How to make the mutation you want and succeed every time

Good research is highly reliant on good tools. Among the most important tools in molecular biology and biochemistry are mutant versions of the genes we study. These mutants help us understand how the proteins encoded by the genes function, both in vivo and in vitro. Today, I shall discuss the site-directed mutagenesis protocol I've been using to get my mutants. My protocol works. Every. Single. Time.

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.

Notice how the primer extents for 10 bases to both side of the mutation? Shorter primers also work, but try to extent it at least 9 bases on both sides of the mutation while keeping the GC content within 40-60% (in this case, it's 52.4%) and the melting temperature between 52-55C (53.5C, in this case). Bear in mind that the mutation you make will not count towards the melting temperature of the primer, as the mutation represents a mismatch! Of course, you can introduce more than one mutation, but the primer will have to be proportionally longer. I've successfully introduced up to seven point mutations with a single primer pair in one reaction using this method.

2. PCR

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.

6. Sequencing

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.

Conclusion

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!

Apoptosis vs. Necroptosis

Cell death is a highly regulated process. Ask any cancer or stroke patient. In the former case, too little cell death is causing problems, in the latter it's too much cell death that's doing the damage. Every day, approximately 100 billion (!) cells die in your body and every day, all those cells are replaced. Every day you die a little and you never even noticed.

Most of those cells die by a process called apoptosis; a programmed form of cell death. Every cell is genetically programmed to undergo apoptosis, a sequence of orchestrated events leading to the cells demise, when the cell has either suffered irreparable internal damage or receives an extrinsic stimulus from its environment. The extrinsic signal to die is given when a cell is, for example, infected with a virus, has become old or redundant or has become dislodged from its usual place in the body. The cell is basically told to quietly commit suicide, dismantle itself and allow its remains to be recycled. Cells are equipped with 'death receptors', members of the TNFa super family, that receive the death signal.

As mentioned above, apoptosis can also be triggered intrinsically. When, for example, a cell's genome or essential organelles have suffered irreparable damage a sequence of events leads to the release of toxic proteins from the mitochondria, such as cytochrome c and SMAC, that induce the cell to undergo apoptosis and remove itself from the population.

Extrinsic versus intrinsic apoptosis (image by author)

Apoptosis is a quiet, dignified form of cell death that does not trigger an inflammatory response. Apoptosis depends on the sequential activation of the caspases; a family of cysteine proteases. At the top of the chain are the initiator caspases (caspase-8 and -10 for death-receptor-induced apoptosis, caspase-9 for intrinsically-triggered apoptosis), at the bottom are the executioner caspases (caspase-3, -6 and -7) that dismantle the cell. Diametrically opposed to apoptosis is necrosis; a messy form of cell death -wherein the cell's contents are spilled into the environment- that does elicit an inflammatory response.

Surprisingly, necrosis can also follow a genetically-encoded program, similar to apoptosis. However, programmed necrosis, now widely known as 'necroptosis', does not depend in caspase activity, but on the activity of a kinase: Receptor Interacting Protein Kinase 1 (RIPK1). RIPK1 functions as the initiator of the pathway, while several downstream kinases (most notable RIPK3) serve as the executioners. We don't know much about necroptosis yet, but new findings on this form of cell death are published almost daily. What we do know for sure is that caspase activity is essential to prevent it. Necroptosis only occurs in the absence of caspase activity (for a free review by some friends of mine, see here).

The initiators of death-receptor-dependent apoptosis, caspase-8 and caspase-10, have both been shown to cleave and inactivate RIPK1 (reviewed by me here). Thus, if those caspases are activated, RIPK1 is inactivated. The opposite is also true: When the gene for caspase-8 is knocked out in mice (mice don't have the gene for caspase-10), the embryo deficient in caspase-8 dies on the eleventh day after gestation (Varfolomeev et al. 1998). This is a crucial day in the development of the murine embryo, since at that time the embryo's own blood circulation kicks in. In the absence of caspase-8 activity, the hearth and blood vessels of the embryonic mice fail to develop. Experiments with conditional knockout mice, mice that are only deficient for caspase-8 in certain tissues, have revealed that caspase-8 activity is also essential for the development of the immune system (see Kang et al.).

This failure of caspase-8 deficient embryos to develop, is entirely due to the activity of RIPK1 and its downstream effector RIPK3. Knock either one of these genes out concomitantly with caspase-8 or the adapter protein FADD (essential for the activation of caspase-8) and the mouse develops just fine (here, here, here and here; only the last one, the one I'm on, is free) . Or at least; it develops past this crucial stage, past day 11, for knockout of RIPK1 is lethal in itself. RIPK1 has pro-life as well as pro-death functions, but knockout of RIPK3 is relatively safe. Mice deficient in RIPK3 as well as caspase-8 develop into relatively healthy individuals. They still have some problems, of course, caspase-8 and RIPK3 are not entirely useless genes you can just dispose of. They have problems dealing with viral infections, for example, and their T cells proliferate unchecked. They may have other problems too, that haven't surfaced yet.

Thus, caspase-8 has a crucial pro-survival role in shutting off RIPK1 and preventing it from inducing necroptosis. But how, then, does a cell wherein caspase-8 is activated not die by apoptosis instead? How does it live to develop into a healthy mouse or human? Caspase-8 activates through dimerization; two molecules of caspase-8 are forcefully brought together to form an active complex. The previously mentioned adapter protein FADD is essential for initiating this process of dimerization, but recent evidence has shown that once a few dimers are formed around clusters of FADD, more caspase-8 dimers can form independent of FADD. An important clue comes from the observation that caspase-8 does not only activate when it dimerises with itself to form a homodimer, but can also when it forms a dimer with its cousin, FLIP (FLICE-like Inhibitory Protein), to form a heterodimer. FLIP is similar to caspase-8 but has no protease activity, it is an inactive caspase homologue. The heterodimer is active, but has a restricetd substrate repertoire; it cleaves the pro-apoptotic substrates of caspase-8 with very low efficiency, while it cleaves the non-apoptotic substrates of caspase-8 just as efficiently as the homodimer. I recently published a very readable review on the proliferative versus the apoptotic functions of caspase-8, find it here.

Apoptosis vs. Necroptosis vs. Survival  (image by author)

Now, this is not the end of it. There is much more to RIPK1 signaling than necroptosis; it plays an important role in immune activation and development too. In addition, recent evidence suggests that direct cleavage of RIPK1 cleavage by caspase-8 may not even be the key to prevention of necroptosis. Instead, caspase-8 may cleave CYLD, a de-ubiquitinating enzyme and an important regulator of RIPK1 activity. As long as CYLD is active, RIPK1 can promote necroptosis but if CYLD is inactivated RIPK1 is more likely to promote cell survival. However, caspase-8 is very bad at cleaving either CYLD or RIPK1. The paracaspase MALT1 can also cleave CYLD, an event that is crucial for the activation of T cells, but it is not so very good at it either. Could there be another caspase, downstream of caspase-8, that cleaves and inactivates RIPK1? Does cleavage of RIPK1 really lead to its inactivation or do the two fragments gain a different function? Does the caspase-8/FLIP heterodimer have other substrates, besides RIPK1 and CYLD? These and other important questions are currently under investigation. We're not even sure yet what the relevance of necroptosis is for either normal human physiology or pathology.

Surely these are exciting times for the fields of cell death and inflammation! I will use this blog to review the latest findings in these fields, both by myself and by others. I hope to attract opinionated readers both inside and outside the fields and get some discussions going.