I have been trying to incorporate two point mutations in a 8kb plasmid using the Quick Change II site directed mutagenesis from Stratagene. The two mutated sites are 8bp apart. Also, I used the Stratagene primer designing software to design the primers. So far I have been unsuccessful getting a band after PCR. Couple of times after DpnI digestion, I did bacterial transformations (even though there was no band in gel), however, did not see any colony. Moreover, using the same PCR set up, I see blue colonies on X-GAL, IPTG coated plates for the control plasmid that comes along with the kit. I pretty much follow the kit guideline to set up the PCR reaction mix. Also, I tried with different amt. of template (5ng-50ng) keeping the primer concentrations constant, but no luck yet! Bottom is the PCR set up I used. Any idea will be highly appreciated.
Initial denaturation: 95C for 3min
18 cycles of 95C for 50sec.
55C for 50sec.
68C for 18min.
Final extension: 68C for 8 min.
Hold:4C.
I always run a gel taking 10ul. of the reaction mix after PCR and use the rest for DpnI digestion.
@Arjan: There is no issue with my control as the colonies are supposed to be blue due to the mutation. Here the stop codon TAA is mutated to CAA which codes for glutamine and the resultant colonies appear blue in an XGAL-IPTG plate by virtue of beta-galatosidase production. Also, I have been using 125ng. of each primer.
So far I have tried with a maximum of 50 ng of DNA. So, I may try increasing the template concentration and also by using thin walled tubes. Thanks
You could also try to increase the amount of PCR mix used for transformation (5-10 ul) and use different types of competents (in our hands TOP10 supercompetent cells work best). I would not worry too much about the band in the gel I had successful transformations several times without seeing any gel band. Also be careful not to mix the antibiotics in the plate (vs resistance gene of your vector).
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