What’s with hot start, anyway? A tale of mispriming and how to avoid it.

Nov. 25, 2019

Ever looked at a PCR thermocycle program and noticed it has a prolonged initial high temperature step before the repeated cycling occurs? In this month’s episode, we’ll delve into what that’s for, and the several ways in which the same effect can be achieved.

Primer, meet template

If the whole mechanics of the PCR process aren’t fresh in your mind, you might want to dip back in the archives to the April 2013 installment of this series, “PCR: the basics of the polymerase chain reaction.” Key points here are that the process requires synthetic primers, or at least their 3’ ends, to anneal down onto a template to recruit DNA polymerase in and commence template-directed elongation of the primer. That primer annealing is a balancing act between random thermal motion of the primers and template (attempting to tear them apart) and bonding energy (primarily H bonds) between primer and template (attempting to hold them together). As temperatures go lower, less bonding energy is needed to tip the balance towards primer annealing; this means that shorter sequences, or in this case, less than all of the primer sequence has to match its complement on the template to anneal.

Imagine then that within a reaction tube, we have all the components needed for PCR—template, buffer, dNTPs, primers, and polymerase—and that we’re going to be manipulating this at room temperature, which is where most lab techs like to work. Most PCR primers have annealing temperatures well above room temperature, often in the 50-65° C range. Put them at 21° C and guess what; they only need to match about half their length to effectively hybridize onto a template. If that half length is towards the 5’ end, or even in the middle—in fact, anywhere but including the 3’ end and its critical hydroxyl—this doesn’t create a substrate for polymerase to do anything. So aside from the fact that this interaction effectively sequesters some primer in non-productive locations, it’s not very harmful. If, however, the 3’ end of the primer is annealed down, a functional substrate for polymerase is formed, and it may be grabbed and nascent strand elongation begins. Your previously carefully designed, highly target specific primer is now starting to amplify someplace it shouldn’t.

At this point, I suspect there’s murmurs of discontent in the gallery. “But PCR uses thermostable polymerases, usually from thermophilic organisms, and they don’t work at ambient room temperatures!” Well, that’s not quite true. Yes, they generally work much better (faster, more processivity) at their evolved optimal temperatures, but they still can catalyze DNA synthesis from a proper template, slowly, at lower temperatures. How much slower probably depends on the buffer, the enzyme and the actual ambient temperature, but it’s rather a moot point. Even if the non-specifically annealed primer is only extended a handful of base pairs, its now created a modified, elongated primer in which the initially mispaired region plus the extended section now are a perfect match for someplace undesirable in the template. That’s right, now even at elevated temperatures designed for specific annealing of the initial primer, this modified primer can anneal it’s lengthened 3’ end and effectively recruit polymerase. In fact, it can’t act to prime amplification from its intended target anymore, as its 3’ end doesn’t match the correct target.


Oh, it gets worse. If our undesired, single strand of misprimed product is able to find any floating DNA fragment capable of annealing to it, that in turn can prime synthesis back across what was the original primer. This creates a perfectly good binding site for any functional original primer, which now competes with the actual target sequence. While this might not seem a highly likely occurrence, recall we’re dealing with literally billions of base pairs of template sequence in the average sample, and statistics is no longer our friend here. Bottom line, in reality, if you let your polymerases sit around in an active state with primers and template at temperatures well below the primer annealing temperature, you tend to lose assay sensitivity (primers get sequestered and rendered unusable), and you lose specificity (modified primers start priming amplification of off target garbage). Obviously, this is a bad thing, and should be avoided if possible.

Solving the problem: hot start

The techniques used to circumvent this problem are generally lumped together under the title of “hot start” methods, meaning the PCR reaction isn’t made wholly functional until the reaction is well above intended primer annealing temperatures for the first time. No functional reaction mix means no modification of misprimed primers and we get a fresh, clean start as the reaction comes to the first programmed annealing temperature. This can significantly improve sensitivity and specificity, so let’s look at some of the ways to do this. For completeness, we’ll cover some methods no longer in vogue, as well as the most commonplace methods in the lab today. 

First approach: set up the reactions on ice, and don’t put the reaction tubes into the thermocycler until it’s well above annealing temperature. Wait, what? Doesn’t putting the reaction on ice make the primer mispriming issue worse, making it need even less nucleotides at its 3’ end to match something to anneal down? Yes, that’s right, but the trade-off is that we’re getting further away from the optimal activity temperature of the polymerase, and it may be that it’s so sluggish at these temperature ranges that it just really hasn’t got much or perhaps any chance to catalyze primer extension.

Lower temperatures also require less kinetic mixing in the sample, meaning productive collisions between a polymerase and (unwanted) substrates occur less frequently. Overall, while this doesn’t unequivocally solve the problem, at least it greatly reduces it. By not placing the reaction tubes into the thermocycler until it’s hot, the small volumes of each reaction increase in temperature rapidly, meaning there’s not much window of opportunity for both primers to be misannealed and for polymerase activity to occur. This method is probably the crudest approach to hot start; it requires hands-on activity and it’s not perfect. But it’s cheap and simple and at least it helps (empirically, sometimes quite a lot).

Second approach: don’t make the reaction contents complete until the mixture gets hot. Variations on this approach hinge on encapsulating critical components (either the primers themselves, or sometimes, the divalent ions such as Mg2+ needed for DNA polymerase catalysis) in something like a wax bead sitting in the reaction tube. This wax has a melting point somewhwere above usual primer Tm ranges but below 95°C, such that the reaction doesn’t become complete until it’s above temperatures where mispriming can occur.

In the dawn of time from a lab PCR standpoint, before heated PCR machine lids or “hot bonnets” were common, this had an additional advantage in that after melting, the wax would float to the top of the reaction and serve as an evaporation barrier. You no longer had to drip mineral oil on top of each reaction vessel to serve this role as part of the reaction setup. While a clever approach (and one still in use in some settings) downsides of this are either that you have to custom-formulate primers into the wax beads for every assay (with appropriate control of concentrations and bead size per reaction), or for the divalent cation beads, lose ability to optimize cation concentration as the beads are premade. (A solution to this, of course, could be to vary the reaction volume such that the premade bead cation content provides the desired final optimum, so it’s not an insurmountable challenge.) Overall though, these wax-embedded reagent methods aren’t overly commonly encountered. 

Third approach: involves doing something to the DNA polymerase itself such that it’s actually unable to function prior to being heated well above primer annealing temperatures—a heat activation step. Since polymerases don’t naturally come with an on/off switch, clever folks in the biotechnology industry have had to come up with a way to do this. Patents being what they are and human ingenuity being what it is, at least two different strategies to do this have been developed and both are referred to as “hot start polymerases.” One of these works by developing thermolabile monoclonal antibodies which have a strong affinity for the DNA polymerase active site, and effectively block the site from accepting substrate while bound. (Nothing says this has to be an antibody; another engineered polypeptide could work as well, but it’s probably technically easiest to do this via antibodies.) 

During polymerase formulation, after it’s purified, it is mixed with an excess of this blocking antibody and binding is allowed to proceed to completion, yielding intact but inhibited polymerase. Subsequent handling of the reagent is such that the antibodies remain bound right through until final reactions are made. When this sample is heated, however, the importance of the antibody being thermolabile becomes apparent; at some temperature above expected primer-specific annealing, the blocking antibodies denature or unfold. This means they no longer have affinity for the polymerase active site, detach and float off into solution. This denaturation is generally thermodynamically irreversible, meaning they won’t come back to interfere with polymerase activity, regardless of subsequent thermal profiles. Hot start polymerase formulations based on this approach generally have fairly short hot start cycle requirements, on the order of less than five minutes.

The second route to this is via a reversible chemical modification of the polymerase, such as esterification of a critical active site hydroxyl (-OH) side chain. This modification completely abrogates enzyme function, but the blocking agent and buffer composition are chosen to make the blocking reaction chemically reversible at elevated temperature. Similar to the idea with antibody blocking, during reagent formulation the purified enzyme is treated with excess blocking agent to leave little or no active enzyme. After final reaction constitution, in a hot environment (e.g. 95°C), the blocking agent is hydrolyzed off to regenerate the critical side chain moiety, and enzyme activity is regained. Compared to conditions during formulation, concentration of released blocking agent is low. That, combined with at least a brief high-temperature denaturation phase in every cycle, acts to ensure enzymes don’t become re-blocked. These chemical-based hot start polymerases usually suggest a somewhat longer initial hot start phase to each reaction (~10 minutes).

Do you always want to hot start, a hot start polymerase?

What if you use a hot start enzyme but dispense with the normal extended hot incubation on your reaction? In this case, the enzyme activation occurs more slowly over each of the cycle denaturation steps, with release of a little more active enzyme each time. This timed enzyme-release approach can actually be desirable sometimes—if only small amounts of template are expected in early reaction cycles, little functional enzyme is needed to sustain effective PCR, and it limits the amount of unoccupied functional polymerase, “wandering around the tube getting in trouble.” This approach usually must be offset by increasing the total cycle number however, which can lead to its own problems. So selecting between this and a longer enzyme activation pre-step is another aspect best left addressed during assay optimization and validation steps, and before general use. 

This third approach, in one of its two guises, is by far the most common hot start method seen in the average laboratory. It allows for ease of ambient sample and reaction handling while avoiding spurious side reactions. Thus, the next time you’re looking at a reaction thermal profile you’ll now know why so much time (sometimes as much as one-third of the total reaction time) is spent in a hot preactivation step, and what this is doing to maximize your assay utility.