A compilation of the most Frequently Asked Questions (FAQs) entailing Real-Time PCR.  .

General Real-Time PCR

How do I determine the efficiency of my real-time PCR assay?

There are a number of ways to determine the efficiency of a real-time PCR assay. The simplest and most commonly used method is the dilution or standard curve method. This method calculates PCR efficiency using the linear regression slope of a dilution series based on either one of the following equations:

E = 10(-1/slope) -1 (or E = 10(-1/slope))

The ideal slope is -3.32, which correlates to an amplification efficiency of 100% (2). Slopes in the range of -3.60 to -3.10 are generally considered acceptable for real-time PCR. These slope values correlate to amplification efficiencies between 90% (1.9) and 110% (2.1).

 

What are the advantages of real-time PCR over traditional PCR?

The advantages of real-time PCR over traditional PCR are it is a closed tube system requiring no post PCR processing. Real-time PCR has higher precision, increased sensitivity (down to 1 copy), increased dynamic range (greater than 8 logs), and high resolution (less than 2 fold differences).

 

What are applications of real-time PCR?

Real-time PCR has been used in quantification of gene expression, viral quantification, validation of array data, pathogen detection, and allelic discrimination.

 

What are some basic real-time PCR terms and their definitions?

Some basic real-time PCR terms and their definitions are:
Amplification plot – plot of fluorescent signal versus cycle number.
Baseline – the initial cycles of PCR where there is little to no change in fluorescence.
Threshold – the arbitrary level of fluorescence used for Cq determination. Should be set above the baseline and within the exponential growth phase of the amplification plot.
Cq – quantification cycle, the fractional cycle number where fluorescence increases above the threshold. Also referred to as Ct (threshold cycle) or Cp (quantification cycle).
R – reporter signal
Rn – normalized reporter signal
ΔRn – baseline subtracted normalized reporter signal
Slope – Used to determine efficiency of RXN. With 10-fold dilutions, a slope of -3.32 indicates a perfect doubling of product per cycle (100% PCR efficiency)
R2 – Reports linearity of standard curve

 

What are the phases of PCR amplification?

There are three phases of PCR amplification: exponential phase, linear phase and plateau phase. The exponential phase is the first phase of PCR amplification. Reaction components are in excess, there is an exact doubling of product each cycle and the reaction is specific and precise. Real-time PCR measures the Cq value at this phase of PCR. The linear phase is the second phase of PCR amplification. The reaction components are being consumed, amplification slows and the reactions become highly variable. The final phase of PCR amplification is the plateau phase. The reaction has completed and no more products are being generated. Traditional PCR takes its measurements during this phase of PCR.

 

I am currently running traditional PCR; can I use the same template, primers and reagents to run a real-time PCR?

In some cases it is possible to convert existing traditional PCR assays into real-time PCR assays with a few considerations around primer design and master mix. Primer design would be one of the first considerations for converting a traditional PCR assay. For real-time PCR it is generally recommended to have relatively short amplicon lengths, in the range of 50 to 150 bp, to maximize PCR efficiency. Larger products can be used if the cycling conditions are changed to accommodate longer extension times but products larger than 300 bp should generally be avoided. In some cases it may be possible to design a TaqMan® probe to hybridize between the two existing PCR primers. If not SYBR® Green I can be used for detection.

The master mix used is another consideration for converting a traditional PCR assay into a real-time PCR assay. If a TaqMan® probe can be designed, it may be possible to use the same master mix used for the traditional PCR assay. If a TaqMan® probe cannot be designed, SYBR® Green I will need to be added to the master mix. In either case, a certain amount of optimization may be needed to obtain good real-time PCR results.

 

Does Helixis sell real-time PCR reagents?

No, Helixis does not currently sell real-time PCR reagents.

 

What are the major detection chemistries used for real-time PCR?

There are two major detection chemistries used for real-time PCR, hydrolysis (TaqMan®) probe based chemistry or SYBR® Green I dye based chemistry.

TaqMan® probe based chemistry, also known as the fluorogenic 5’ nuclease assay, uses an oligonucleotide probe that is designed to anneal to a specific sequence downstream to one of the PCR primers. The oligonucleotide is labeled with a fluorescent reporter dye at the 5’ end and a quencher dye at the 3’. When the probe is intact, the reporter is in close proximity to the quencher and the fluorescent signal is low as the energy from the reporter will be transferred to the quencher through Fluorescent Resonant Energy Transfer (FRET). During PCR, as Taq DNA polymerase extends from the primers, the 5’ exonuclease activity of the enzyme will cleave the annealed probe separating the reporter dye from the quencher dye, increasing the fluorescent signal.

SYBR® Green I is a dye that binds only to double stranded DNA (dsDNA) and its fluorescent signal increases only when bound to dsDNA. During PCR, as more dsDNA amplicon is being produced, the fluorescent signal of SYBR® Green I will increase.

In addition to TaqMan® and SYBR® Green I based detection systems, there are a few other lesser-used detection chemistries. These include Molecular Beacons, Scorpion® probes and LUX™ primers.

 

What is real-time polymerase chain reaction (PCR)?

Real-time PCR uses various fluorescent detection chemistries that allow for the monitoring of the PCR reaction as it progresses. The amount of fluorescent signal generated is directly proportional to the amount of DNA being synthesized during the PCR reaction. Data is collected at each cycle as the reaction proceeds as opposed to the end of the reaction as in traditional PCR. This allows for samples to be characterized by the point in time when amplification is first detected as opposed to the amount of product generated after PCR cycling. The greater the amount of the target sequence the earlier amplification will be detected.

 

 

Detection Chemistries

What are the advantages and disadvantages of TaqMan® based and SYBR® Green I chemistries?

The main advantage of TaqMan® based chemistry is that a fluorescent signal is generated only when there is specific hybridization of the probe to the target sequence. No signal is generated from any non-specific amplification products that may be formed during the reaction. Another advantage is that probes can be labeled with different, spectrally distinct reporter dyes, which allows for the amplification of multiple target sequences within a single tube (multiplex real-time PCR). The main disadvantage of TaqMan® based chemistry is that design and synthesis of different dual-labeled probes is required for each target sequence, which increases assay setup and cost.

The main advantage of SYBR® Green I based chemistry is that it only requires the design and synthesis of two PCR primers, which decreases assay setup and cost. Another advantage for SYBR® Green I based chemistry is the ability to perform melt curves. The main disadvantage of SYBR® Green I based chemistry is that since SYBR® Green I binds to any dsDNA present during the reaction it will bind to and generate a signal for any non-specific amplification that occurs.

 

What is melt curve analysis?

Melt curve analysis is a post PCR analysis that is compatible with SYBR® Green I based assays. During a melt curve, amplicons produced during PCR are dissociated (or melted) by slowly ramping the instrument from a low temperature to high temperature and the fluorescence is monitored throughout. As the amplification products transition from dsDNA to single stranded DNA, there is a sharp decrease in fluorescence as SYBR® Green I is no longer bound. The midpoint of this transition is known as the melting temperature (Tm) of the sample and is characteristic of a given DNA sequence. Melt curves are useful for determining the specificity of a PCR reaction, as any non-specific amplification products will have a different melt curve profile than the target sequence.

 

When using SYBR® Green I based assays, I get amplification in my no template control (NTC) reactions. What is it and how can I eliminate or reduce it?

Amplification in NTC reactions can either be from contamination or non-specific amplification. Performing melt curve analysis can help to identify if the signal is from contamination or non-specific amplification. If it is contamination, the melting curve of the NTC reaction will have the same Tm as your target sequence. Good aseptic technique, using aerosol resistant pipette tips and a real-time PCR master mix with dUTP and UDG can help to reduce any potential contamination.

If the signal is due to non-specific amplification, the melting curve of the NTC reaction will have a different Tm than the target sequence. The most common type of non-specific amplification is primer-dimer formation. There are a number of ways of reducing primer-dimer formation. Optimal primer design is an important first step in preventing primer dimer formation. Primer pairs should be screened using oligonucleotide analysis tools for any secondary structure, self-annealing or cross annealing. Using a hot-start DNA polymerase will prevent the extension of non-specific primer annealing during reaction setup. Decreasing the primer and Mg++ concentrations can reduce primer dimer formation, but can also affect adversely amplification efficiency. Also, additives such as glycerol and DMSO have been known to reduce primer-dimer formation. If reduction in primer-dimer formation cannot be accomplished, re-designing the primers is necessary to obtain good results.

 

Are SYBR® Green I based real-time PCR assays less specific than TaqMan® probe based assays?

The specificity of any real-time PCR assay, whether TaqMan® probe or SYBR® Green I based, is determined by the quality of the assay design. Non-specific amplification can occur for both SYBR® Green I or TaqMan® probe based methods if assay design is poor. Though TaqMan® based assays will not generate a signal for any non-specific amplification whereas SYBR® Green I based assays will. Although not detected, non-specific amplification will affect the amplification efficiency and sensitivity of TaqMan® based assays in the same way as SYBR® Green I based assays. It is important when designing primers for either system to avoid primer sets that generate any non-specific amplification products. With SYBR® Green I based assays, the ability to perform melt curve analysis is advantageous when trying to optimize primer design as any non-specific amplification can be detected and identified in the melt curve. For TaqMan® based assays detecting non-specific amplification usually requires another post PCR analysis method, such as agarose gel electrophoresis of the PCR products. Alternatively, the primers could be used in PCR with SYBR® Green I and melt curve analysis performed after amplification to determine if any non-specific amplification occurs. This allows for the optimization of primer design without the expense of the labeled probe.

 

How do I design primers/probes for a real-time PCR assay?

There are numerous primer design tools commercially available for purchase or freely accessible on the web. These tools simplify assay design significantly. Some widely used primer design tools are Primer Express® (Applied Biosystems), Beacon Designer™ (Premier Biosoft) and Real-Time Design™(BioSearch Technologies). Also there are numerous websites that contain databases of validated primer sets, including RTPrimerDB (http://medgen.ugent.be/rtprimerdb/) and The Quantitative PCR Primer Database (http://web.ncifcrf.gov/rtp/gel/primerdb/).

If designing primers and probes manually, the following criteria should be followed (taken from Primer Express manual):

• amplicon length should be between 50-150 bp to maximize amplification efficiency.
• The optimal primer length is 20 nucleotides and the Tm should be between 58°C to 60°C
• The optimal probe length is between 13 and 30 nucleotides for standard TaqMan® probes and should have a Tm between 68°C to 70°C.
• Primer and probe sequences should not overlap
• The GC content of primers and probes should be between 30% and 80%.
• To prevent quenching, the 5’ end of the probe cannot be a G residue.
• Avoid runs of identical nucleotides. Primer and probe sequences must contain fewer than 4 consecutive G residues.
• Ideally, to avoid genomic DNA amplification primers should span an exon-exon junction in cDNA.

Additionally, a BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi) analysis should be performed on primer and probe sequences to determine if the primers/probe will amplify sequences other than the target sequence.

 

 

Reaction Formats

What is required in developing multiplex real-time PCR assays?

Developing multiplex real-time PCR assays can be difficult and time consuming. As the reaction complexity increases, significant optimization may be required to generate reliable data. It can be a challenge to develop multiplex assays that amplify all targets with equal efficiency. Several factors need to be considered when developing multiplex real-time PCR assays including primer design, the relative expression levels of target sequences and master mix / reagent conditions.

When designing primers and probes for multiplex real-time PCR assays it is important to carefully design each primer/probe set using the same design criteria. If possible, all sequences should be screened against each other to determine any potential primer-dimer formation. Additionally, a BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi) analysis should be performed to determine primer specificity.

If the expression levels of the target sequences are significantly different, the most abundant target will be preferentially amplified and deplete all the reaction components, compromising amplification of the lesser abundant targets. One way to address this issue is to limit the primer concentrations of the most abundant target. The lowest primer concentration that produces the same Cq and PCR efficiency should be used. Primer limiting allows the highest abundant target to amplify and go to completion without depleting all the reagents needed for the other sequences.

Amplification of multiple target sequences creates additional demand for reaction components. Taq DNA polymerase, Mg++ and dNTP concentrations may need to be optimized to improve amplification of all targets. Recently, commercially available master mixes optimized specifically for multiplex real-time PCR have become available. These multiplex master mixes can reduce the amount of time required for optimization.

 

What are applications of multiplex real-time PCR?

Multiplex real-time PCR can be applied to relative quantification experiments where the gene of interest and reference gene are co-amplified in the same reaction. Multiplex real-time PCR can also be used for allelic discrimination assays, where two differentially labeled probes are used to detect two alleles of a single nucleotide polymorphism. Another application of multiplex real-time PCR is pathogen detection, where multiple pathogens can be detected in one reaction.

 

What is the difference between one-step and two-step real-time RT-qPCR?

The difference between one-step and two-step real-time RT-qPCR lies mainly in the reverse transcription step. In one-step RT-qPCR, a short reverse transcription (5-30 min) reaction is followed by a PCR reaction in a single tube. In two-step RT-qPCR the reverse transcription reaction takes place in a separate tube. Each method has advantages and disadvantages depending on the application.

One-step real-time RT-qPCR is useful when analyzing a few genes over a large number of samples. Since both the RT and PCR reactions occur in the same tube, there is less pipetting and sample manipulation, possibly reducing variation and potential contamination. One-step RT-qPCR requires the use of gene specific primers. One-step RT-qPCR may not be as sensitive as two-step since the reverse transcription step is much shorter than in two-step. Also, the reaction conditions needed to support both the RT and PCR reactions may not be optimal for either reaction. Another drawback of one-step RT-qPCR is that it is not possible to archive the cDNA produced during the reverse transcription reaction.

Two-step real-time RT-qPCR is useful when analyzing a large number of genes over a few samples. Two-step RT-qPCR has flexibility in the priming strategy, allowing for oligo-dT, random primers or gene specific primers. Two-step RT-qPCR is generally more sensitive than one-step since the RT reaction is much longer and the RT and PCR reactions occur separately and can be optimized individually. Also, the cDNA produced is more stable than the initial RNA sample and can be more easily archived for future use.

 

What is multiplex real-time PCR?

Multiplex real-time PCR is a technique in which more than one target sequence is amplified and detected in a single PCR reaction. Amplified sequences are distinguished from one another by the use of different dyes conjugated to the TaqMan® probes. The number of targets that can be detected in a single reaction is technically limited only by the availability of spectrally distinct dyes and the ability of the real-time PCR instrument to effectively excite and detect those dyes. Some advantages of multiplex real-time PCR are reduced reagent costs, reduction in sample use, and increased throughput.

 

 

Sample Prep

What are the different starting materials that I can use for real-time PCR?

The starting materials for real-time PCR can be RNA, genomic DNA or plasmid DNA. RNA must be reverse transcribed into complementary DNA (cDNA) before PCR. More recently, kits have become available that can perform RT-qPCR directly from cells without the need for a separate RNA synthesis step.

 

How do I quantify my RNA sample?

The most widely used method to quantify RNA is by traditional UV spectroscopy. A diluted RNA sample is quantified by measuring its absorbance at 260 nm and 280 nm and the concentration is calculated using the equation:

[RNA] µg/ml = A260 x dilution factor x 40
(40 = average extinction coefficient for RNA)

In addition the A260/A280 ratio can be used to estimate RNA purity. An A260/A280 ratio between 1.8 and 2.1 is indicative of a highly pure RNA sample. UV spectroscopy is relatively simple to perform but there are several drawbacks.

It does not discriminate between RNA and DNA so it is advisable to DNAse treat RNA samples before quantifying. DNA in the sample will lead to an overestimation of RNA concentration. Since proteins and residual phenol from the purification can interfere with absorbance readings, it is important to take care in purification to remove these contaminants. Also, absorbance readings are dependent on pH and ionic strength. It is recommended to dilute RNA samples in TE (pH 8.0) and use TE to blank the spectrophotometer before taking absorbance readings.

An alternative method to quantify RNA samples is to use fluorescent dyes, such as RiboGreen® (Invitrogen). RiboGreen® exhibits a strong fluorescent signal when bound to nucleic acids. Samples are quantified in a fluorescence microplate reader or standard spectrophotometer relative to a nucleic acid standard curve of known concentrations. The linear range of quantification using RiboGreen® is three orders of magnitude from 1µg/ml down to 1 ng/ml. The major advantage of fluorescent dyes over absorbance-based methods is that it is not affected by contaminating proteins or organic solvents carried over from the purification process. DNAse treatment is still recommended as RiboGreen® does not discriminate between RNA and DNA.

 

How do I assess the quality of my RNA sample?

RNA quality is perhaps the most important factor in generating reliable and reproducible real-time PCR data. Traditionally, RNA quality was assessed using gel electrophoresis and comparing the 28S and 18S ribosomal RNA bands. Gel electrophoresis is a laborious, time consuming, and low throughput method that requires fairly large amounts of RNA. Recently, automated lab-on-chip capillary electrophoresis systems, such as the Bioanalyzer (Agilent) and Experion (BioRad) have become increasingly popular tools for determining RNA quality. These systems use microfluidic technology to perform electrophoresis on glass chips in a miniaturized scale that overcome some of the issues of traditional electrophoresis. Data is presented as an electrophoretic trace of the RNA sample.

The Agilent Bioanalyzer provides a quantitative measure of RNA integrity known as the RNA Integrity Number (RIN). A proprietary software algorithm examines the entire electrophoretic trace to determine RNA degradation and gives a numerical value between 1 and 10 that is indicative of RNA quality. An RNA sample with a RIN value of 10 is considered a highly intact sample where as a sample with a RIN value of 1 is considered a highly degraded sample.

 

How do I determine if my RNA sample is contaminated with genomic DNA?

Genomic DNA contamination can be problematic for real-time RT-qPCR since genomic DNA can potentially be co-amplified during the PCR reaction leading to erroneous results. To determine if an RNA sample is contaminated with genomic DNA it is important to include a no reverse transcriptase control during the RT step. A no RT control is an important control to include in all RT-qPCR experiments. If the RNA sample is free of genomic DNA contamination there should be no signal generated after real-time PCR for the no RT control samples. If the RNA sample is contaminated with genomic DNA the no RT control samples will generate a signal after real-time PCR due to amplification of the genomic DNA. To avoid genomic DNA contamination RNA samples should be DNAse treated before reverse transcription. Alternatively, genomic DNA amplification can be avoided if the PCR primers are designed to anneal to sequences of the transcript that span a large intron. Primers designed in this way can only amplify cDNA.

 

 

Quantification

What are the minimum controls that I need to run to have confidence in my data?

The controls that are required depend on the type of real-time PCR experiment. No template control reactions are an important control to run for any real-time PCR experiment. If performing reverse transcription, a no reverse transcriptase reaction is important for determining if genomic DNA is present. When performing relative quantification a control (reference) sample as well as a control (reference) gene is required for normalization. Other controls include no template controls to determine if there is contamination or non-specific amplification and positive and negative amplification controls to minimize false negative or false positive results.

 

What is the difference between a relative quantification assay and an absolute quantification assay?

A relative quantification (also known as comparative quantification) assay quantifies changes in gene expression relative to a reference gene and reference sample. An absolute quantification assay uses a standard curve of known quantities to determine the quantity of unknown samples. Relative quantification results report fold changes in expression relative to the reference gene and reference sample. Absolute quantification reports results as an absolute quantity (copies, ng, pg etc) extrapolated from the standard curve.

 

How do I choose an appropriate reference gene?

Selection of a reference gene or genes is a critical step for expression analysis using real-time PCR. Validation of reference genes for each experimental condition is critical for obtaining accurate real-time PCR data. Validation requires determining if expression of the reference gene is stable between cells of different tissues and if any experimental treatment affects expression.

A good summary of the validation process can be found in the following paper:
Dheda K, et al. Validation of housekeeping genes for normalizing RNA expression in real-time PCR. Biotechniques 2004; 37: 112–119.

The process first requires extraction and quantification of RNA samples from the samples under investigation (diseased v. normal; treated v. untreated). Then normalizing the input of RNA into the reverse transcription reaction. The expression of a panel of different reference genes is then measured by real-time PCR and the differences in Cq across the different samples is determined for each gene.

There are a number of software programs available for selecting appropriate reference genes.
GeNorm
BestKeeper
Norm-Finder
They are all useful applications for selecting reference genes for a given experiment.

 

 

High Resolution Melt

What are the applications of HRM?

High resolution melt curve analysis was originally developed for SNP genotyping but has since been applied to different applications of mutational analysis. Applications for HRM include mutation discovery, DNA fingerprinting, species identification, HLA compatibility typing, allelic prevalence, and DNA methylation analysis among others.

 

What is High Resolution Melt (HRM) curve analysis?

HRM is a recent advancement to the traditional melt curve analysis that significantly increases the amount of detail and information that can be captured. It is sensitive enough to be able to differentiate sequence differences within PCR amplicons down to a single nucleotide. Mutations within amplicons are detected as either a shift in the Tm of the product or a change in the shape of the melting curve.

 

What is required to perform HRM?

High resolution melt curve analysis requires a different class of dsDNA binding dyes, extremely precise instrumentation and specialized software.

HRM analysis is generally performed using dsDNA binding dyes other than SYBR® Green I. These dyes are generally known as saturating dsDNA-binding dyes. Some examples of these dyes are SYTO® 9 (Invitrogen), LCGreen™ (Idaho Tech.), EvaGreen™ (Biotium Inc.). These dyes differ from SYBR® Green I in that they are significantly less inhibitory to PCR. This reduced inhibition allows them to be used at significantly higher concentrations than SYBR® Green I that saturate the dsDNA amplicons. Greater dye saturation provides greater sensitivity and resolution of melt curve profiles.

Extremely precise instrumentation is important for HRM. Since some mutations only cause Tm shifts of a fraction of a degree, any thermal or optical uniformity will reduce the ability to detect these sequence changes. To be able to perform HRM analysis an instrument needs to have a fast acquisition rate, precise temperature control and an absolute minimum of sample-to-sample thermal and optical variation.

Also required to perform HRM is software with specialized analysis algorithms that can analyze the shape of melt profiles and group similar melt profiles together. HRM data can be viewed as either normalized melt curves or difference plots. Difference plots show the difference in fluorescence from a selected reference sample. Some software also features an auto-call feature, which can automatically assign genotypes based on melt profiles.