A Definitive, Practical Guide to Designing Primers for PCR & qPCR.
This is the first installment of my Definitive, Practical Guide series on molecular biology techniques.
What if I told you that I earned myself an entire PhD without ever intricately understanding the principles of designing primers for PCR and qPCR at a fundamental level? I blame it on the web applications such as NCBI Primer BLAST, which allows us to too easily construct primers that will at the very least give us a workable reaction most of the time without ever examining us on the underlying principles of any of the parameters and values even mean. Other times, we copy a published protocol and the set of primers wholesale.
And I get it. As PhD students juggling work, relationships, and other aspects of life (*cough* your empty wallet… *cough*), a PCR/qPCR reaction that works is way more important than why it works. Admit it; usually, things work, and we don’t know why, nor do we care! However, sometimes, things DON’T work. But I’ve come to realize that these instances more often than not turn out to be blessings in disguise. It is precisely when the AI-designed primers are not working or when replication of a published protocol fails to yield the result you are looking for that you are forced to troubleshoot by learning the entire process at a fundamental level. It is when learning happens, and you grow. This article is a record of my learning journey, published in the hope that you will find it useful for your journey as well.
NOTE: Take this as practical, unsolicited advice from a lab senior rather than as a peer-reviewed protocol, i.e. it’s a collection of the rule of thumbs, tips and tricks, and what has worked for me, as opposed to a systematic and complete review of the science with highly substantiated references. Also, being a microbiologist myself, I principally work with prokaryotes or viruses. Hence, considerations that are unique to designing primers (like introns) for eukaryotes like humans, animals, fungi, etc. not be reflected in this guide.
What will the primers be used for?
The first step of designing the primers is to question whether they will be used for the conventional Polymerase Chain Reaction (PCR) or Quantitative PCR (qPCR). Real-time PCR (not to be confused with reverse-transcriptase PCR typically annotated as rt-PCR).
As this is not a guide on the different types of PCRs, I will be brief. Refer to here for more information.
A conventional PCR involves amplifying a segment of a gene of interest, staining it with DNA stain and then visualizing it by gel electrophoresis. It is chiefly used to determine if a gene is present and, therefore, amplified to visualize an end-product of known size in gel and vice versa. The size of the PCR product/amplicon is entirely within your control. Typically, primers are designed to yield PCR products ranging from 100–1000 bp. A DNA ladder of known sizes is run alongside in the gel to identify the PCR product size. If you designed the primers to amplify a 500 bp portion of the gene of interest but yielded an 800 bp band or multiple bands of 100, 1200, and 3000 bp in gel electrophoresis instead, you know something is up. You may also design PCR products of different lengths for different genes and run them together in the same run for easier identification of different gene products. For instance, primers can be designed for Gene 1 and Gene 2 to yield 150 and 400 bp products, respectively, if present. They can be differentiated very clearly in the subsequent gel electrophoresis step.
On the other hand, qPCR introduces the quantification component, hence, named quantitative PCR. It tells you if a gene is present and how much by correlating the amount to florescence detected by the machine. Another use of the qPCR machine is to quantify gene expression levels (mRNA levels) in which mRNA is first reverse transcribed into DNA, after which it is amplified and monitored (reverse transcriptase PCR). The size of PCR products in primer design for qPCR is typically much smaller at about 90 to 110 bp than conventional PCR. In particular, when designing multiple primers to monitor the gene expression of multiple genes in the same run, you have to make sure that the PCR products of all the genes are very similar in size (~100 bp), unlike conventional PCR, so that florescence dye will similarly bind to all the amplicons from different genes, making them comparable.
Depending on the application, you now know exactly what the size of amplicon you want for your primer design.
Which one’s forward and which one’s reverse?
At the end of the day, since both forward and reverse primers are added into the reaction mix, does it matter if we label forward as reverse and vice versa? How do we know which one’s which? Truth be told, it doesn’t matter 99% of the time. But there is a rare 1% of the time when it does matter, like genetic recombineering.
The forward primer is found on the plus strand, while the reverse primer is on the minus strand. The plus-strand is also called the coding strand or the sense strand, while the minus strand is called the template strand, non-coding strand (rarer in my experience), or the anti-sense strand. So, how do we know which one is which?
The plus/coding/sense strand is the strand that corresponds directly to the mRNA transcript of the gene (except, of course, T will be U, but you know that, right? If not, you are in the wrong article!). For more information, see here.
With that in mind, consider the example below.
Consider for a moment that you are trying to create a primer pair to amplify a 250 bp segment of this gene using the NCBI Primer BLAST tool. My question is, which sequence of the gene will you input as PCR template? The plus-strand sequence (left) or the minus strand sequence (right)? I hope you realize that depending on the input, the software will be giving you wildly different outcomes. In one case, the forward primer is the actual forward and reverse the reverse, while in the other, forward is reverse, and reverse is forward.
And the answer is, you make sure you are using the plus strand as input to the software to yield the correct forward and reverse orientations of the primers. Always look for your start and stop codon, and if you see them in your sequences, you are good to go. The start codon is pretty much always ATG, while the common stop codons are TAG, TAA, and TGA.
Melting temperatures (Tm) of the primers: are my primer sets compatible?
Have you ever run a PCR reaction involving 5 different sets of primers for 5 separate genes in 5 tubes, expecting bands for all but realizing that only 3 showed up? You probably have compatibility issues among your primer sets.
By running 5 primer pairs in the same PCR run, you are subjecting the same PCR protocol (temperatures and times) to all 5 primers pairs. For example, your PCR protocol dictates that the annealing temperature will be 51 °C for 30 secs. But what if I told you that only 3 pairs out of 5 work properly at 51 °C while the other 2 which failed to amplify needed 55 °C. That, in a nutshell, is the primer compatibility issue.
Every primer has a parameter called the melting temperature (Tm). Technically, it is the temperature at which 50% of the primer will dissociate from its DNA duplex form to become single strands.
As a rule of thumb, all the primers used in a single PCR run should have as similar a Tm as possible for best results. In general, the Tm of primers should be between 52 to 58 °C. Personally, I try to design primers with a Tm of 56 °C and no more than 1 °C difference between any given primer used in the same PCR run.
The NCBI Primer BLAST tool does supply us with the Tm, but I found it to be inadequate because it does not take into account the live reaction conditions like primer concentration, Mg2+ and Ca2+ ion concentrations into account. A better tool to determine the actual Tm in a practical PCR reaction condition is the OligoAnalyzer by IDT (need an account). You also need this tool for the next part.
Secondary structures: are my primers designed to fail?
The primers are designed to obediently bind to your DNA template and subserviently amplify the amplicon you want, yes? Sometimes, however, the primers take on a mind of their own and rebel against their creator in a manner not dissimilar to the great rebellion of angels in heaven against God. Do you know what the mark of defiance in primers looks like? It’s…. a hairpin loop!?
The OligoAnalyzer tool, in addition to determining realistic Tm, also allows the visualization of the potential hairpin loop secondary structures that your primers may form. In my opinion, the critical parameter you have to watch out for is the highest Tm of the possible secondary hairpin loop structures. The lower the Tm of these structures, the easier it is to dissociate into the free and available single-strand primer. You have to ensure that the highest Tm of the structure is well-below (<10 °C) that of the annealing temperature you are using. I once mistakenly used a primer whose hairpin Tm of 56 °C was as high as its native Tm of 56 °C (i.e. its hairpin loop structure was equally strong/hard to dissociate than its native duplex structure). At an annealing temperature of 51 °C (5 °C lower than native Tm), the primer was happily binding unto itself. Needless to say, I did not yield good results and did not know why until I learnt of this phenomenon.
Another typical form of secondary structure that plagues many a PCR reaction is called the primer-dimer. This occurs when a 3' end of a primer binds to its 3' end, which is then extended by DNA polymerase to become a duplex product. A picture speaks a thousand words, and I found a very concise and clear illustration of the phenomenon, as illustrated below.
How do you avoid this? The OligoAnalyzer tool allows one to check the self-dimerization potential of your primer. I have not developed a rule of thumb or cut-off values of ΔG, which I can confidently recommend here yet. Use your discretion and make sure the primers won’t easily form primer-dimers based on the sequences.
You can tell that your PCR reaction is being plagued by primer-dimers if you see visible bands of very low molecular weight in addition to the expected band.
Conclusion
I hope this little guide on designing PCR primers have been helpful for you. This is by no means exhaustive, but I hope it can definitively get you started on the right track. Thank you, dear reader, for your time and attention, and I wish you the best in your academic endeavors.
