Hello
I am trying to do insertion (1 basepair change, 2 basepair changes) and deletion (50 base deletion) mutations using the Agilent Quikchange Lightning kit. However the mutagenesis does not seem to work. I have colonies from all of the mutations, but they are all false positives. Could someone please recommend any suggestions.
I constructed the primers using the agilent website (TM > than 78 degrees) and the primers were purified according to their protocol. I also checked for secondary structures such as hairpins or self-dimerization and couldn’t find anything that would contribute to a low mutagenesis efficiency. (see attachment for primers).
My plasmid is about 10kb , so I used the following program:
1 cycle - 95 degrees - 2 min
18 cycles } - 95 degress - 20 sec
- 60 degrees - 10 sec
- 68 degrees - 5 min (30 sec for 1KB)
1 cycle - 68 degrees - 5 min
Troubleshooting:
I have increased the annealing temperature to 68 degrees as recommended by the kit and i still don't have any colonies.
The recommended dpn1 digestion time with the kit is 5 min at 37 degrees. I have increased the dpn1 digest up to an hour. I have fewer colonies with increased dpn1 digest but again they were false positives.
I have tried using different concentrations of the template (10ng, 50ng and 100ng) and I still have colonies with no mutations.
The control plasmids (pWhitescript 4.5-kb control plasmid ) provided with the kit worked well (see attachment for picture). My template lacks the LacZ region and i cannot select colonies using x-Gal/IPTG. I selected almost all the colonies and sequenced them.
I would be really grateful for any help/advice.
Thank you
Cheers
Pranetha
Hi DpnI treatment seems to be the case. Try to add more and run a control reaction to make sure, all template has been cleaved (Ie, prepare the same reaction either without polymerase or without primers - DpnI digest and transform. You should get no colony at all if DpnI digestion is sufficient.). Do not be afraid of adding too much enzyme - in this case overkill is much better than incomplete cleavage.
Another possibility to explain if you are trying to do blunt end cloning at a nucleotide site which has blunt ends is that the primers are not 5' phosphorylated before you start PCR. In other words at least one strand has to be able to join in the ligation, without 5'P cannot.
Given that you have only have false positives your parental plasmid is definetely not digested enough by dpn1 as Martin Lenicek said.
This however might not be the only problem. How did you design your primers? Is the desired insert in the middle of your primer or at the 3' kissing end with a small overhang ?
Hello Michiel
I designed the primer using the agilent software.. It designs the mutation to be in the middle of the primer.
I have good experience with the Q5 site directed mutagensis kit .
https://www.neb.com/products/e0554-q5-site-directed-mutagenesis-kit
Maybe you can use the method of primer design in your kit, or maybe try the specified kit. Might also be easier with the included Kinase Ligase Dpn1 mix. Of course you can also make this mix yourself.
Hope this helps! Let me know
Although this method works in many cases, but it seems not working for you. The problem is more likely in the PCR amplification step in which your designed primers are prone to form primer-primer dimer but not for plasmid DNA amplification. If that is the case actually there is no or very limited newly synthesized plasmid DNA (can check in agarose gel) with your primers, all the transformable DNA actually are the remained parental DNA after DpnI digestion and the colonies are all the wild type. The quick way to get your work done is to redesign your primers based the linked article or the protocol and redo your SDM as described. The attached link is a modified protocol with some explanations and proved much more efficiency, https://bmcbiotechnol.biomedcentral.com/articles/10.1186/1472-6750-8-91, the attached lab protocol is summarized based on that publication and much easier to follow if you like.
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