A Definitive, Practical Guide to Conventional PCR

This is the second installment of my Definitive, Practical Guide series on molecular biology techniques. For the first one on designing primers for PCR and qPCR, see here.

Now that you have (hopefully) read my previous article, designed, and received the primers in the mail, what now? What next? Without further ado, let’s get you these glorious bands that you deserve, shall we?

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 eukaryotes will not be reflected in this guide.

Primers

Let’s start with the primers you just received. Typically, the forward and reverse primers are shipped in the form of desalted powder for stability at room temperature during shipping. Product sheets accompanying the primers will include information such as its Tm, GC content, but more importantly, the volume of nuclease-free molecular-grade water you need to add to reconstitute the powdered primer into a set concentration, typically 100 µM. Once reconstituted, you now should store them at -20 °C for long term storage. However, these native concentrations of 100 µM is too concentrated to be used directly in a PCR reaction and thus, requires further dilution. Add 4 µL of the concentrated primer into 36 µL of nuclease-free water to obtain primers at 10 µM concentration.

PROTIP: If you add 4 µL each of both the concentrated (100 µM) forward and reverse primer into 32 µL of nuclease-free water, you now have a 40 µL volume primer mix of 10 µM of each primer in a compact, ready-to-use single vial. However, this significantly increases the chances of formation of primer-dimers in the subsequent reaction so use this tip at your own risk.

PCR Master Mix

Typically, PCR master mix is a premixed ready-to-use solution containing the Taq DNA Polymerase, dNTPs (DNA nucleotides), magnesium chloride and other buffers at concentrations and ratios suitable for PCR reactions. This will save you a tonne of effort from having to mix in your own polymerase enzymes, nucleotides, and buffers. The author strongly recommends purchasing one commercially. They typically comes in 2X concentration. What this means is that, if the final volume of the entire PCR reaction is 20 µL, then half of this should be the mater mix (i.e. 10 µL) so that its final concentration will now be 1X. They also comes pre-mixed with the loading dye like the GoTaq Green Master Mix, which happens to be the author’s favourite master mix.

GoTaq Green Master Mix (2X) comes premixed with loading dye which allows the contents of the PCR tube after the reaction to be directly loaded into the wells during gel electrophoresis without diluting it with loading dyes. Source: Google image search.

DNA Template

Obviously, if you are doing PCR, you will need your source of DNA. Being a microbiologist myself, I will limit the discussion to microbial DNA, specifically, bacterial DNA.

Commercial DNA Extraction Kits

There is a plethora of bacterial extraction kits such as this which will do a mighty good job at giving you clean and pure bacterial DNA that is even good enough for long read whole genome sequencing. However, for routine convention PCR where you just want to know if the gene for which you designed your primer for is present, you honestly do not need to waste time on using commercial DNA extraction kits.

Colony PCR

Colony PCR is the practice of using a particular, well-isolated single colony from the agar for subsequent PCR as opposed to a test tube full of homogeneously mixed bacteria cells. There are two main ways of getting DNA from a single colony in colony PCR.

Colony PCR investigates individual colonies on an agar via PCR. This is useful when a mixed culture of bacteria is involved where the bacterial cells growing in the same agar are not necessarily identical. Source: Animated Biology with Arpan.

One way is to use a pipette tip to touch a single colony of interest on the agar and then suspend it directly into the PCR mix and let the heating during PCR to do the “extraction” of DNA from the live bacterial cells. This is also less commonly called the “Touch PCR”. However, Touch PCR has a few serious limitations. The live bacterial cells often end up inhibiting the PCR reaction and may cause failure to amplify the gene even if it is present, especially if you end up using too many bacterial cells in too little a volume of PCR reaction mixture (like 10 µL). Secondly, if your bacterial colonies have been grown on selective and/or differential agar like say, Salmonella spp. on XLD agar, the bacterial colony may not be “pure” enough for PCR. For instance, Salmonella spp. growing on XLD will form black centers due to H2S production. If you use these Salmonella XLD colonies directly for PCR, the iron sulfide in black color center from the colonies will inhibit the PCR reaction. Hence, it is important to only use colonies from non-selective agars such as TSA, LB or nutrient agar for Touch PCR.

For these reasons, the author prefers the second method which involves performing a rough extract of DNA from the colony by heating in sterile DI water first before using it in PCR. Take a 2–3 mm sized, well-isolated single colony from a preferably non-selective agar and resuspend in 500 µL of sterile DI water. Heat the suspension at 98 °C for 10 min, cool on ice (preferred) or room temperature water for 5 min, followed by centrifugation on a bench top centrifuge for 5 min (exact speed matters less). All the cellular debris and other potential PCR inhibitors will sink to the bottom of the tube, leaving behind a relatively much cleaner DNA extract that is now suitable for PCR. This method also works for “unclean” colonies on selective agars because all the potential inhibitors like bile salts or iron sulfide etc. will be significantly diluted in the DI water.

PCR Mixs & Typical Recipes

The PCR mixes will have the following typical recipes as depicted:

Typical PCR mix recipes as viable starting points. Author recommends that the order of reagents to be added be from top to bottom. There are very technical reasons for why the PCR master mix, for example, should be added last/just before PCR that the author has read in a textbook somewhere long ago. Or maybe the author might have dreamed it up, who knows. Source: Author’s work.

Noteworthy Pointers (in no particular order)

1. In Touch PCR, the PCR mix is created without allocation for DNA template volume. Intact bacterial cells are inoculated directly into the mix.
2. In the author’s experience, using a full 8 µL of DNA template without any nuclease-free water in standard PCR leads to inefficient amplifications or sometimes, even failure to amplify, plausibly due to PCR inhibitors or too high a DNA concentration. Start with 1:1 ratio of water to template first, and adjust accordingly in subsequent optimization, if needed.
3. It is practical to make a batch of the PCR mix for the total required number of samples and then dispensing 16 µL each into separate PCR tubes later rather than individually mixing all the components in the PCR tubes.
4. PCR master mix should be thawed on ice and added last but before DNA template. This minimizes premature DNA polymerase action before commencement of PCR.
5. The physical space to prepare the mixture of nuclease-free water, primers, and PCR master mix should be located far away and separately from the DNA template adding area to avoid contamination of DNA template into subsequently prepared PCR mixes.
6. If the forward and reverse primer mix is used as described above, the volume of primer mix should be 2 µL as opposed to individual 1 µL portions of each primer.
7. When adding the PCR master mix into the water + primer mix, pipette at least 20X up and down for thorough mixing. Why 20X? An old textbook recommended it and it has not failed me since, so I’m passing along.
8. Ensure that the final PCR mix with DNA template is thoroughly centrifuged to prevent air bubbles or leftover reagent on tube walls before PCR.
9. A total volume of 20 µL is recommended as this allows two gel electrophoresis runs of 10 µL each. In the event that the first one goes wrong from improperly casted gel or other random occurrences, you will have backup without having to repeat the entire PCR. Throw away the 10 µL backup only after you have successfully obtained the bands.

PCR Conditions and Run Parameters

Typical PCR run parameters are as follows:

A typical PCR run parameters as a good starting point. Source: Author’s lab records.

Noteworthy Pointers (in no particular order)

1. In Touch PCR, where live intact bacterial cells are used as template source, the duration of the initial denaturation step can be extended to 5 min to more completely lyse the bacterial cells.
2. High temperatures of 98 °C has been used in some protocols but is not recommended as it may cause denaturation of the Taq polymerase. If 98 °C is to be used, significantly reduce the duration of initial denaturation and subsequent denaturation step to 30 s and 10 s, respectively. Reference: Zwe, et al. Food Control 90 (2018): 233–240.
3. Annealing temperature of 51 °C is used as the Tm of primers have been specifically designed to be at 56 °C. In general, the annealing temperature should be 5 °C below that of the Tm of primers being used. See the first part of this guide here for more information on primer design.
4. The duration of the extension step should be at a rate of 30 s per 500 bp of the amplicon. Therefore, if a PCR product of 1000 bp is expected, the duration should be increased to 1 min. However, for amplicons < 500 bp, a minimum duration of 30 s is still recommended.

Conclusion

Successfully performing your first PCR reaction is a rite of passage for any aspiring researcher in the field of molecular biology and life sciences in general. I hope this little guide on PCR has benefitted you. Thank you, dear reader, for your time and attention, and I wish you the best in your academic endeavors.