In this webinar you will learn:
- About the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines
- How to optimize your qPCR and multiplexing protocols for rapid and reliable results
- How to leverage technology to improve your PCR analysis efficiency
Real-time reverse transcription quantitative polymerase chain reaction (RT-qPCR) has been indispensable in the fight against COVID-19. However, there happens to be numerous falsehoods, misconceptions, assumptions, and exaggerated expectations with regards to its capabilities, performance, and usefulness.
PCR today is an essential tool for countless industries and research applications. Utilizing the biological DNA replication machinery, PCR technology amplifies small fragments of genetic material to facilitate downstream analysis and processes.
This molecular technique was recently thrust into the limelight when the World Health Organization (WHO) declared the outbreak of COVID-19 a pandemic, and PCR testing became the preferred technology for detection of the disease-causing virus SARS-CoV-2.
It has also been leveraged in other aspects of disease control and prevention; screening for variants and environmental monitoring are just a few ways scientists have used PCR to combat this global crisis. It is, therefore, essential that the true strengths and limitations of PCR are restated in the context of the current COVID-19 pandemic.
Over the course of the last twelve months, demand for PCR-based techniques and PCR capabilities has skyrocketed. So, whether you are an experienced scientist or a newbie in the lab, learning how to optimize your PCR assays and multiplexing protocols could benefit both your productivity in the lab and the quality of your results.
Antylia Scientific™ (formerly Cole-Parmer) offers a range of innovative tools that can be leveraged for rapid, reliable, and reproducible PCR analysis.
Meet the presenters
Dr. Stephen Bustin – Professor of Molecular Medicine at Anglia Ruskin University
Stephen Bustin is Professor of Molecular Medicine at Anglia Ruskin University in the UK, where his research interests center around developing novel approaches for the early diagnosis of infectious diseases. He has authored numerous papers, review articles, and book chapters aimed at improving the reproducibility and robustness of molecular methods, especially those based on the use of PCR and has presented hundreds of talks and workshops worldwide on this subject. He has published three books, the “A-Z of Quantitative PCR” (2004), “The PCR Revolution” (2011) and “PCR Technology” (2013). Professor Bustin was an expert witness advising the UK High Court and the US Department of Justice on PCR technology in the Measles Mumps Rubella (MMR) vaccine - Autism class action as well as at the MMR trial in Washington DC in 2007, respectively, and has acted as an expert witness in several international patent and criminal trials. He led an international consortium developing the MIQE guidelines for the use and reporting of real-time (2009) and digital PCR (2013). Professor Bustin has extensive editorial involvements as Editor-in Chief (Gene Expression), International Journal of Molecular Sciences, and as a member of the editorial boards of several peer-reviewed journals. He acts as a consultant to advise a number of UK-based and international companies.
Madalina Enache – Product Sales Manager (EMEA) – PCRmax®, Techne®, and Argos Technologies®
Madalina holds a BSc (with Hons) in Biology and a MSc in Microbiology. She joined Antylia Scientific™ three years ago, when it was known as Cole-Parmer, as part of the application sales team. She has risen through several commercial roles, focused on PCR products, to become the EMEA Product Sales Manager for our PCRmax, Techne, and Argos Technologies brands.
Questions & Answers from the live event
Regarding sample preparation – is it always necessary to perform a complete extraction and purification of the sample, especially if the sample has been taken for rapid testing? If not, what effect might it have on the result? No, it is not always necessary for diagnostic purposes if no quantification is required. If appropriate controls are included and the expected level of sensitivity is achieved, then there are methods and reagents that allow an RT-qPCR assay to be performed without extraction. For quantitative analysis, I would not recommend this.
Why do some primer pairs result in a wider spread of Cq values? If there is a secondary structure at one of the primer binding sites, then it could cause the primers to hybridize with unequal efficiency. The sequence of the primer itself may matter, but it is not predictable without performing the wet lab analysis.
As well as primer sequence, will relative primer concentrations influence efficiency, and should you optimize this as well? Primer concentration can affect the reaction, especially if the primers are poorly chosen. But a generic 200 to 400 nM (SYBR) and 300 to 600 nM (probe) primer concentration usually works well. If the template is tricky and poor primers cannot be substituted, then of course it can be important to assess the effects of changing the concentration of the primers.
It is concerning that different lots of commercial reagents can vary considerably. Do you propose that a validation test is performed each time new reagents are used? I perform a validation test each time new reagents are used.
Can the ramp rate of your instrument affect the denaturation and polymerization times? It depends. Most of the denaturation and polymerization occur during the ramping. Check Carl Wittwer’s paper on this. The PCRmax® Eco 48 Real-Time PCR system is very fast, and I know of another competitive brand real-time system that is slower, but both work with one-second denaturation and one-second annealing/polymerization. The reagents make the biggest difference, as does of course the G/C content of the template.
Changing the threshold position can change the Cq values. Do you have any guidelines on how much or little the threshold should be changed when analyzing the data? Not really, but most people tend to keep the same threshold for diagnostics. For quantification, you need to include standards so you can relate all your Cqs to one another. If your assays are 100% efficient, the slopes of the amplification plots will be similar. Then, it does not matter where the threshold is drawn when you do your ∆Cq analyses.
Do you have a recommendation concerning spikes as internal controls and their accuracy? I use RNA spikes for RNA and DNA spikes for DNA, and quantify them using ddPCR. If RT and PCR efficiencies are similar, they work well. However, I have noticed that RT reactions on synthetic RNA spikes are variable, hence it is important to include proper error propagation calculations with your results.
Can one use normal autoclaved pipette tips instead of RNAse free ones? How big is the mistake? I would never do that because you cannot guarantee removal of all condensation from the tips, and if you are pipetting µL volumes, you will be inaccurate.
What do you mean by high RT variability: a difference in results between different lots of RT from the same supplier or high differences in results between suppliers? Try it yourself. Take RNA and carry out four RTs using random primers, which you then amplify separately. You will see that different RTs end up recording different Cqs. This can be minor or major, and is assay, template, and protocol dependent. I have not tested different lots of RNA for variability.
In terms of data analysis, could you please comment on the effectiveness of the Fourier transform approach and possibly indicate a few references to read, thank you. I always use standard curves, for reasons I have explained in numerous papers. For alternative approaches, read papers by R.G. Rutledge and J.M. Ruijter.
When designing primers, do you have any suggestions on the GC content percentage? Depends on target. Fungi have high G/C content and so you need to denature for longer, but I have not found much of a difference between a 50% and 60% G/C content primer. I try not to have runs of G/Cs.
When choosing a region of the target to amplify, which one will work better, short or long PCR products? Any suggestions? I always choose very short amplicons (60 to 90 bp). I want to complete the run as fast as possible.
What are the most important factors to design a fast protocol? Factors include short amplicon, fast enzyme (KAPA, Bioline), and SYBR if possible.
What is the sample volume for the ECO 48 real-time PCR system? I use as little as 1 µL, but standard volumes in my lab are 5 µL.
If the commercial reagents can vary so much, can they be useful in clinical diagnostics? They all work for detection. The problem arises for quantification.
When doing bacterial ID, how do you correlate PCR results to CFUs? You would need to experimentally determine a correlation between qPCR results and microbial growth. CFUs will only be live bacteria whereas qPCR will detect DNA from dead cells as well. To avoid this, you would need to grow the cells then perform a serial dilution of the culture. Perform PCR on each dilution and plate out the cells to get the CFU. You will also be assuming one copy of target per cell, which may not be accurate. See this reference:
I am having a hard time getting tips. I have tips that are RNase and DNase free, but they are coming in bags and I have to box them myself. Should I avoid autoclaving them after I fill the boxes? Essentially, can I fill a box and then use the tips? We don’t recommend using autoclaved tips, because it can lead to inaccuracies in pipetting. Also, for PCR, filter tips are recommended to prevent pipette contamination and you cannot autoclave filter tips.
When reducing the amount of master mix, do you reduce the overall reaction volume or dilute the master mix with some sort of buffer to maintain optimal reaction volume? You can top up with water to same reaction volume.
For master mix variation, are you saying this applies to Ct results or calibration for copy numbers using plasmid too? I don’t target plasmids. This variation is only a problem for quantification, not for detection.
If different master mixes provide different results, how do we choose the right master mix? Master mixes provide different quantitative results, so you should use more than one master mix and include the error calculations, so the final result is not a 3-fold difference but a 3-fold
(1.5 to 7-fold) difference.
When assessing performance, what should we look for when speeding up cycling? Do amplification inhibitors increase their effect with shorter cycles? You need to check that the efficiency of the reaction does not suffer. You can do this by running a series of standards under all the conditions and checking the slope of the curve. Also, the sensitivity may be lower, so you may also need to check your LOD and LOQ.
Generally, my customers request a conventional 96 position PCR system. How can I suggest a real-time 48-well system? The Eco 48 Real-Time PCR system, having a small block, allows very rapid PCR run times. With smaller sample volumes, it saves on reagents. It is possible to throughput the same number of samples in the same amount of time as a conventional 96-well qPCR instrument. The included software also allows you to combine several plates of data together for analysis if you can’t fit all the samples on a single plate.