Although real-time quantitative reverse transcription PCR, or RT-qPCR, is a powerful tool for many downstream applications, it can be frustrating to run a good RT-qPCR due to its multi-process nature.
Several factors to consider when running RT-qPCR, which influence the outcome, include
- Sampling
- Storage
- Quality and extraction during sample preparation
- Yield
- Inhibitors during RT-qPCR
Fortunately, GoldBio is always thinking about how to make your research easier. So, if you are working with RT-qPCR, take a few minutes to read these tips for running a good RT-qPCR.
Keep it handy too, because it works as a compendium of several articles reporting essential tips for running a good RT-qPCR.
In this article:
Tip 1: Define the chemistry of your RT-qPCR
Tip 2: Chose one or two step RT-qPCR
Tip 3: Always perform controls
Tip 4: Use a master mix qPCR reaction
Tip 5: Double-check qPCR conditions
Tip 6: Optimize and care about your primers
Tip 7: Minimize cross-contamination
Tip 9: Perform normalizations and use reference genes
Tip 10: Print our checklist to run a good RT-qPCR
Tip 1: Define the chemistry of your RT-qPCR
There are two main chemistries for RT-qPCR that are well-known. One is based on fluorescent dyes (typically green), and the other is based on probes.
Fluorescent labeling
With dye-based RT-qPCR, the quantification of the amplified DNA fragments is based on fluorescent labeling.
Commonly, SYBR green is one of the most used fluorogenic dyes that binds to double-stranded DNA, in this case, complementary DNA (cDNA). Consequently, the fluorescence is measured during each amplification cycle.
The fluorescence signal increases proportionally with the amount of replicated DNA. The more copies of DNA molecules, the more intense the quantified fluorescence in real-time will be.
A drawback to dye-based RT-qPCR comes from the generality of the dye because it binds to any dsDNA molecule. Therefore, the dye can also bind to primer-dimers or contaminants such as gDNA (genomic DNA).
To ensure the target DNA is being measured, it is necessary to perform melt/dissociation curves.
Advantages to dye-based RT-qPCR is that they are easy to handle, inexpensive, and only need primers to run.
Probe-based
Regarding probe-based RT-qPCR chemistry, the most common type is a hydrolysis probe.
The hydrolysis probe consists of two components, a fluorophore, and a quencher.
As sequence polymerization proceeds, the probe is hydrolyzed, separating the fluorophore and the quencher, and increasing the fluorescence.
The fluorescence signal is proportional to the amount of probe-target sequences present in the sample. The fluorescence increases as more DNA copies are produced.
Although the probe-based RT-qPCR is more specific than dye-based chemistry, it requires you to design primers and the probe, which also makes it more expensive.
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Tip 2: Chose one or two step RT-qPCR
RT-qPCR requires an extra step compared to conventional qPCR. This step consists of generating complementary DNA (cDNA) using RNA as a starting material. Generating DNA from RNA is necessary due to RNA instability.
Consequently, there are two types of RT-qPCR where both cDNA synthesis and qPCR reactions can be done in one step (in the same tube) or two steps (in separate tubes).
Each of them has advantages and disadvantages.
For instance, one-step RT-qPCR simplifies the process, reduces experimentation time, and reduces contamination risk, because all of it is done in the same tube. However, if you need isolated cDNA for other applications, you won't get it using one-step RT-qPCR, because all of the cDNA produced will be used for the qPCR reaction.
Two-step RT-qPCR allows you to obtain two products, cDNA and the qPCR reaction mixture. Although you must pipette more, you can perform more than a single reverse transcription reaction in your PCR assay.
Tip 3: Always perform controls
My advisor used to emphasize the importance of controls. In short, he used to say "80% controls and 20% treatments." Therefore, before you start your RT-qPCR reactions, plan your controls carefully.
The controls idea is to truly verify whether the results you are seeing are due to the target gene or if they are artifacts coming from contaminants or errors made during the RT-qPCR process.
This is also essential if you work with dye-based qPCR chemistry. Let us take a closer look at each of them.
- No template control: In this control, you will not add your RNA template (or cDNA in two-step RT-qPCR), but the rest remains the same. The idea here is to identify if there are contaminants like genomic DNA or primer-dimers bound to the dye.
- No-RT control: Here, you won't add the reverse transcriptase enzyme. With this control, if a product is detected, it probably indicates that contaminating DNA is present in the sample.
- Internal control: A second primer pair can be incorporated to match the reference gene (or housekeeping gene). It will let you verify the quality of your input cDNA template as well as the reaction components.
Our recommendation is to have all three controls plus the experimental in your qPCR design.
Tip 4: Use a master mix qPCR reaction
A master mix is the mixture of all reagents intervening in a qPCR reaction. A master mix will reduce the variability associated with pipetting well-to-well because it is prepared using a larger volume of each reagent. It will also help in the reproducibility of your experiment.
Furthermore, it is critical to check all reagents during master mix preparation and thoroughly mix all reagents before pipetting into the well plate.
Using master mixes also reduces errors caused by forgetting a reagent such as primers or dNTPs. And proper mixing will help prevent uneven distribution of reagents within each well.
NOTE: For general qPCR, if you’re new to the process or are introducing it to student researchers in your lab, you may want to consider the Goof-Proof qPCR Master Mix series. This product uses a color-changing reaction to help prevent the mistake of forgetting which well you’ve pipetted into.
Tip 5: Double-check qPCR conditions
It surprised me once when I received an email from a colleague who was very frustrated because someone changed his temperature settings on the shared qPCR machine without communicating these changes.
After giving a long speech about ethics, honesty, camaraderie, and good laboratory practices, he finally concluded all his work went to waste, and he had to start from scratch with all the difficulties associated with getting the reagents and samples ready again.
From this, comes a very important lesson: Verify that your cycling conditions are correct, especially if you are using shared devices.
Check and double-check before starting your qPCR run, and remember, a wise researcher plans.
Tip 6: Optimize and care about your primers
In RT-qPCR reactions, the primer is the short sequence you design to bind to your target cDNA template. Even though I said "you design the primers," there are some great primers that may already exist in published articles.
To design and optimize a primer pair, target an amplicon size between 70-150 bp, and span an exon-exon junction.
Illustration depicts a primer spanning an exon-exon junction.
Furthermore, you should select a primer with a GC content between 40-60%.
This is because sequences with a high GC content tend to form secondary structures, such as hairpin loops. Consequently, GC-rich double-strands are difficult to completely separate during the denaturation phase of the qPCR. As a result, DNA polymerase cannot synthesize the new strand without hindrance (Neidher, 2017).
For the above reasons, double-check your primer sequence using online tools such as RNAfold web server or OligoAnalyzerTM tool. These tools will show you the likelihood of having a secondary structure on your primer sequence.
Verify the specificity of your primer using the Primer designing tool from NCBI. This will help you reduce mismatching your primer with the template during the annealing process.
Ultimately, care about your primer sets. Prepare stock solutions of your primers at 100μM, and then create working solutions diluting the original primers at 10–20μM in nuclease-free water.
Store all primer solutions at -20°C.
Working in this way will help you avoid further contamination and freeze/thaw cycles.
Tip 7: Minimize cross-contamination
Many factors can be sources of cross-contamination.
Bare hands have a lot of RNases that can degrade your RNA. Also, using the same pipette set for all parts of the qPCR workflow also can contaminate the tubes.
Contamination can increase because the qPCR technique is so sensitive. Therefore, reducing the potential for contamination is critical for generating qPCR results that you can trust.
To minimize cross-contamination during qPCR, I will provide six tips when setting up your experiment.
- Always wear gloves and change them frequently.
- Use aerosol-resistant pipette tips.
- Apply the three-room rule, where there is a dedicated area for nucleic acid extraction, one for reaction set up for the qPCR reaction, and one for the qPCR cycler.
- Use DNase before cDNA synthesis to remove genomic DNA (gDNA) contamination.
- Decontaminate surfaces and your bench with DNase/RNase killer solutions before proceeding with your cDNA synthesis and qPCR.
- Always use DEPC-treated water or DNase/RNase-free water to perform your experiments.
Tip 8: Verify the RNA quality
The quality and purity of the RNA template is essential for the success of RT-qPCR. Otherwise, degraded or impure RNA can reduce the efficiency of the qPCR reaction. Furthermore, degraded RNA can prevent an accurate representation of gene expression.
RNA can be prepared from fresh tissue or from tissue stored in RNA stabilization solutions.
In either case, work quickly and consider possible sources of RNases and DNases (bare hands, hair, skin, dirty surfaces).
Also, perform RNA quality and quantity validations using Nanodrop or chip-based techniques such as a Bioanalyzer.
Similarly, it is advisable to run gel electrophoresis to verify the presence of two clear bands without smearing.
Tip 9: Perform normalizations and use reference genes
The variation introduced in the qPCR process due to variable RNA inputs can be corrected by normalization using a reference gene.
Although the starting tissue amount used for the RNA extraction can also be used as a normalization parameter, reference genes are more commonly used.
A reference gene is a gene whose expression does not change along with the experimental conditions. This stability in the reference gene expression is used to calculate the ratios of the gene-specific signal to the reference gene signal.
The reliability of your experiment can also improve by including more than one reference gene, which is usually recommended.
Although there are reported reference genes validated for different tissues and species in scientific articles, I recommend you verify that your reference gene does not vary across the analyzed samples.
Some known genes working as references include GAPDH, ß-actin, and 18S rRNA. Start by confirming if the above genes work well for your experiment, otherwise search for another stable gene for your experiment.
Tip 10: Print our checklists to run a good RT-qPCR
It is very easy to generate data with RT-qPCR because it is a high-throughput technique that allows you to test several genes at the same time.
Therefore, performing a careful process when running an RT-qPCR is vital to avoid misleading results and wrong conclusions. As a multi-step technique, each one can add variability to the final output.
To address the issues of variability associated with the qPCR workflow, the "Minimum Information for Publication of Quantitative Real-time PCR Experiments" or (MIQE) guidelines was developed (Bustin et al., 2009). This guide provides parameters associated with
- The experimental design
- Sample
- Nucleic acid extraction
- Reverse transcription
- qPCR target information
- qPCR oligonucleotides
- qPCR protocol
- qPCR validation
- Data analysis
And gives them a level of importance as "essential" or "desirable.”
At GoldBio, we value your time, and therefore, I extracted those parameters considered as "Essential" from the MIQE guidelines and made a packet of checklists for each section.
All you need is to print it, add the data and check the corresponding item when finished.
Furthermore, this checklist will save you a lot of time because, based on this, you can build the materials and methods section of your publication. Also, it will facilitate reproducibility in further qPCR experiments.
In our printable checklist, you will find three columns. The first column presents the parameters associated with each step in the RT-qPCR workflow. The second column is the description and should be filled with the information according to the parameter. The third column is for checking the parameter when ready.
References
Azure biosystems. (n.d.). 5 Essential Tips for Real-time PCR Success. Azure Biosystems.
Bustin, S. A., Benes, V., Garson, J. A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R., Nolan, T., Pfaffl, M. W., Shipley, G. L., Vandesompele, J., & Wittwer, C. T. (2009). The MIQE guidelines: Minimum information for publication of quantitative real-time PCR experiments. Clinical Chemistry, 55(4), 611–622. https://doi.org/10.1373/clinchem.2008.112797
Čepin, U. (2017). Real-time PCR (qPCR) technology basics. BioSistemika.
Grooms, K. (2015). Top 10 Tips to Improve Your qPCR or RT-qPCR Assays. Promega Connections.
Mathieu, M. (2014). 10 Tips for Successful Real Time qPCR. Enzo Life Science. http://www.enzolifesciences.com/science-center/technotes/2015/may/10-tips-for-successful-elisa-assay/
Oswald, N. (2016). 10 Tips for Consistent Real-Time PCR. BiteSizeBio.
S.A. Deepak, K.R. Kottapalli, R. Rakwal, G. Oros, K.S. Rangappa, H. Iwahashi, Y. Masuo, & G.K. Agrawal. (2007). Real-Time PCR: Revolutionizing Detection and Expression Analysis of Genes. Current Genomics, 8(4), 234–251. https://doi.org/10.2174/138920207781386960
Schoales, J. (2019). Ten Tips for Successful qPCR - Behind the Bench. ThermoFisher Scientific.
Sundquist, T., & Saldanha, G. (2012). 10 Tips to Improve Your qPCR and RT-qPCR Results (pp. 15–17). http://www.genengnews.com/gen-articles/10-tips-to-...
