Let's say I have a DNA sequence with the following structure:
$$ 5' - N_n - S_1 - N_{1000} - S_2 - N_{1000} - S_3 - N_n - 3' $$
Here, the $N$s represent stretches of arbitrary sequence of the indicated length. $S_1$, $S_2$ and $S_3$ are all unique sequences 20 bp in length.
I design a forward primer PF
that anneals to the positive strand at $S_1$. I design two reverse primers PR1
and PR2
that anneal to the negative strand at $S_2$ and $S_3$ respectively. Their annealing temperatures are reasonably close.
Clearly, PF
and PR1
together will produce a 1040 bp PCR product, and PF
with PR2
will produce a 2060 bp product.
Now let's say I add 0.2 μM PF
(standard concentration), 0.1 μM PR1
and 0.1 μM PR2
all in the same reaction. I then run a PCR, with extension time sufficient for the 2 kb product.
I can see the following possible outcomes:
- I get a 50/50 mix of 1 kb and 2 kb products.
PR1
competes aggressively and I get hardly any 2 kb product and a lot of 1 kb product.- After a few cycles, the 1 kb product that has accumulated serves as an efficient forward primer for
PR2
, and I end up getting mostly 2 kb product and little 1 kb product.
Which one will happen? If it depends, what are the major factors on which it depends? Primer concentrations, extension time, annealing temperature, dNTP amount, number of cycles?
This question is basically a very simplified multiplex PCR. While much has been written about practical applications of multiplex PCR, I am hoping that answers will help me understand the considerations that go into deducing optimal multiplex PCR conditions without doing any empirical calibration. This isn't to circumvent the calibration, but to understand the process conceptually.