Need help with dCAPS pcr, seeing huge bands on gel - (Dec/02/2013 )
I am a research assistant at a university here in the US. I have been trying to optimize a certain dCAPS PCR and digest protocol for some time now and I am having a lot of trouble. I have submitted some pictures of my gels so you can see what I mean.
This first pic is the result I have been getting for some time. There are always huge fragments appearing on the gel (the last row of fragments that I have circled is in the right range for the mutation I'm looking for) and I can't figure out what the problem is.
The second picture (the one with two prints) is a nice result I got last week while I was tweaking our protocol yet again. The top picture is the PCR product and the bottom is after the restriction digest. Since the overall volume of the kit I am using is very small, I decided to double it so I could use a bigger piece of zebrafish tail fin. The result was much cleaner that the others I had been getting Since it worked pretty well I decided to repeat it using new tissue samples but I got the third picture as my result, having giant fragments appearing once again.
What I am using: Sigma Red Extract-N-Amp kit and NEB restriction enzyme DraI digesting for 2.5-3 hours (the buffer doesn't seem to make a lot of difference, I have tried both the NEB Cutsmart buffer and the Lifetech REact Buffer 1). I am running all products on 2% agarose gel.
I don't have a lot of experience with PCR (I have a bachelor's in Zoology and I am interested in evolution so I am trying to get more experience with microbio techniques) so I'm not really sure what to do. There is an associate in my lab who has been "helping" me, but she is very belligerent and difficult and she believes the kit doesn't work and keeps trying to make me use her homemade preps. Her homemade preps work for a different mutant we work with in the lab, but when I use them for this mutant I just get a big smear and no bands appear at all. Plus I'm just sick of her laughing at me when the gel fails and acting like I'm an idiot. I have confidence in the kit since our grad student uses it with no problems and she was the one who recommended doubling the kit volume. Could it be contamination or something else? This mutant is notoriously difficult to pin down but I don't think anyone has had these problems before. The fish are getting old and I need to freeze down some sperm from the mutants as soon as possible, so any advice you can give me would be great.
OK, I'm not familiar with the exact dCAPs procedure, but it seems to be a combination of PCR with a primer that incorporates a RE site followed by digestion to detect the mutants, which will cleave, while normals won't (or the other way around, it makes little difference for optimization).
The first thing I would test is the PCR - I'm not sure from your images whether these multi-bands are post PCR or post digestion, but assuming post-PCR. It would seem that your PCR is not reliable - you should run a temperature gradient for the annealing temperature to check if there is a more optimal temperature for the primers. You may also need to try some different polymerases, as these can have lower error rates (note: you probably want to avoid proofreading polymerases).
Would it be possible to use a different primer? Often non-specific bands are due to poor primer design, a few bases added or subtracted from the end of a primer can make all the difference sometimes.
Make sure that you aren't loading too much DNA, for genomic DNA you want to have less than 500 ng per reaction. Often less DNA will give you a cleaner result. Also make sure that you don't have more than 30 cycles, after this point you could be amplifying just about anything non-specifically.
For the digest - make sure that your RE is working and that you aren't overloading the enzyme, you typically want a 2 - 5 fold amount of enzyme:DNA (e.g. if you have 1 ug DNA, use 2 - 5 units of enzyme). Check the enzyme for star activity, as overdigestion can cause this, so you need to keep the digest short (1 h should be enough).