument and analyzed manually.
The Real-Time PCR Amplification Curve
A typical real-time PCR reaction is carried out over 40 cycles, referred to collectively as ‘a run’. An amplification curve is the initial data output of each real-time PCR reaction within a run. One curve represents one reaction, and since real-time PCR is usually carried out in multi-well strips or plates, a single real-time run can generate a large number of curves. For simplicity, let’s look at an example of one amplification curve (Figure 1).
Figure 1. The real-time PCR amplification curve.
The Threshold Line
The threshold line indicates the maximum level of fluorescence that can be considered as background e.g., fluorescence that will be detected upon the binding of a fluorescent detector molecule to dsDNA in the absence of amplification. The real-time PCR instrument usually calculates the threshold automatically, or the user may set it manually. The baseline or negative control represents a successful negative control, where no fluorescence is detected. Negative controls should always appear below the threshold line.
The Cycle Threshold (Ct)
The first few cycles of the run represent the initiation phase, where all reaction components are in abundance, but the number of amplicons present in each reaction is too low to generate a fluorescent signal that exceeds the threshold line. As the run progresses, the amount of fluorescence detected by the instrument will exceed the threshold line. The cycle threshold (Ct) is the cycle number at which the fluorescent signal emitted by a given PCR reaction reaches the threshold line.
As a very general rule of thumb, samples that generate low Cts probably contain a high abundance of the target sequence, while samples with high Cts contain low amounts of the target sequence. The Ct value is also known as the quantification cycle (Cq) or the crossing point (Cp).
PCR Efficiency
During the exponential phase, all PCR reaction components are present in abundance, and the number of PCR amplicons produced increases exponentially, with a theoretical doubling of the amplicon during each cycle. Here, the PCR efficiency* is often taken to be 2. Real-time PCR analysis relies upon the exponential phase because it provides the most accurate data for quantification, and it is during the exponential phase that the threshold line and Ct are calculated.
* PCR efficiency will be covered in more detail in part 2 of this series.
During the linear phase, amplification slows down as reaction components become limited. The cycle number at which a PCR reaction enters the linear phase may differ from one sample to the next, depending on many factors e.g., the amount of template DNA present to begin with. The reaction eventually reaches the plateau phase where the fluorescence signal remains constant.
Real-Time PCR is More Precise than End-Point PCR
During the linear phase of any PCR run (end-point or real-time), the number of amplicons won’t necessarily increase at the same rate in every reaction, and some reactions are likely to reach the linear phase before others in the same run. Differences between amplification rates are even more pronounced during the plateau phase because many of the PCR components may be depleted, and some PCR reactions may have come to a complete halt. Importantly, initial differences in target abundance between different samples will not be reflected in the linear or plateau phases.
Since quantification occurs during the exponential phase in real-time PCR, we get a true picture of the amount of amplicon present before the differences mentioned above come into affect. However, in end-point PCR, amplicons are usually analyzed on an agarose gel at the end of the run, which may be during the plateau phase. Figure 2 illustrates that while the many samples shown in these amplification curves probably have similar Cps, one would get a completely different picture if these samples were analyzed on an agarose gel!
Figure 2. Real-time PCR vs. End-point PCR
Additional Advantages of Real-Time PCR over End-Point PCR
Besides those alluded to already, real-time PCR boasts a range of advantages over end-point PCR. The most significant of these are outlined in Table 1 below.
Table 1: Real-Time PCR vs. End-Point PCR |
|
|
Increased sensitivity |
High-confidence detection of low-copy targets Very wide dynamic range possible: between 1 and 1011 copies of a target are detectable within a run |
| Highly accurate quantitative data |
Accurate measurements of absolute gene copy number and relative gene expression fold changes because the increase in fluorescence intensity is directly proportional to the amount of target present Closed-tube format eliminates risk of cross-contamination i.e., data is acquired without the need to reopen PCR tubes and process for agarose gels |
| Speed | No time spent preparing and running agarose gels |
How Do the Fluorescent Labeling Systems Work?
As mentioned previously, there are two main two choices of fluorescent reporter for real-time PCR – the DNA-binding dyes and the sequence-specific fluorescence quencher probes. Which one is right for your application will depend on many factors, and before we go further, let’s have a look at how these two systems work:
DNA-Binding Dyes
✔ These dyes, of which SYBR Green is the best known, bind non-specifically to double-stranded DNA (dsDNA).
✔ Dye molecules that are not bound to dsDNA emit very low background fluorescence.
✔ During amplification, the dye intercalates the newly synthesized dsDNA, and the resulting DNA-dye complex emits fluorescence that is detected and recorded by the real-time PCR instrument.
✔ The main disadvantage of DNA-binding dyes is that they don’t exhibit sequence specificity. This means that they can bind to primer dimers and off-target amplicons. Primer design is therefore crucial when using these dyes, and while it is possible to monitor off-target amplification and primer dimer formation with post-run quality control checks, DNA-specific probes may be the best choice when there are concerns about specificity.
Sequence-Specific Fluorescence Quencher Probes
✔ These probes bind DNA with sequence-specificity, and are therefore usually custom-designed for every target sequence to be analyzed.
✔ Probes are typically comprised of 3 parts: the DNA-binding region, a 5′ covalently attached fluorophore, and a 3′ quencher.
✔ An intact probe does not emit fluorescence because the quencher prevents the fluorophore from doing so.
✔ During PCR extension, dsDNA is synthesized and bound by the complementary probe. As Taq polymerase extends along the template, it comes into proximity of the probe, where it cleaves the probe via its inherent 5′ to 3′ exonuclease activity. This releases the fluorophore from the quencher and fluorescence is emitted.
For both detection systems, the amount of fluorescence emitted is proportional to the amount of amplicon present in the reaction.
So Which Fluorescence Detection Molecule Should I Use?
Research groups that analyze large numbers of genes, where the focus shifts regularly, often opt for DNA-binding dyes for convenience and cost-effectiveness. Those that work with the same handful of genes for longer periods of time may find that the time and effort invested in probe design and validation is well worth the rewards of increased specificity in the long run. Table 2 below gives an overview of the pros and cons of each detection system.
Table 2: DNA-binding dyes vs. sequence-specific probes |
||
| Pros | Cons | |
| DNA-binding dyes | Flexibility – suitable for analysis of any dsDNA sequence and no need to custom-design | False positives due to inability of dye to discriminate target and off-target amplicons and primer dimers |
| Economical compared to custom-designed probes | May turn out to be very time-consuming Primers may require extensive optimization to exclude off-target amplification |
|
| Straightforward No need for challenging probe design |
Post-run melt curve essential to monitor and exclude off-target amplification | |
| Sequence-specific probes | Increased specificity because specific hybridization between probe and target is required for signal | May be time-consuming, depending on how challenging probe design is |
| Multiplex reactions possible Probes can be labeled with distinguishable dyes |
Costly because a different probe is needed for each target | |
| Can save time and money depending on how often a given probe will be used No need for post-run melt curves because specificity is ensured by the probe |
Risk of false negatives as probe may not identify splice variants (validation needed) | |
That was it for part 1. Stay tuned for part 2 where we will complete this series by looking at the various real-time quantification methods available, setup tips, and quality control steps that will help you to achieve high-quality real-time PCR results every time.
References
1. Bustin SA, Benes V, Garson JA, Hellemans J, Huggett J, Kubista M, et al. The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin Chem. 2009;55(4):611-22.