Primer Design

 

Primer design is an imperfect art driven by knowledge of nucleotide binding properties and thermodynamics. Designing good primers is crucial to the success of PCR. Time spent on this can save a lot of time wasted later trying to optimise bad primers. For long PCRs or PCRs from complex templates (eg. soil) the need for excellent primers is even more important. Perfect primer design is often the product of trial-and-error, working from the best picks of an algorithm.

If you are new to PCR or are using untested equipment/consumables, I highly recommend that you start by using well quantified primers from a publication rather than ones you designed yourself. Once you know that everything is working and how a good idea of what success & failure look like - then you are ready to start primer design

General Design Considerations

  • 3’ Homology is much more important than 5’ Homology - You need 15-20 bases at the 3' end of the primer that are absolutely conserved in the target sequence, but you can add stuff at the 5' end (up to another approx. 30 bases) that may or may not be found in the target sequence. This is why we’re able to use PCR to add flanking restriction sites.

  • Primers should have homology to opposite strands of DNA with the correct orientation - Your forward primer should have the same sequence as the target site on the forward strand of the target DNA, but your reverse primer has the same sequence as the reverse complement of the target site. In both cases, the primer goes in the 5’ > 3’ direction. A simple tool to use for reverse complementing sequences is this: http://bioinformatics.org/sms/

  • GC Content - Aim to have primers with a GC content approx 50-60%, and if possible avoid repeat regions (e.g. AAAAAA). High GC regions are also problematic for primer design, since any dimers or hairpins in these regions will be much stronger than in ‘normal’ DNA region.

  • GC Clamp - Aim to end the primer on at least one G or C at the 3’ end – this helps keep the 3’ end of the primer firmly anchored on the template.

  • Matching Tm Value - Your pair of primers should have very similar melting temps (Tm) +/- about 3°C, so they work well at the same annealing temp. You can use the IDT website or Snapgene to check melting temps. Note that the predicted primer melting temperature varies depending on the solution chemistry – the values printed on the primer spec sheets are for DNA in 50 mM NaCl. If PCR is performed using Taq, the optimal annealing temperature for the PCR reaction is about 5°C below this Tm in 50 mM NaCl.  For newer PCR polymerases, e.g Phusion or Q5, the optimal annealing temperature is higher due to a processivity domain which makes the enzyme bind more tightly to the DNA – in these cases, the optimum annealing temp could be same or even higher than the Tm value.

Computation Tools for ΔG

Unfortunately, designing primers that associate perfectly with your template is only half the journey. The second and much more challenging facet of primer design is ensuring that the primers don’t form secondary structures such as self-dimers, cross-dimers, hairpins or turns. When in a single-stranded state, the primers will happily bind with any homologous sequence they find - woe be to you if your primer sees its own 3’ end as a better binding target than your template.

The real art of primer design is in modelling the potential sub-structures that will compete with the primer/template structure - for this, you need a computer. Here is an incomplete list of software that can help you calculate the ΔG of every weird little shape your primer might want to twist into. They are in no particular order as none stand out to me as infallible.

The dimers should have higher (less negative) ΔG values than approx -5 kcal/mol to be acceptable. eg. a primer where the worst self-dimers are -8 kcal/mol would not work well, but a primer where the worst self-dimer is -3 kcal/mol should be OK.

You need to check for both self-dimers and pair dimers - See if the primers anneal to themselves alone, then see if they anneal to each other. A good pair of primers will want nothing to do with each other.

For hairpins, the cutoff is about -2 kcal/mol - Any length of DNA that routinely forms a hairpin with itself will be a rubbish primer.

If your primer has bad dimers or hairpins, there are a number of ways to deal with this problem. The first step in all cases is to identify which particular bases are problematic (using the software/websites mentioned above). Then you can try the below:

  • Slide the primer location up or down a little bit to move it away from those problematic bases. This only works if the problem bases are near the ends of the primer. if the problem bases are in the middle of the primer, you will probably need to choose a new priming location

  • If the problem bases are towards the 5’ end, you can sometimes mutate them to another base which doesn’t cause the dimer/hairpin to form. you need to be very careful with this approach though that the mutations you introduce don’t cause problems later.

  • You can (at your own peril) ignore the hairpins and dimers and just order the primer and see what happens… sometimes primers that look bad in silico work OK enough in the lab.

  • If the problems are arising from the bases in a restriction site, can you change this to a different restriction site?

  • If the problems are arising from your extra bases at the 5’ end that are added to facilitate digestion, you can change these to any bases you like (they get cleaved off after digestion)