Site-directed mutagenesis, also called site-specific mutagenesis or oligonucleotide-directed mutagenesis, is a molecular biology technique in which a mutation is created at a defined site in a DNA molecule. In general, this form of mutagenesis requires that the wild type gene sequence be known. It is commonly used in protein engineering.
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Early attempts at mutagenesis were non-site specific using radiation or chemical mutagens.[1] Analogs of nucleotides and other chemicals were later used to generate localized point mutations,[2] examples of such chemicals are aminopurine,[3] nitrosoguanidine,[4] and bisulfite.[5] Site-directed mutagenesis was achieved in 1973 in the laboratory of Charles Weissmann using N4-hydroxycytidine which induces transition of GC to AT.[6][7] These methods of mutagenesis however are limited by the kind of mutation they can achieve.
In 1971, Clyde Hutchison and Marshall Edgell showed that it is possible to produce mutants with small fragments of phage ϕX174 and restriction nucleases.[8][9] Hutchison later produced with his collaborator Michael Smith in 1978 a more flexible approach to site-directed mutagenesis by using oligonucleotides in a primer extension method with DNA polymerase.[10] For his part in the development of this process, Michael Smith later shared the Nobel Prize in Chemistry in October 1993 with Kary B. Mullis, who invented polymerase chain reaction.
The basic procedure requires the synthesis of a short DNA primer which is complementary to the template DNA around the site where the mutation is to be introduced. The mutation may be a single base change (a point mutation), deletion or insertion, containing the desired mutation such as a base change. This synthetic primer is complementary to the template DNA around the base change so it can hybridize with the DNA containing the gene of interest. The single-stranded primer is then extended using a DNA polymerase, which copies the rest of the gene. The gene thus copied contains the mutated site, and is then introduced into a host cell as a vector and cloned. Finally, mutants are selected.
The original method using single-primer extension was inefficient due to a lower yield of mutants. The resulting mixture may contain both the original unmutated template as well as the mutant strand, producing a mix population of mutant and non-mutant progenies. The mutants may also be counter-selected due to presence of mismatch repair system which favors the methylated template DNA. Many approaches have since been developed to improve the efficiency of mutagenesis.
A large number of methods are available to effect site-directed mutagenesis,[11] although most of them are now rarely used in laboratories since the early 2000s as newer techniques allow for simpler and easier way of introducing site-specific mutation into genes.
In 1987 Kunkel et al. introduced a technique which reduces the need to select for the mutants.[12] The vector DNA to be mutated is inserted into an E. coli strain deficient in two enzymes, dUTPase and uracil deglycosidase. The dUTPase deficiency prevents the breakdown of dUTP, a nucleotide that normally replaces dTTP in RNA, resulting in an abundance of dUTP; the uracil deglycosidase deficiency prevents the removal of uracil from newly-synthesized DNA. As the double-mutant E. coli replicates the vector DNA, its enzymatic machinery may therefore misincorporates dUTP instead of dTTP, resulting in DNA which contains some uracils. This copy is extracted, and then used as template for mutagenesis. An oligonucleotide containing the desired mutation is use for primer extension. The heteroduplex DNA formed may be chimeric containing one strand unmutated and containing UTP, and the other strand mutated and containing dTTP. The DNA is then first treated with uracil deglycosidase which removes the uracil in the template, then with alkali which degrades the template DNA as it has its uracil removed making it sensitive to alkali. The resulting DNA therefore consists of only the mutated strand.
Unlike other methods, cassette mutagenesis need not involve primer extension using DNA polymerase. In this method, a fragment of DNA is synthesized, and then inserted into a plasmid.[13] It involves the cleavage by a restriction enzyme at a site in the plasmid and subsequent ligation of a pair of complementary oligonucleotides containing the mutation in the gene of interest to the plasmid. Usually the restriction enzymes that cuts at the plasmid and the oligonucleotide are the same, permitting sticky ends of the plasmid and insert to ligate to one another. This method can generate mutants at close to 100% efficiency, but is limited by the availability of suitable restriction sites flanking the site that is to be mutated.
The limitation of restriction sites in cassette mutagenesis may be overcome using polymerase chain reaction with oligonucleotide "primers", such that a larger fragment may be generated covering two convenient restriction sites. The exponential amplification in PCR produces a fragment containing the desired mutation in sufficient quantity to be separate from the original, unmutated plasmid by gel electrophoresis, which may then be inserted in the original context using standard recombinant molecular biology techniques. There are many variations of the same technique. The simplest method places the mutation site towards one of the ends of the fragment whereby one of two oligonucleotides used for generating the fragment contains the mutation. This involves a single step of PCR, but still has the inherent problem of requiring a suitable restriction site near the mutation site unless a very long primer is used. Other variations therefore employ three or four oligionucleotides, two of which may be non-mutagenic oligonucleotides that cover two convenient restriction sites and generate a fragment that can be digested and ligated into a plasmid, while the mutagenic oligonucleotide may be complementary to a location within that fragment well away from any convenient restriction site. These methods require multiple steps of PCR so that the final fragment to be ligated can contain the desired mutation.
For plasmid manipulations, other site-directed mutagenesis techniques have been largely supplanted by techniques which are highly efficient but relatively simple, easy to use, and commercially available as a kit. An example of these techniques is the Quikchange method,[14] where a pair of complementary mutagenic primers are used to amplify the entire plasmid in a thermocycling reaction using a high-fidelity DNA polymerase such as pfu polymerase. The reaction generates a nicked, circular DNA which is relaxed. The template DNA must be eliminated by enzymatic digestion with a restriction enzyme such as DpnI which is specific for methylated DNA. All DNA produced from Escherichia coli would be methylated; the template plasmid which is biosynthesized in E. coli will therefore be digested, while the mutated plasmid is generated in vitro and is therefore unmethylated would be left undigested. Note that in these double-stranded plasmid mutagenesis methods, while the thermocycling reaction may be used, the DNA need not be exponentially amplified as in a PCR, instead the amplification is linear, and it is therefore inaccurate to describe them as a PCR reaction since there is no chain reaction.
Site-directed mutagenesis is used to generate mutations that may produce rationally designed protein that has improved or special properties.
Investigative tools - specific mutations in DNA allow the function and properties of a DNA sequence or a protein to be investigated in a rational approach.
Commercial applications - proteins may be engineered to produce proteins that are tailored for a specific application. For example, commonly-used laundry detergents may contain subtilisin whose wild-type form has a methionine that can be oxidized by bleach, inactivating the protein in the process. This methionine may be replaced by alanine, thereby making the protein active in the presence of bleach.
