Some plasmid backbones are hard to mutagenise (eg. some pcDNA3 backbones due to high GC content). If the problem is lack of colonies, why not avoid the PCR cleanup step. That is unnecessary and may affect the transformation efficiency of large templates. Good Luck.

I agree with Manoj about the cleanup being redundant. 

In addition, you can frequently resolve high CG issues by adding DMSO to the PCR reaction - 3% final conc.  Also, you caun increase the number of cycles to get more product for the transformation. 

Best of luck. 

The main problem of Quikchange is its poor PCR amplification.Try re-design your primers as described http://bmcbiotechnol.biomedcentral.com/articles/10.1186/1472-6750-8-91 and read the explanations. You should get your work done easily just following that modified method. The attached file is a lab protocol I generated for our institute based on that publication, it is much easier to follow than the original paper.

Hi Charles,

I have been doing quite a lot of Quikchange Site Directed Mutagenesis.

I always use their software to generate the primers and never had any trouble. All my mutagenesis always worked almost right away.

As Manoj mentioned, you don't have to PCR purify after the DnpI digest. Directly transform 2uL of your DpnI-treated SDM PCR (50uL).

I have always used XL10-Gold cells, sold with the kit. 

If it still doesn't work, I would suggest you to increase your extension time. I had once difficulties with a particular plasmid. It ended up being that the plasmid was over 1000bp bigger than what my map said. I increased the extension time and it ended up working fine. As a consequence of a bigger plasmid, I had to test different amount of template for the PCR as well . Maybe you are also not putting enough?

Good luck !

Best,

Cécile

I have done many quickchange mutagenesis with variable results, depending mainly on the pair of mutagenic oligonucleotides. I normally use 50 as the annealing temperature for the first attempt. Playing with higher temperatures is sometimes useful, but in general in my experience the differences have not been significant.

Can you see the amplified products in an agarose gel? Even though colonies and mutated plasmids can be obtained even form reaction products that are not visible on a gel, if you can see a good amplification you will get more colonies and everything will be easier.

With poor amplification sometimes its really hard to get colonies. If tranformation fails, I go to electroporation. I agree that you dont need to purify reaction products before transformation, but if you go for electroporation  to obtain a much higher transformation efficiency you need to cean-up the products or to use a very small volume of non-purified reaction (1-3 microliters depending on the cuvette). In that way you can get colonies from reactions that did not produce them after thermal shock transformation.

If colonies are not obtained from electroporation or the clones are not mutated, sometimes its necessary to redesign the primers, extending the regions flanking the mutation. Its better if the ends of the mutagenic primers contain G/C.

Hi Charles,

Why don't you try different method for mutagenesis? Try the following method and use Q5 instead of Pfu.

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3672168/

I used this and it works well.good luck

hi Jahnavi

If you see the smear in the gel;

You need to change your reaction conditions

You can start by increasing the anneling temperature or reduce the primer concentration.

You also need to check the primer and redesign your primers as needed.....

good luck