qPCR and RT-qPCR

Double-stranded DNA binding dyes bind to the double-stranded DNA product of PCR amplification. The unbound dye fluoresces at a much lower level than bound dye, therefore the level of fluorescence is directly proportional to the amount of amplified DNA produced. One advantage of using these dyes is that they will work with standard PCR primers, eliminating the need to acquire specially labeled primers or probes. However, the signal detected is not sequence-specific and you will need to perform additional analysis such as melt curve analysis to confirm that the correct sequence has been amplified.

Probe-based qPCR relies on a reporter-quencher mechanism where sequence-specific probe that is labeled with a fluorescent reporter and a quencher molecule bind to the template. The fluorescent signal appears as probes specifically bind and interact with amplified target produced in the reaction. You will also need amplification primers for your target, but the probe-based detection means your results are more specific compared to the dye-based methos. One main advantage of using probe-based qPCR is that it provides the opportunity for multiplexing. By incorporating different sequence-specific probes labeled with different colored fluorescent reporter dyes, you can increase throughput by detecting several distinct sequences from the same qPCR.

Why use Melt Curve Analysis following qPCR?

You can verify that you have the correct qPCR amplification product by performing melt curve analysis. These curves are generated by allowing the PCR products to form double-stranded DNA at a lower temperature (approximately 60°C) and then slowly ramping the temperature to denaturing levels (approximately 95°C). As the DNA strands separate, the double-stranded DNA binding dye dissociates, and fluorescent signal is lost. The temperature at which 50% of double-stranded DNA (dsDNA) dissociates into single-stranded DNA, is known as the melt temperature (T_m). Both product length and sequence affect T_m, so the melt curve can be used to characterize amplicons present. Nonspecific amplification products can be identified by broad peaks in the melt curve or peaks with unexpected Tm values. By distinguishing specific and nonspecific amplification products, melt curve analysis adds a quality control aspect during routine use. The generation of melt curves is not possible with some probe-based detection methods that rely on the 5´→3´ exonuclease activity of Taq DNA polymerase, such as TaqMan® technology.

RT-qPCR can be done in a single step or by dividing that process into two steps. The 1-step method saves time and work by combining the reverse transcription and amplification reactions, but you can’t save the cDNA to analyze later. In the 2-step process, you perform the reverse transcription reaction first, and then use the cDNA product in a separate amplification reaction. With the 2-step process, you have the option to save the cDNA produced during the reverse transcription reaction to use later, making it possible to analyze different target sequences within an amplified sample.

Quantitative polymerase chain reaction (qPCR) or quantitative real time PCR (qRT-PCR), is a method for measuring the amount of DNA that is present in a sample. Performing PCR in real time means that product formation is measured at each cycle as it is occurring by measuring a change in a fluorescent reporter signal. With traditional, end-point PCR the signal is measured only once after the cycling program has ended. qPCR, sometimes referred to as quantitative real-time PCR or qRT-PCR, should not be confused with RT-qPCR, which is reverse transcription qPCR, where the starting template is RNA that is reverse transcribed into a cDNA target prior to qPCR.

There are two general types of fluorescent reporters that are used in qPCR assays:

Double-stranded DNA-binding Dyes
Double-stranded DNA-binding dyes are dyes that bind to the minor groove of double-stranded DNA (dsDNA). The free dye has a very low fluorescence, whereas the bound dye has a high fluorescence. The more double-stranded PCR product that is produced, the greater the fluorescence signal will be. BRYT Green® Dye is an example of a dsDNA-binding dye. These dyes allow you to use standard PCR primers, so there is no need to order special labeled primers. The disadvantage is that the signal is not sequence-specific, so you will need to verify that the reaction is producing the desired product.

Labeled Primer or Probe
The second type of reporter method uses a labeled primer or probe to detect the amplification product. There are many different ways to modify reporter signal generation. Real-time PCR using labeled oligonucleotide primers or probes employs two different fluorescent reporters and relies on energy transfer from one reporter (the energy donor) to a second reporter (the energy acceptor) when the reporters are in close proximity. The second reporter can be either a quencher or a fluor. If the second reporter is a quencher, the energy from the first reporter is absorbed but re-emitted as heat rather than light, leading to a decrease in fluorescent signal. Alternatively, if the second reporter is a fluor, the energy can be absorbed and re-emitted at another wavelength through fluorescent resonance energy transfer (FRET) and the progress of the reaction can be monitored by the decrease in fluorescence of the energy donor or the increase in fluorescence of the energy acceptor. Some qPCR strategies employ complementary nucleic acid probes to quantify the DNA target. These probes also can be used to detect single nucleotide polymorphisms (3,4). There are many different types of real-time PCR probes, including hydrolysis, hairpin and simple hybridization probes. Probes contain a complementary sequence that allows the probe to anneal to the accumulating PCR product but can differ in the number and location of the fluorescent reporters. TaqMan® Assays are an example of a method that uses a labeled probe. The big advantage that label-based methods offer is the ability to multiplex two or more targets in a single reaction by using different fluorescent reporters. This would allow you to measure an experimental gene and a control gene for normalization in the same reaction.

The amplification threshold is calculated based on the background fluorescence over the baseline region of the amplification curves. The threshold represents the point where the reporter signal is significantly higher than the background levels.

The amplification threshold is a set fluorescence value that the PCR curve must cross. To find the cycle threshold (C_t), the software determines when the fluorescence signal rises above this threshold. There is no specific single equation for the threshold itself, as it is user-defined or software-defined based on the baseline noise of the reaction.

RT-qPCR is the same as qPCR but using a starting template of RNA that is reverse transcribed (RT) to cDNA prior to amplification. A successful RT-qPCR assay is dependent on the success of the RT reaction, and successful reverse transcription depends on RNA integrity and purity. There are several methods for evaluating your RNA before proceeding to the RT step. RNA integrity can be assessed qualitatively by gel electrophoresis or quantitatively using systems such as the Agilent Bioanalyzer, which uses microfluidics to size‐separate and quantitate RNA. RNA quantity and purity can be assessed by absorbance. The quantity is determined by absorbance at A₂₆₀, and quality is measured using a ratio of absorbance at A₂₆₀/A₂₈₀. High‐quality RNA should have a minimum A₂₆₀/₂₈₀ ratio of about 2. Spiking experiments, in which RNA of unknown quality is added to a control assay with template also commonly referred to as an internal amplification control or internal positive control assay, example IAC RT-qPCR Control cat.# AM2040, can be used to detect inhibitors. It is a good idea to include a no-RT qPCR control reaction to detect gDNA contamination. Finally, quantifying the RNA will help normalize the amount of RNA you are including in each reaction.

Anytime you are working with RNA, RNA degradation is a concern. We recommend that you have a dedicated, ribonuclease-free (RNase-free), area for all RNA work. Procedures for creating and maintaining a RNase-free environment to minimize RNA degradation should be followed faithfully. Using an RNase inhibitor (e.g., Recombinant RNasin® Ribonuclease Inhibitor, Cat. #N2111) is strongly recommended.

The fundamental difference between these two approaches is whether the RT step is performed separately from the PCR or in the same tube. For one-step RT-PCR, the RT and PCR steps are performed sequentially in the same tube using the entire amount of the cDNA synthesis products as the template for PCR. For two-step RT-PCR, the RT and PCR steps are also performed sequentially, but only a portion of the cDNA products is used as a template for PCR, which is performed in a separate tube. For this reason, one-step RT-qPCR is is ideal when only one or a few targets is being measured and sample is not limited. However, two-step RT-PCR allows multiple PCRs from a single RT reaction, which works well for quantifying multiple targets or for doing replicate assays.

One-step RT-PCR commonly uses gene-specific primers for both the RT and PCR steps, with one of the PCR primers also acting as the RT primer. Two-step RT-qPCR can use oligo(dT) primers or random primers for the RT step. 

Absolute quantitation in qPCR refers to the precise determination of the absolute number of target DNA copies in a sample. This method requires the construction of a standard curve from serial dilutions of a known concentration of target DNA. The amount of target DNA in unknown samples is then interpolated from this standard curve, providing an exact number of copies.

Relative quantitation measures the change in expression of a target gene relative to a reference gene or control sample. This method is commonly used to compare gene expression levels across different samples or conditions, providing insights into gene regulation and expression patterns.

PCR inhibitors can significantly disrupt qPCR experiments, causing underestimation of nucleic acid targets or amplification failure. These inhibitors may originate from sample materials, co-purify with the DNA/RNA template, or arise during reaction setup. Detecting and mitigating their effects is crucial for successful qPCR. Key Indicators of Inhibition:

1. Changes in C_q Values: An elevated quantification cycle (C_q) value may suggest inhibition but could also reflect reduced target amounts. Distinguishing these requires internal PCR controls (IPC) or template dilution tests.
2. Slope of Standard Curves: A standard curve slope outside the range of -3.3 to -3.6 indicates poor amplification efficiency and potential inhibition.
3. Amplification Curve Morphology: Alterations in curve shape, such as flattening or failure to cross the detection threshold, often signal inhibition.

Common Types of Inhibition:

• Polymerase Inhibition: Often visible as a delayed or altered amplification curve.
• Reverse Transcriptase Inhibition: Manifests as shifts in C_q values, affecting target RNA quantification.
• Signal Interference: Inhibitors can interfere with fluorescence dyes or probes, reducing signal strength.

Strategies for Detection and Mitigation:

• Include IPCs to identify inhibition without template quantity concerns.
• Perform serial dilutions to evaluate C_q consistency and efficiency.
• Optimize sample preparation to minimize carryover of inhibitory substances.

GoTaq® Endure polymerase is engineered for robust performance, providing reliable amplification even in the presence of common PCR inhibitors, ensuring consistent and accurate results with challenging samples.