Why hot-start is so hot?

Imagine you are preparing your samples for PCR during a hot summer day. You have already mixed all the necessary components together, now you just have to insert your samples in the PCR machine. Suddenly your co-worker shows up and says that they need only a few minutes of your time to discuss a very important matter. As per usual a few minutes then turns into fifteen minutes. You then finally turn to your samples and insert them into the PCR machine. A little while later it’s time to analyse the results, but for some reason they are messy and don’t make sense. You sigh since you have wasted a perfectly good summer day, but gotten nothing out of it. So how to make sure this doesn’t happen again?
First let’s talk about what went wrong. Presuming that there was no contamination and all the primers and PCR mix components were working properly, there was only the issue of letting your samples sit at room temperature for fifteen minutes. During that time your primers probably randomly bound to DNA templates or to each other forming primer dimers and starting non-specific amplification, because there was nothing to stop them from doing that. In the end you ended up with lower yield and non-specific results, since part of the PCR efficiency went to amplifying unnecessary segments [1]. 
To avoid that the easy answer is using hot-start PCR technique. With this technique you don’t have to worry about leaving your samples on the table while talking to a colleague or worry that once you get to the last samples the first ones might already be messed up. The main idea behind hot-start is that the reaction needs a hot start to work [1][2]. There will be no non-specific amplification during reaction setup since the specially modified DNA polymerase won’t work until heated at 95 °C in the PCR machine [1][2]. So using hot-start PCR technique means that not only can you work calmly on your samples at a room temperature, but you will also end up with higher yields, better specificity and sensitivity [1][2].  
There are still some things to consider when using hot-start, since there isn't only a yes and no option. There are many different hot-start variants: antibody, chemical, oligo, wax bead etc., each with their advantages and disadvantages.
  • Let's start with antibody mediated hot-start. A monoclonal antibody is bound to the Taq DNA polymerase's active site and prevents catalysis [2][3]. By raising the temperature the antibody denatures leaving the active site free. The advantage of this is that denaturing the antibody is a very quick process (takes 1-3 minutes) and the antibody doesn't alter the polymerase [2]. The disadvantage is that antibodies can contaminate the reaction when detecting mammalian DNA targets, since antibodies are mostly produced in hybridoma cells of mammalian origin. [4]. Also since animal cells are used for the production of antibodies it makes the production more expensive and less ethical. 
  • Chemically modified Taq polymerase has a small organic compound attached to it, which prevents its activity. The advantage of this is that there are no contamination risks. The disadvantages are that getting rid of the compound can take over ten minutes and during this time the DNA you are researching can get heat-damaged, which is why this method is not good for long targets. Also it may be impossible to completely remove the organic compound from the polymerase and therefore it cannot work in full capacity. [2]
  • Oligo (aptamer) hot-start uses oligonucleotides to block the Taq polymerase until raising the temperature will free the polymerase. In general it works just like the antibody mediated hot-start. The advantages of this method are that oligonucleotides aren't as strong as small organic compounds in chemical hot-start, so enzyme activation will only take around 30 seconds (sometimes longer incubation times are used to also enable longer target DNA molecules to denature). Using oligonucleotides is also more ethical than using antibodies. The disadvantages are that since oligonucleotides aren't very strong, the reaction might not be stable for a long time and there could be some nonspecific amplification. [2]
  • Wax bead based hot-start is probably the strangest technique. It has Taq polymerase inside wax spheres. Once the wax melts around the polymerase it can be used for action. Advantage of this is again that the polymerase activation process doesn't take long. Disadvantage is that using wax is messy, it's not soluble in water and can make removing the amplified DNA difficult. [2]
For your convenience most Solis BioDyne PCR and qPCR mixes already contain reagents required for hot-start. We use chemical and oligo hot-start methods to avoid experimenting on animals. In addition, due to Stability TAG technology all our enzymes and master mixes have an enhanced stability at room temperature with no activity loss up to 1 month, so you don’t have to worry about not using ice while preparing your samples. Also be sure to check out our SolisFAST® line products that are not only thermostable, but so fast that you can repeat your perfect result and still have time left to enjoy the autumn.
For fast endpoint PCR:
For fast qPCR:
Literature used:
[1] Kubu, C. J. (2008). HotStart-IT®: A Novel Hot Start PCR Method Based on Primer Sequestration. BioTechniques, 44(2), 275–277. doi:10.2144/000112827 
[2] Paul N, Shum J, Le T. Hot start PCR. Methods Mol Biol. 2010;630:301-18. doi: 10.1007/978-1-60761-629-0_19. PMID: 20301005.
[3] Dahiya, R., Deng, G., Chen, K., Haughney, P. C., Cunha, G. R., & Narayan, P. (1995). Terms and techniques: New approach to hot-start polymerase chain reaction using Taq DNA polymerase antibody. Urologic Oncology: Seminars and Original Investigations, 1(1), 42–46. doi:10.1016/1078-1439(95)00001-x 
[4] Witt, N., Rodger, G., Vandesompele, J., Benes, V., Zumla, A., Rook, G. A., & Huggett, J. F. (2009). An assessment of air as a source of DNA contamination encountered when performing PCR. Journal of biomolecular techniques : JBT, 20(5), 236–240.