When it comes to site-directed mutagenesis many people prefer the easy way out and use a kit. The QuickChange kit from Stratagene, for example, is a popular choice. However, this kit is rather expensive, unnecessarily complex and really not as good as advertised. The whole procedure can be performed with materials you probably already have in your lab at a fraction of the cost, if you follow the exact steps outlined below.
1. Primer Design
The first step towards generating a successful mutation is designing your primers. The same rules of thumb apply when designing a mutagenesis primer as for designing a cloning primer: you're looking for a primer that extents at least 9 bases on both sites of the mutation, with a GC content of 40-60% and a melting temperature of about 52-55C. I always use complementary primers, a pair of primers that are each others exact opposites. For example, in figure 1, I show you the primer I designed to mutate an arginine residue, R54, to a cysteine in human Ubiquitin B. I did this by changing a single 'C' in the primer to a 'T', as indicated by the yellow bar. The codon 'CGC' encodes an arginine, whereas ' TGC' encodes a cysteine. Easy as that!
|Figure 1. A primer designed to mutate arginine 54 to a cysteine (R54C) in human Ubiquitin B.|
Once your primer order has come in and you have your template ready, it's time to run a PCR reaction. For site-directed mutagenesis reactions, it's wise to use a reliable polymerase with proof-reading. Several options are available, but my favorite enzyme is Phusion from New England Biolabs. This enzyme is twice as fast as a regular DNA polymerase and a whopping four times as fast as other proof-reading enzymes, such as Pfu. It has, in my experience, also proven to be highly reliable. Using the Phusion enzyme and buffer, set up the following reaction:
200 ng DNA template
125 ng forward primer
125 ng reverse primer
2 µL dNTP (2.5 mM each)
10 µL 5X Phusion buffer
1 µL Phusion
x µL water (to 50 µL total)
The ideal amount of template to be used depends on a lot of factors, but I find 50-200 ng to be sufficient. If you're worried about background you can use a little less, but more generally works better.
Set up one reaction with and one without polymerase as a control! This is important, so make sure not to forget this control.
The following PCR program can be used for Phusion (Figure 2), adjust to primer melting temperature (2 degrees below the melting temperature of your primers is ideal) and plasmid size (30 seconds/kb, remember you're amplifying the entire plasmid plus your insert!). 18 cycles is generally sufficient.
|Figure 2. General PCR program used for site-directed mutagenesis with Phusion.|
It's best to run the PCR in the morning. The program only takes about 2 hours in total, when using Phusion; enough time for a hearty lunch and a cup of coffee!
3. Analyze Product on Gel
After the PCR is done, take 10 µL of your product and analyze it on a 1% agarose gel. This is a very important step in the mutagenesis process! If the mutagenesis reaction has worked you'll see a signifcant increase in the amount of DNA in your reaction with polymerase, as compared to your control reaction without polymerase. If you don't see this increase, the reaction hasn't worked and there's no point in proceeding! Have a good look at your primer design or try again with more template DNA. If the reaction has worked, you should see something similar to Figure 3.
|Figure 3. PCR product of the mutagenesis reaction analyzed on gel. Samples 1-3 were run without polymerase, samples 4-6 were run with Phusion polymerase added to the reaction.|
4. Digest Product with DpnI
If, and only if, you got significant amplification of your template after PCR, you can digest your product with the restriction enzyme DpnI. This enzyme only digests methylated DNA, so your template (which was amplified in and purified from E. coli) gets digested, whereas your mutation-bearing product, formed in the PCR reaction, remains intact. Simply add 1µL DpnI (I get mine from Promega these days, but any DpnI is fine) directly to your PCR tubes and put them at 37C for 2-3 hours. Don't forget them and leave them overnight, as there will be nothing left (I know from experience...).
5. Transform into DH5a and Mini-prep
After digestion, simply take 5-10 µL of your product and transform into competent DH5a E. coli to amplify the DNA. Any strain of competent cells will work, really. Plate the cells in LB agar plates after the transformation and let them grow overnight at 37C. The plate of the no polymerase control should have no colonies, whereas the other plate should have plenty.
I know from experience that if you digest a product that did not look like it was amplified on gel you will still get no colonies on the no polymerase control plate and some on the other plate. However, none of these colonies will harbor the mutation. I don't know why this is, yet it happens.
The following day, pick some 12 colonies from your plate and grow mini-prep cultures overnight. You'll need enough DNA for sequencing (50 ng/µL) and I usually grow and prep 2 mL culture for high copy plasmids and 4 mL culture for low copy plasmids. I like to use 2xTY medium for my cultures, rather than LB, as the yield is higher.
For your mini-preps you can use any kit you like, but be aware that the popular Qiagen kits are expensive but absolutely no better than any other kit out there. I generally like the kits from Machery-Nagel or, if you really want to safe some money, you can buy the Econo-Prep columns from Epoch Life Sciences and make your own buffers. Mini-prep columns can be recycled and re-used several times, so there's really no use in spending a lot of money here.
After you've done your mini-preps, it's time to send out your DNA for sequencing. Most labs these days use commercial sequencing services, and prices have dropped significantly in recent years, so I don't find it necessary to safe money in this step at the risk of losing time. In general, the mutagenesis reaction is rather efficient, but I like to sequence about twelve clones of each reaction to be on the safe side. I find that at least one in four clones generally contains the desired mutation.
If you don't have access to fast and cheap sequencing facilities, you can also consider designing your mutation so that you introduce a restriction site. That way, you can analyze your clones by restriction digest. However, nowadays, this is hardly cheaper, faster or more reliable than sequencing and you'll have to sequence anyway in the end.
Well, that's it! With this protocol, my mutagenesis reactions always work, provided I designed my primers right and used enough template. If you're introducing multiple mutations with very long primers, you might consider adding 1% DMSO to your PCR reaction. This prevents the formation of DNA super structures during amplification, but also decreases the efficiency of the polymerase. Good luck making some mutants of your own!