I've been pushed to the edge of sanity with this PCR that I'm trying to do. I have to amplify the coding region (that means exons only, no introns and no UTRs) of a gene. In addition, I'm trying to add on two restriction cut sites to the ends as well. The gene itself isn't enormous (1.3 kb), but I've spent 4 weeks attempting to amplify and have ended up with very nice blank gels. One primer has a Tm of 72 (it is 27 bp long with a 59% GC content) while the other has a Tm of 65 (40 bp long, 33% GC). Since I only want to amplify exons only, I can't use alternative primers. I've never used primers longer than 40 bp and don't feel comfortable shortening the other below 27 bp. I've done qPCR to quantify expression levels of this gene in different cDNAs and the results are what is to be expected. I've selected the cDNAs that seem to have the highest expression levels for this gene.
My conditions are as follows: 95 degree melt, an annealing temp gradient between 55 - 65 (30 secs), and an extension temp of 72 (1:45 minutes). I'm using the Denville Taq and its respective Taq buffer (with Mg salts already mixed in).
Here's what I've tried in the last few weeks:
5% DMSO - I get a very very strong band at around 200 bp at all temperatures in my gradient. No other products. My positive control (beta-actin) shows up very strongly with no nonspecfic banding.
1 M betaine - No different from my normal PCR - very pretty ladders and nothing else. Positive control again shows up beautifully.
Pfx (high fidelity polymerase) - No different from my normal PCR. Weaker amplification of positive control.
Extension time of 2:30 - Random products show up, especially at my lower temperatures.
Any feedback or advice?
Edit: I get weak primer dimers in all my PCR results, almost without fail.
Edited by jamesmhyu, 28 October 2010 - 02:00 PM.