Can I directly sequence a PCR product?
Direct Sequencing of PCR Products
It is quite possible to directly sequence a PCR product without first cloning the fragment. Indeed, there are some distinct advantages to this approach. However, you need to be aware of some of the drawbacks as well. Direct PCR sequencing is rarely successful unless you spend some time ensuring that you aren’t falling into one of the many traps. This document will explain how to get sequence directly from a PCR product with a reasonable chance of success.
Make SURE you amplified the right fragment.
You would be amazed at how often a PCR reaction SEEMS to produce the correct band, when in fact it has amplified something spurious. Sometimes when you sequence the band, you will discover that the sequence is completely unexpected and nonsensical. At other times, sequencing with one of your PCR primers will give a completely blank lane, while the other primer will give two simultaneous and superimposed (and thus unreadable) sequences. That happens when only one of your primers acted as *both* ends of an illegitimate amplification.
Make sure you really are amplifying the fragment you expected. For example, if you know of a restriction site in the fragment, try to cut it and look for the correct product bands. Alternatively, use nested primers to re-amplify the desired product to verify its identity and coincidentally eliminate any illegitimate products.
You must remove all residual PCR primers and unincorporated nucleotides.
Sequencing uses one primer, while PCR utilizes two. If we try to sequence with two primers present, you’ll get the two sequences back, superimposed on each other and completely unreadable.
There are many ways to purify a PCR reaction prior to sequencing it. Several manufacturers make kits to do this task. We prefer to not recommend specific products, but ask your neighbors for their advice. Some people just gel-elute the PCR band, which not only removes the extraneous primers and nucleotides but also eliminates illegitimate PCR products (see below).
If the PCR primers will also be the sequencing primer(s), make sure they match our conditions.
You may be able to adjust your PCR conditions to optimize reactions, but we, unfortunately, cannot do this. Please make sure your primer(s) are appropriately designed for automated sequencing. They should have a Tm between 60 and 70 degrees (or at least between 55 and 75 at the outside) and should have little propensity for primer-dimer formation. Click here for complete information on primer design for DNA sequencing.
Don’t over-concentrate the sample! Double-check template the concentrations.
PCR fragments are smaller, and thus more effective sequencing templates that are the usual plasmids. Consequently, they don’t need to be at as high a concentration. For example, a 200 bp fragment only needs to be 1 ng/ul! If your samples are at too high a concentration, not only will they NOT sequence any better, they may cause problems for other peoples’ samples nearby. PLEASE estimate your PCR product concentrations on an analytical gel, and dilute them according to recommendations.
Inefficient primers are sometimes OK for PCR, but the same primers may fail in sequencing.
Because PCR is intrinsically an exponential process, and because it is usually carried well beyond completion, even rather poor primers will produce amplification in a PCR reaction. Sequencing. however, is strictly linear, and is much more unforgiving of poor primers. If you have to cycle more than 35 or so times to get an amplification product, or if you have to use unusual additives or odd conditions to achieve success, your primer may not be efficient enough to use for sequencing.
If your primer is mismatched to your *original* template, after the PCR reaction, the product (which of course now incorporates your primers) will indeed match the primers perfectly. In other words, mismatch primers aren’t a problem *if* they’re the ones with which you amplified.
Please DON’T try to use a spectrophotometer to measure the concentration of a PCR product!
Typical laboratory spectrophotometers cannot with any accuracy measure the small amount of DNA that a PCR reaction generates. Unless you own one of the newer micro-specs (e.g. “Nanodrop” or similar), you should simply use an analytical agarose gel to estimate the concentration of your templates. Please compare them to a reference DNA fragment of similar size and known concentration!
PCR Reactions RARELY produce only single-bands.
You may think that your reaction produced just a single product, but there are very often other things there. When you use an agarose gel to assess the PCR result, you can’t detect any of these:
- Problem: Small, illegitimate products
Amplification at an illegitimate site that gives rise to a small (<100 nt) fragment will never show up on the typical 1% agarose gel, but it will sequence much better than your larger, ‘legitimate’ product.
Solution: Sequence it anyway. If you see interfering peaks for the first 20-100 nt (especially if they are extremely large), you should assume there’s interference from a small amplification product. If you get the sequence you needed despite the problem, great. If not, you can always cut the desired band out of a preparative gel, or go back and redesign your PCR reaction to avoid that interfering product. You may need to use a 2-3% agarose gel to see smaller fragments reliably.
Warning, however: those small products can produce such bright bands that they cause interference with someone else’s samples nearby! If we complain about your ‘overconcentrated’ samples, please consider whether the small fragments are causing problems, and please don’t keep sending such samples if they are!
- Problem: A diffuse, low-level background of illegitimate products
When you assess the outcome of a PCR reaction, look closely at your analytical gel for a dim background smear of ethidium bromide staining. If it’s there, it’s probably DNA, and (because it’s distributed and diffuse) it’s probably a LOT of DNA in many PCR bands. It could easily comprise the majority of DNA in your sample, and the sequencing result could be very bad.
Solution: well, you could sequence it anyway; sequencing is very cheap these days. If you do see multiple superimposed peaks instead ofthe clean sequence, you need to clean up your product some more. Cut the desired band out of a preparative gel, or go back and redesign your PCR reaction to avoid that interfering product.
- Problem: Your “single band” is really two superimposed products
It’s surprising, but this is not a particularly rare event. Everything looks great, but when you sequence it, all you see is multiple, superimposed sequences. This may occur if you are amplifying two related – but non-identical – genes, or if you have a homopolymer tract (e.g. poly-A or poly-T that causes the polymerase to ‘stutter’, or simply that you were unlucky.
Solution: If you have some way to PROVE that you amplified only the right product, it is always worth the effort to do so. Don’t just assume that the right size means the right band. For instance, if you happen to know of a restriction site within your expected PCR fragment, then cut the PCR product and verify that you get the expected fragment(s). If there are unexpected bands, unexpected sizes or bands that do not cut, be suspicious of the identity or purity of your product.
The use of nested PCR primers can minimize this problem. It is less likely that an illegitimate product will co-amplify through a second round of PCR with internally- nested primers.
Sounds a bit dismal, right? Not necessarily. There are some very good reasons you might want to go ahead and sequence directly from a PCR product. Here are some:
Direct sequencing is much quicker. If you’re screening hundreds of patient samples for mutations in a gene, you do NOT want to be gel-purifying all those PCR reactions, and you CERTAINLY don’t want to clone them all before sequencing.
Direct sequencing doesn’t show any PCR mutations. Common PCR protocols (Taq polymerase under standard cycling conditions) generate mis-incorporations occasionally (about once per 3 kb, in my hands). If you clone those PCR products and sequence several of them, you will see point mutations in some of the clones.
If you directly sequence the PCR product, though, what you’ll see is the consensus base at each position. Although many of the individual products have mutated nucleotides, these mutations are scattered randomly and are different for each individual product fragment. Consequently, at any one nucleotide, most of the clones will be correct, and you’ll be seeing the original sequence with no mutations.
We have many clients who have successfully sequenced thousands – even tens of thousands – of PCR products, with outstanding results. Be critical of the quality of your PCR product and, if necessary, optimize the PCR conditions or gel-purify the desired fragment. Prove you have the right fragment before you invest in large-scale sequencing, and you’ll be pleased with the results.