Site directed mutagenesis - (Jul/30/2013 )
I have a big problem with my mutagenesis. I'm supposed to create 12 mutants, each of them one single point mutation. Unfortunateley it doesn't work and I'm very desperate now.
I designed my primers with the Quick change sdm programme. And all of them are above 80°C.And reverse and forward primers bind to the same sequence on opposite strands.
I have two plasmids one 8.5kb and the other one 7.9kb. So I make 6 mutations separatley in each of them. I followed the kit instructions (without having the kit, but having Pfu Turbo) and I could create 2 mutants in the 7.9kb plasmid and one in the 8.5kb very easily. After that eveything went down. I didn't get any colonies or I got some, which I couldn't recover in liquid media.
I begged my supervisor to buy the kit, but even with the kit it didn't work.
I started to optimise my PCR using normal Pfu and DMSO. And running the cycles 20 times (in the kit it says 12 times) to see if I can detect bands on agarose gel and yes I can (correct size). After DpnI (brand new enzyme) digest I transformed once into electrocompetent XL1blue and into chemical competent XL1blue cells. I got colonies and sequenced the area, but I didn't get any correct clones (they had a mutation, but on the wrong spot).
I thought I needed a more accurate polymerase and now I'm using the Phusion Hot start II high fidelity DNAP.
The protocol looks like that:
5x phusion buffer: 1x
10mM dNTPs: 200um each
primer A: 0,5uM
primer B: 0,5uM
Polymerase: 0,02 U/ul
4)72 4,25 min (30s/kb)
4 till 2 20x.
I digested with DpnI for 2 1/2 hrs and then purified with a PCR purification kit and measured with Nanodrop, so I still have 30ng/ul as concentration. I used 45ng for transformation. But I don't get any colonies.
I was thinking maybe I can increase the transformation efficiency if I ligate before I transform.
I have no idea what to do next.
It's difficult to really tell what you are doing with the mix of volumes and concentrations in your protocol. If you are providing a protocol, listing both would help.
I don't know what to make of the template: 20 ul line. This is probably vastly more template than you should be using. What is the concentration of the template?
I would guess also that you are using too much PCR product in the transformation. With chemical transformation, more than 5% volume of DNA added is inhibitory. What is the competence of your cells? Are these commercial cells, or are you making them yourself?
5x phusion buffer: 10µl (final conc 1x)
10mM dNTPs: 1 µl (200µM each)
10 µM primer A: 2,5µl (0,5µM )
primer B: 2,5µl (0,5µM)
DMSO : 1,5µl (3%)
2U/µl enzyme: 0,5µl (0,02U/µl)
20ng plasmid DNA: 1µl (20ng)
rest is water till 50µl
yeah I just saw in the recommendation that you should add template from 1pg-10ng in 50µl reaction volume, so i added too much.
For the transformation, I added the same amount of DNA (my control plasmid, which has no mutation) to my positive control and I have no problems to get colonies, but I will try to decrease the amount and see what happens.
I made the cells myself and they have a transformation efficiency of 2.1x10^8 cfu/µg.
I don't know how your primers are prepared (desalted, etc), but spend a bit of money to have them PAGE- or HPLC-purified.
*Most* mutagenesis reactions work with just desalted primers, but not all.
I was in the same situation as you with no successful mutagenesis reaction (using only desalted primers) for a month. A new set of primers with the same sequence that were PAGE-purified corrected the problem.
yeahhh I checked my primers a hundred times and they are HPLC purified. I'm really running out of options :(
Take an alternative approach. For example, design primers to amplify your vector at the mutation site. Use an offset restriction enzyme such as BsaI in the 5' end of construct. Design the mutation into the cut region (on both ends). Cut with BsaI, ligate, go.