PCR Troubleshooting

It’s funny, but a lot of organisms don’t come ready-made for PCR amplification of their genes or DNA sequences. As you know by now, we have to go through extensive procedures to extract the DNA in the first place. But even after these extractions are complete, the DNA may not be of sufficient purity for efficient amplification.

How can you tell if your PCR has been inhibited? The problem begins when you see no bands in your gel. And in general, that is not a likely result of PCR. Thus, we’ll begin with the most extreme problem: no bands at all, no product.

After excluding obvious technical errors on our part (did we add the primers? the polymerase? the dNTPs?), we turn to the more challenging troubleshooting steps. There can be several reasons for a result of no product, but these are not equally likely.

First, no product could mean not enough template DNA was used. This is possible, but the result is generally weak bands, not the complete absence of bands. Stain the gel longer, then image again. PCR is very sensitive, and amplification of DNA from even one cell is routine.

Second, no product could mean that the primers were not matched to the template/target DNA. However, primer/template mismatch will likely show up as amplification of multiple, weak bands (stuttering).

Third, there could be a problem with the DNA polymerase, either from a nonoptimal amount of key cofactor salts (MgCl₂, KCl) or the DNA polymerase has been exposed to moisture and it has degraded.

Fourth, the DNA we used could be so degraded that few strands are long enough to serve as templates for the primers.

Fifth, inhibitors may be present if we see few, faint bands, where we expect several, strong bands, and in particular, if only small-sized products are present.

Inhibitors imply inhibition of the activity of the DNA polymerase (Kermekchiev et al 2009). For example, in whole blood, hemoglobin is a strong inhibitor of Taq polymerase. In plants like beans, a potential food source for your GMO detective work, the inhibitor problem is likely from the polysaccharides that are present in the seeds.

What are our options?

Removal of impurities from our extractions implies additional steps. One option is to use spin columns that selectively trap the DNA but let smaller molecules to leach out. These work well, but tend to be expensive. Dialysis is also an option, but tends to lose small DNA fragments.

Another option is actually a very simple choice. Instead of using more DNA, use less DNA. This seems counter-intuitive… to use less DNA to overcome a problem of no products in our PCR?

LMP agarose

Here, I will introduce you to another method to deal with impurities, use of low melting point (LMP) agarose. LMP agarose has a low melting point (60 C) compared to 80 C or more for typical agarose used in electrophoresis. LMP agarose is expensive compared to regular agarose (about 10X greater), but it is a far cheaper solution compared to the spin columns. And, we don’t lose DNA! (All spin column approaches will lose DNA at least the smaller fragments). LMP agarose can be present in the PCR reaction tube and it does not seem to interfere with DNA annealing and synthesis in standard PCR. Because the melting point of 50 C is still higher than the relatively cool 36 C we use for the annealing temperatures in the RAPD protocol, we may indeed see some issues with LMP agarose. Either way, this would be an interesting technical result.

Making agarose plugs

1. Find your DNA samples (freezer or ice bucket) and warm the tubes containing DNA/water to 65 C (at this temp, DNA is still double stranded). Use the heat blocks.

2. Add equal volume of melted 1.6% LMP agarose (in TE) to your tube containing your DNA/water — I want 1 ml total volume.

3. Mix well, then quickly withdraw the agarose+DNA into a 1 ml tuberculin syringe.

4. Let the agarose gel at room temperature.

5. Then push the plug into a new microtube. Cut the end off the syringe, use the plunger to gently push the plug out.

6. Add TE to the microtube enough to cover the agarose plug.

7. Leave the tube at 50 C for 24 hours, then remove the TE and repeat for another 24 hours with clean TE.

8. After 48 hours, replace with clean TE — you now have a microtube with an agarose plug in TE.

The rationale behind this is as follows. The TE will leach through the agarose pellet — the DNA is trapped in the gel matrix while the small molecule impurities will wash away. After the TE washes are done, we will have pure DNA in the agarose plugs.

If all goes well, we will run PCR again on your target amplicons next week, in time for you to see results by your next lab. This time PCR will be done to compare amplification of dilute DNA samples not treated for impurities vs. samples taken from the agarose+DNA plugs. Note: TE stands for tris-EDTA, which is a standard storage buffer for nucleic acids.

References

Kermekchiev MB, Kirilova LI, Vail EE, Barnes WM (2009) Mutants of Taq DNA polymerase resistant to PCR inhibitors allow DNA amplification from whole blood and crude soil samples. — Nucl. Acids Res. (2009) 37 (5): e40.).