Methods and Concepts in the Life Sciences/Enzymes for DNA Manipulation
Enzymes for DNA Manipulation
A restriction enzyme (or restriction endonuclease) is an enzyme that cuts DNA at or near a specific recognition nucleotide sequence known as restriction site. To cut DNA, all restriction enzymes make two incisions, once through each sugar-phosphate backbone of the DNA double helix.
These enzymes are found in bacteria and archaea and provide a defense mechanism against invading viruses. Inside a prokaryote, the restriction enzymes selectively cut up foreign DNA in a process called restriction; while host DNA is protected by a modification enzyme (a methylase) that modifies the prokaryotic DNA and blocks cleavage. Together, these two processes form the restriction modification system.
Over 3000 restriction enzymes have been studied in detail, and more than 600 of these are available commercially. These enzymes are routinely used for DNA modification in laboratories, and are a vital tool in molecular cloning.
Restriction enzymes are commonly classified into four types, which differ in their structure and whether they cut their DNA substrate at their recognition site, or if the recognition and cleavage sites are separate from one another.
- Type I enzymes cleave at sites remote from the recognition site and require both ATP and S-adenosyl-L-methionine (SAM) to function. They are multifunctional proteins with both restriction and methylase activities.
- Type II enzymes cleave within or at short specific distances from recognition site; most require magnesium. They are single function (restriction) enzymes independent of methylase.
- Type III enzymes cleave at sites a short distance from the recognition site. They require ATP (but do not hydrolyse it); S-adenosyl-L-methionine stimulates reaction but is not required. They exist as part of a complex with a modification methylase.
- Type IV enzymes target modified DNA, e.g. methylated, hydroxymethylated and glucosyl-hydroxymethylated DNA.
Type II restriction enzymes are the most common in molecular biology since they have a clearly defined cleavage site and do not require ATP. Their recognition sites are usually palindromic and 4–8 nucleotides in length. As can be seen from the image, they are homodimers.
Restriction enzymes are named after the bacterium from which they were isolated. The first letter refers to the bacterial genus, the second and third to the species and a forth letter may be added to identify the strain. Enzymes from the same bacterium can be distinguished by a Latin number which indicates the order of isolation.
Restriction enzymes which have the same recognition sequence and cut it in the same way are called isoschizomers. Neoschizomers are enzymes which recognize the same sequence but cleave it differently.
The following table gives an overview of typical protocols for analytical and preparative digests of plasmid DNA.
|Analytical digest||Preparative digest|
|plasmid DNA||0.1 – 0.5 µg||1 – 5 µg|
|restriction enzyme A||0.5 µl||1 – 2 µl|
|restriction enzyme B||0.5 µl||1 – 2 µl|
|10x buffer||10 % of total volume||10 % of total volume|
|water||fill up to total volume||fill up to total volume|
|total volume||10 µl||25 – 50 µl|
|incubation temperature||37 °C||37 °C|
|incubation time||10 min – 2 h||10 min – overnight|
If the aim is to create a vector for molecular cloning, it is important that the plasmid is digested completely, otherwise plasmids without an insert might be transformed and lead to false positive colonies. The digestion of the insert is less critical and can therefore be shorter.
Some suppliers offer optimized restriction enzymes, which are all active in the same buffer and only need very short incubation times. For a double digest with classical enzymes, it is necessary to check their compatibility.
The enzyme volume should not exceed 10 % of the total reaction volume to prevent inhibition or star activity due to excess glycerol (restriction enzymes are delivered in 50 % glycerol). Restriction enzymes are typically stored at -20 °C and should be kept on ice when they are not in the freezer. They should be the last component added to the reaction. Vortexing is not advised.
Some restriction enzymes do not recognize their restriction site if the DNA is methylated. This should be considered when working with an E. coli strain that expresses methylases.
Contaminants such as ethanol, EDTA, phenol, chloroform or detergents interfere with the activity of the enzymes, thus the DNA should be free of them.
- REBASE is the most comprehensive restriction enzyme database
- NEBcutter finds restriction sites in a DNA sequence
- The websites of the commercial suppliers NEB and Thermo Scientific contain extensive information about restriction enzymes
DNA ligase is an enzyme that facilitates the joining of DNA strands by catalyzing the formation of a phosphodiester bond. It plays a role in repairing single-strand breaks in duplex DNA in living organisms, but some forms (such as DNA ligase IV) may specifically repair double-strand breaks (i.e. a break in both complementary strands of DNA). Single-strand breaks are repaired by DNA ligase using the complementary strand of the double helix as a template, with DNA ligase creating the final phosphodiester bond to fully repair the DNA.
DNA ligase has applications in both DNA repair and DNA replication. In addition, DNA ligase has extensive use in molecular biology laboratories for recombinant DNA experiments.
The mechanism of DNA ligase is to form two covalent phosphodiester bonds between 3' hydroxyl ends of one nucleotide, ("acceptor") with the 5' phosphate end of another ("donor"). ATP is required for the ligase reaction, which proceeds in three steps:
- adenylation (addition of AMP) of a lysine residue in the active center of the enzyme, pyrophosphate is released;
- transfer of the AMP to the 5' phosphate of the so-called donor, formation of a pyrophosphate bond;
- formation of a phosphodiester bond between the 5' phosphate of the donor and the 3' hydroxyl of the acceptor.
Ligase will also work with blunt ends, although higher enzyme concentrations and different reaction conditions are required.
Types of DNA ligases
E. coli DNA ligase
The E. coli DNA ligase is encoded by the lig gene. DNA ligase in E. coli, as well as most prokaryotes, uses energy gained by cleaving nicotinamide adenine dinucleotide (NAD) to create the phosphodiester bond. It does not ligate blunt-ended DNA except under conditions of molecular crowding with polyethylene glycol, and cannot join RNA to DNA efficiently.
T4 DNA ligase
The DNA ligase from bacteriophage T4 is the ligase most-commonly used in laboratory research. It can ligate cohesive or "sticky" ends of DNA, oligonucleotides, as well as RNA and RNA-DNA hybrids, but not single-stranded nucleic acids. It can also ligate blunt-ended DNA with much greater efficiency than E. coli DNA ligase. Unlike E. coli DNA ligase, T4 DNA ligase cannot utilize NAD and it has an absolute requirement for ATP as a cofactor. Some engineering has been done to improve the in vivo activity of T4 DNA ligase; one successful approach, for example, tested T4 DNA ligase fused to several alternative DNA binding proteins and found that the constructs with either p50 or NF-kB as fusion partners were over 160% more active in blunt-end ligations for cloning purposes than wild type T4 DNA ligase.
A phosphatase is an enzyme that removes a phosphate group from its substrate by hydrolysing phosphoric acid monoesters into a phosphate ion and a molecule with a free hydroxyl group (dephosphorylation). This action is directly opposite to that of phosphorylases and kinases, which attach phosphate groups to their substrates by using energetic molecules like ATP. A common phosphatase in many organisms is alkaline phosphatase. Another large group of proteins present in archaea, bacteria, and eukaryote exhibits deoxyribonucleotide and ribonucleotide phosphatase or pyrophosphatase activities that catalyse the decomposition of dNTP/NTP into dNDP/NDP and a free phosphate ion or dNMP/NMP and a free pyrophosphate ion.
Cysteine-dependent phosphatases (CDPs) catalyse the hydrolysis of a phosphoester bond via a phospho-cysteine intermediate.
The free cysteine nucleophile forms a bond with the phosphorus atom of the phosphate moiety, and the P-O bond linking the phosphate group to the tyrosine is protonated, either by a suitably positioned acidic amino acid residue (Asp in the diagram below) or a water molecule. The phospho-cysteine intermediate is then hydrolysed by another water molecule, thus regenerating the active site for another dephosphorylation reaction.
Metallo-phosphatases (e.g. PP2C) co-ordinate 2 catalytically essential metal ions within their active site. There is currently some confusion of the identity of these metal ions, as successive attempts to identify them yield different answers. There is currently evidence that these metals could be Magnesium, Manganese, Iron, Zinc, or any combination thereof. It is thought that a hydroxyl ion bridging the two metal ions takes part in nucleophilic attack on the phosphorus ion.
Alkaline phosphatase (EC 220.127.116.11) can remove phosphate groups from many types of molecules, including nucleotides, proteins, and alkaloids. As the name suggests, alkaline phosphatases are most effective in an alkaline environment.
Typical use in the lab for alkaline phosphatases includes removing phosphate monoester to prevent self-ligation.
Common alkaline phosphatases used in research includes:
- Shrimp alkaline phosphatase (SAP), from a species of Arctic shrimp (Pandalus borealis). This phosphatase is easily inactivated by heat, a useful feature in some applications.
- Calf-intestinal alkaline phosphatase (CIP)
- Placental alkaline phosphatase (PLAP) and its C terminally truncated version that lacks the last 24 amino acids (constituting the domain that targets for GPI membrane anchoring) - the secreted alkaline phosphatase (SEAP)
Alkaline phosphatase has become a useful tool in molecular biology laboratories, since DNA normally possesses phosphate groups on the 5' end. Removing these phosphates prevents the DNA from ligating (the 5' end attaching to the 3' end), thereby keeping DNA molecules linear until the next step of the process for which they are being prepared; also, removal of the phosphate groups allows radiolabeling (replacement by radioactive phosphate groups) in order to measure the presence of the labeled DNA through further steps in the process or experiment. For these purposes, the alkaline phosphatase from shrimp is the most useful, as it is the easiest to inactivate once it has done its job.
Another important use of alkaline phosphatase is as a label for enzyme immunoassays.
Undifferentiated pluripotent stem cells have elevated levels of alkaline phosphatase on their cell membrane, therefore alkaline phosphatase staining is used to detect these cells and to test pluripotency (i.e., embryonic stem cells or embryonal carcinoma cells).
Exonuclease III (ExoIII) catalyzes the stepwise removal of mononucleotides from 3´-hydroxyl termini of duplex DNA. A limited number of nucleotides are removed during each binding event, resulting in coordinated progressive deletions within the population of DNA molecules. The preferred substrates are blunt or recessed 3´-termini, although ExoIII also acts at nicks in duplex DNA to produce single-strand gaps. The enzyme is not active on single-stranded DNA, and thus 3´-protruding termini are resistant to cleavage. The degree of resistance depends on the length of the extension, with extensions 4 bases or longer being essentially resistant to cleavage. This property is used to produce unidirectional deletions from a linear molecule with one resistant (3´-overhang) and one susceptible (blunt or 5´-overhang) terminus.
Nuclease S1 is an endonuclease derived from Aspergillus oryzae that splits single-stranded DNA and RNA into oligo- or mononucleotides. Nuclease S1 is used to remove single stranded tails from DNA molecules to create blunt ends, for opening hairpin loops generated during synthesis of double stranded cDNA and as a reagent in nuclease protection assays.
Taq polymerase is a thermostable DNA polymerase which was isolated from the thermophilic bacterium Thermus aquaticus in 1965.
T. aquaticus is a bacterium that lives in hot springs and hydrothermal vents, and Taq polymerase was identified as an enzyme able to withstand the protein-denaturing conditions (high temperature) required during PCR. Therefore it replaced the DNA polymerase from E. coli originally used in PCR. Taq's optimum temperature for activity is 75–80 °C, with a half-life of greater than 2 hours at 92.5 °C, 40 minutes at 95 °C and 9 minutes at 97.5 °C, and can replicate a 1000 base pair strand of DNA in less than 10 seconds at 72 °C.
Taq polymerase lacks the of 3' to 5' exonuclease proofreading activity, resulting in relatively low replication fidelity. Originally its error rate was measured at about 1 in 9,000 nucleotides. Taq has a terminal transferase activity, which means that it adds adenine overhangs to 3' ends.
Pfu DNA polymerase was isolated from the hyperthermophilic archaeon Pyrococcus furiosus. It has superior thermostability and proofreading properties compared to Taq DNA polymerase. Unlike Taq DNA polymerase, Pfu DNA polymerase possesses 3' to 5' exonuclease proofreading activity, meaning that it works its way along the DNA from the 5' end to the 3' end and corrects nucleotide-misincorporation errors. This means that Pfu DNA polymerase-generated PCR fragments will have fewer errors than Taq-generated PCR inserts. However, Pfu is slower and typically requires 1–2 minutes per cycle to amplify 1 kb of DNA at 72 °C. Using Pfu DNA polymerase in PCR reactions results in blunt-ended PCR products.