Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

Structure and function of nucleases in DNA repair: shape, grip and blade of the DNA scissors

Abstract

DNA nucleases catalyze the cleavage of phosphodiester bonds. These enzymes play crucial roles in various DNA repair processes, which involve DNA replication, base excision repair, nucleotide excision repair, mismatch repair, and double strand break repair. In recent years, new nucleases involved in various DNA repair processes have been reported, including the Mus81 : Mms4 (Eme1) complex, which functions during the meiotic phase and the Artemis : DNA-PK complex, which processes a V(D)J recombination intermediate. Defects of these nucleases cause genetic instability or severe immunodeficiency. Thus, structural biology on various nuclease actions is essential for the elucidation of the molecular mechanism of complex DNA repair machinery. Three-dimensional structural information of nucleases is also rapidly accumulating, thus providing important insights into the molecular architectures, as well as the DNA recognition and cleavage mechanisms. This review focuses on the three-dimensional structure-function relationships of nucleases crucial for DNA repair processes.

Introduction

Quality control of genetic material is a function conserved in all living organisms. DNA suffers from many environmental stresses, including attacks by reactive oxygen species, radiation, UV light, and carcinogens, which modify the DNA. In addition, there are intrinsic errors and unusual structures, which are formed during replication or recombination, and they must be corrected by the various repair protein machineries to avoid alterations of the base sequences or entanglement of the DNA. These DNA repair proteins may function independently, but in many cases, they form complexes to perform more efficient repair reactions. In the repair complexes, nucleases play important roles in eliminating the damaged or mismatched nucleotides. They also recognize the replication or recombination intermediates to facilitate the following reaction steps through the cleavage of DNA strands (Table 1).

Table 1 Nucleases involved in DNA repair

Nucleases can be regarded as molecular scissors, which cleave phosphodiester bonds between the sugars and the phosphate moieties of DNA. They contain conserved minimal motifs, which usually consist of acidic and basic residues forming the active site. These active site residues coordinate catalytically essential divalent cations, such as magnesium, calcium, manganese or zinc, as a cofactor. However, the requirements for actual cleavage, such as the types and the numbers of metals, are very complicated, but are not common among the nucleases. It appears that the major role of the metals is to stabilize intermediates, thereby facilitating the phosphoryl transfer reactions. Cleavage reactions occur either at the end or within DNA, and thus DNA nucleases are categorized as exonucleases and endonucleases, respectively (Figure 1). Exonucleases can be further classified as 5′ end processing or 3′ end processing enzymes, according to their polarity of consecutive cleavage.

Figure 1
figure1

Schematic diagram of the nuclease activity. The two strands of DNA are schematically drawn. The cleavage made by the nuclease is represented by arrowhead

This review describes the three-dimensional (3D) structural views of the actions of various nucleases involved in many DNA repair pathways. The rapidly accumulating genomic, biochemical and structural data have allowed us to classify various nucleases into folding families. In general, the nucleases involved in DNA repair recognize the damaged moiety through the remarkably large deformation of DNA duplexes, and thus in terms of their DNA recognition mode, they apparently differ from the sequence-specific endonucleases, such as the restriction enzymes. The active sites of DNA repair nucleases have some similarity with other nucleases, including the metal-coordinating residues; however, they also display pronounced diversity.

Nucleases in various categories of DNA repair

Replication

DNA polymerase replicates a new strand of DNA, the sequence of which is complementary to the template DNA. Most DNA polymerases in prokaryotes and eukaryotes are composed of two different enzymes, a polymerase and an exonuclease, encoded within the same polypeptide, but sometimes they are formed by different subunits. The exonuclease degrades misincorporated DNA strand in the 3′ to 5′ direction (Figure 2) (reviewed in Shevelev and Hubscher, 2002). Deletion of these proofreading nucleases results in lethal or strong mutator phenotypes in bacteria (Fijalkowska and Schaaper, 1996) and in yeast (Morrison et al., 1993), and causes cancer in mice (Goldsby et al., 2001).

Figure 2
figure2

Nuclease associated DNA repair pathways. The substrate DNAs are drawn schematically and the arrowheads denote nuclease cleavage. RNA regions are drawn in bold line

The removal of Okazaki fragments is another important process in replication. This DNA : RNA hybrid is required to initialize DNA polymerization, but once the replication starts, it is rapidly degraded. Most of the Okazaki fragments are eliminated by RNaseH, enzyme ubiquitously present in all living organisms. RNaseH produces nicks in the RNA region of Okazaki fragments (Figure 2). In eukaryotes and in archaea, FEN1 endonucleases also participate in the removal of Okazaki fragments (reviewed in Lieber, 1997). FEN1 is a multi-functional enzyme. In addition to the 5′ to 3′ exonuclease activity to remove the Okazaki fragments, the enzyme can also generate an incision at the junction point of a 5′ flap DNA structure. This latter activity is required to eliminate non-homologous tails in base excision repair and in recombination intermediates.

The replication process is stalled by various modes of DNA damage. Upon the halt of fork progression, the DNA polymerase and other protein complexes abandon the replication fork. The remaining fork must be processed by various fork-specific protein machineries. The most notable protein among them is Mus81, which was recently found as a new fork/junction specific endonuclease (Boddy et al., 2000, 2001; Interthal and Heyer, 2000; Kaliraman et al., 2001; Mullen et al., 2001). Genetic and biochemical analyses have revealed that this endonuclease is completely conserved in eukaryotes, while its homolog has been found in archaea. The loss of Mus81 in yeast causes UV or methylation damage sensitivity (Interthal and Heyer, 2000) and defects in sporulation (Mullen et al., 2001).

Base excision repair

Abasic sites within DNA duplexes are frequently produced by the actions of various DNA glycosylases involved in the base excision repair pathway, in addition to the spontaneous hydrolysis of bases. These apyrimidine or apurine (AP) sites are removed by AP endonucleases which cleave the phosphdiester bond next to an abasic site (Figure 2) (reviewed in Mol et al., 2000a). E. coli cells contain two AP endonucleases: endonuclease IV (endoIV) and exonuclease III (exoIII). Interestingly, these two enzymes show no sequence similarity to each other; although their AP endonuclease activities are quite similar. In eukaryotes, there seems to be a single, major AP endonuclease working in each organism. APN1, the yeast homolog of E. coli endoIV, shows sequence and catalytic activity similarity to endoIV. The absence of APN1 results in enhanced sensitivity to oxidative damage and alkylating agents (Ramotar et al., 1991). Mammalian organisms, including humans, bear Ape1, which shares sequence similarity with E. coli exoIII but lacks the intrinsic 3′ to 5′ exonuclease activity. In addition to the AP endonuclease activity, Ape1 also plays a major role in sensing the redox state of the cell (Xanthoudakis et al., 1992). The loss of Ape1 generates embryonic lethality in mice (Wilson and Thompson, 1997).

Mismatch repair

In prokaryotes, mismatch repair is conducted mainly by the MutSLH proteins, while the Vsr protein is responsible for mismatches in certain sequences (reviewed in Modrich and Lahue, 1996; Yang, 2000; Tsutakawa and Morikawa, 2001). In the MutSLH system, the MutS protein recognizes and binds mismatched base moieties of DNA. MutL mediates the interaction between the MutS and MutH proteins. MutH recognizes a hemimethylated GATC sequence, and cleaves next to the G of the non-methylated strand (Figure 2). The cleavage activity of MutH is enhanced by the MutL protein, although its mechanism remains unclear. Vsr is a mismatch-specific endonuclease involved in very short patch repair, and recognizes a TG mismatch at the specific sequence CT(A/T)GG, where the mismatch occurs at the second thymine, upon spontaneous deamination. Vsr makes an incision next to the mismatched base. In both cases, after the nick has been introduced, these sites are degraded by the RecJ, ExoVII, or ExoI nuclease and are resynthesized by the DNA polymerase.

Nucleotide excision repair

Nucleotide excision repair (NER) is primarily used to process DNA damage that is not repaired by base excision repair. These forms of damage involve those generated by the UV radiation and the large adducts produced by various chemicals. In the NER pathway, a short stretch of DNA containing the damaged nucleotide is removed. During this process, two incisions, on the 5′ side and the 3′ side are made by two different nuclease reactions (Figure 2) (reviewed in Petit and Sancar, 1999; Prakash and Prakash, 2000; de Boer and Hoeijmakers, 2000). In bacteria, this dual incision is performed by the UvrB-UvrC complex. In budding yeast, Rad2 and the Rad1-Rad10 complex make the 5′ and 3′ incisions, respectively. The same process in mammalian cells is conducted by their homologs, XPG and XPF-ERCC1, respectively. Deletions or mutations introduced into these nucleases cause sensitivity to UV damage, and result in cancer formation. In addition, abnormalities of these proteins cause defects in neural development.

Double strand break repair

Double strand breaks are generated by the accidental halt of fork progression during replication or by ionizing radiation and strand incision chemicals. They are also generated as an intermediate state during meiosis and V(D)J recombination. These double strand breaks are repaired through the two main pathways of non-homologous end joining and homologous recombination. In either case, the ends of the double strand breaks must be processed to initiate the repair reaction (Figure 2). Mre11 is a multi-functional nuclease involved in the processing of the DNA ends or hairpin structures (reviewed in D'Amours and Jackson, 2002). While Mre11 itself exhibits a ssDNA exonuclease activity, its complex with Rad50 processes double strand break ends. Moreover, in the presence of ATP, Rad50 activates the cleavage activity of Mre11. Mutations introduced into Mre11 cause an ataxia-telangiectasia-like disorder (Stewart et al., 1999).

V(D)J recombination involves a reaction process, in which hairpin DNAs are opened, and subsequently, both ends are connected. Recently, the Artemis : DNA-PK complex was shown to participate in this opening reaction (Ma et al., 2002). Although Artemis alone possesses a ssDNA exonuclease activity, its complex formation with DNA-PK allows the processing of the double strand break ends to open the hairpin structure. Defects in each protein cause severe immunodeficiency (Blunt et al., 1995; Kirchgessner et al., 1995; Moshous et al., 2001).

In homologous recombination, two homologous DNA strands are paired and are connected by D-loop structures or Holliday junction intermediates. In bacteria, the RuvC protein cleaves the Holliday junction at two symmetrical sites near the junction center to resolve the junction into two dsDNAs (Figure 2). Similar junction resolving enzymes have also been found in other bacteria, bacteriophages, and archaea (reviewed in Sharples, 2001). In eukaryotes, FEN1, XPF-ERCC1, and Mus81 are known to cleave the D-loop structure, while Cce1/Ydc2 processes Holliday junctions in mitochondria.

Structural classification of DNA repair nucleases

The primary sequences of nucleases are often poorly conserved, except for the motifs related to catalytic sites. The functional and biochemical properties of many nucleases have been studied extensively. However, in some cases, it is very difficult to identify the actual functional targets of the nucleases, because of their broad substrate specificity. Nevertheless, many candidates for nucleases are available from various genome sequences, and their functional properties can be inferred by sequence comparisons with other well-studied nucleases. For instance, Koonin and his associates have successfully classified nucleases, phosphoesterases, and phosphatases into several families, based on extensive data base analyses of the primary sequences (Aravind and Koonin, 1998a,b; Aravind et al., 1999). This classification has also revealed the relationships between nucleases and identified several new nuclease families.

In addition to the classifications of primary sequences, 3D structural data have been rapidly accumulating with respect to the proteins involved in DNA repair, including nucleases. Most of their structures were solved in the DNA-free states, although a number of them were determined in complex with cofactors or/and DNA (Table 2). The classification of nucleases in terms of their 3D structures provides more defined properties, since it is accepted that the 3D structures are much less diverged and more closely related to the functions than the primary sequences. As a matter of fact, in the type II restriction endonucleases, all of the structures share the common core motif, which includes the active sites, and thus could be grouped into a single folding family, despite its primary sequence diversity (reviewed in Pingoud and Jeltsch, 2001). In the following section, we describe each of the folding families of the DNA repair nucleases, classifications based on the SCOP database (Figure 3 and Table 3) (Murzin et al., 1995).

Table 2 Structural analysis of DNA repair proteins
Figure 3
figure3

Folding patterns of DNA repair nucleases. The core folding is drawn schematically. The yellow arrows indicate the core β-sheet, where the strand orders are numbered on the top. α-helices are shown as blue cylinders. The positions of the bound metals are marked by black circles. Representative repair nuclease of the folding is written in parenthesis

Table 3 Structural classification of DNA repair nucleases

RNaseH-like fold

The RNaseH-like fold, which is one of the most ubiquitous architectures in the protein world, has been found in RuvC, RNaseH, integrase, transposase, and proofreading exonucleases (Figure 3a). The core structure contains a five-stranded β-sheet flanked by several α-helices. The strand order is 32145, with strand 2 anti-parallel to the others. The active site residues, which are constituted according to the DDE motif, are located on one side of the sheet. These three (or sometimes four) acidic residues coordinate the metals, which are essential for the catalytic reaction. For instance, the crystal structures of RNaseHI exhibit one (Katayanagi et al., 1993) or two (Goedken and Marqusee, 2001) metals bound to the active site. Similarly, the active site of the proofreading subunit of DNA polymerase III coordinates two metals (Hamdan et al., 2002). The cocrystal structure with TMP revealed that the phosphate moiety is directly coordinated between the two metals, as it mimics the product DNA. A similar structure is also observed in the DNA complexes of the Klenow fragment of polymerase I (Beese and Steitz, 1991) and the RB69 DNA polymerase (Shamoo and Steitz, 1999).

Resolvase-like fold

This fold has been found in γδ resolvase, 5′–3′ exonucleases, and FEN1 (Figure 3b). It is similar to the RNaseH-like fold, with a five-stranded β-sheet. However, it possesses a different strand order, which is defined as 21345 with strand 5 anti-parallel to the others. FEN1 possesses two acidic clusters formed by four or three conserved aspartate/glutamate residues. These clusters each coordinate a metal, and are separated by 5 Å from each other (Hwang et al., 1998; Hosfield et al., 1998).

Restriction endonuclease-like fold

The structures of restriction endonucleases revealed that their catalytic domains share common fold architecture (Figure 3c). The core fold comprises a five-stranded β-sheet flanked by several α-helices. The strand order is 12345, with strand 2, and in some cases, strand 5, anti-parallel to the others. A conserved PDXn(D/E)XK sequence is located on one side of the β-sheet, and is involved in the formation of the catalytic centers in most restriction endonucleases. Similar sequences are also found in several DNA repair nucleases, such as MutH, Hjc, and T7 endoI, which are categorized into essentially the same folding family. The Vsr endonuclease also shares a similar fold, whereas the (D/E)XK sequence is replaced by FXH, where histidine participates in catalysis (Tsutakawa et al., 1999b). The active sites in endonucleases with the restriction endonuclease-like fold coordinate up to three metals depending upon the enzyme.

RecJ-like fold

This fold was recently identified by the determination of the RecJ nuclease structure (Yamagata et al., 2002) (Figure 3d). Previous sequence analyses have shown that this family includes RecJ and the phosphoesterases, which contain conserved phosphoesterase motifs (Aravind and Koonin, 1998a). The structure revealed a novel fold, which consists of a five-stranded parallel β-sheet flanked by six α-helices. The strand order of the β-sheet is 21345. On one side of the β-sheet, four phosphoesterase motifs form a cluster, which contains five invariant aspartates and two conserved histidines. The structure of the crystal, grown in the presence of 100 mM MnCl2, exhibits a strong metal peak coordinating three of the aspartates and one of the histidines. These residues, which constitute part of the active site, are likely to participate in the cleavage reaction.

Metallo-dependent phosphatase fold

Mre11 and several phosphatases, including the purple acid phosphatase and the ser/thr phosphatases, share this fold (Figure 3e). The core structure contains two β-sheets, which are sandwiched by α-helices to form a four-layered structure. The primary sequence of this family contains the conserved phosphoesterase motifs usually constituted by six histidines, three aspartates, and an asparagine, which form a cluster on one side of the β-sheet. The cocrystal structure of Mre11 with Mn and dAMP shows two manganese ions bound to the active site, and these two metals are simultaneously coordinated to the phosphate moiety, thus mimicking the product-bound state (Hopfner et al., 2001). The active sites of the ser/thr phosphatases bind two metals (zinc and iron) with a similar coordination scheme (Griffith et al., 1995).

DNaseI-like fold

This fold is found in DNaseI, ExoIII, and Ape1 (Figure 3F). It is also observed in some phosphatases, such as inositol 5-phosphatase. These nucleases share a four-layered structure containing an α/β sandwich, as found in the metallo-dependent phosphatases, although the β-sheet topology and the environments around the active sites are different. The active site is located on one side of the β-sheet, which assembles several conserved acidic residues. The crystal structures of DNaseI (Suck et al., 1988) and ExoIII (Mol et al., 1995) revealed a single metal ion bound to the active site. On the other hand, one (Gorman et al., 1997) or two (Beernink et al., 2001) metals were observed in the free form of Ape1. The Ape1-DNA complex structure revealed one metal, coordinated with the acidic residues and the cleaved phosphate in the active site (Mol et al., 2000b).

TIM β/α barrel fold

The TIM barrel was first observed in triosephosphate isomerase, and is now known to be the most ubiquitous fold adopted by various enzymes with diverse functions (Farber and Petsko, 1990) (Figure 3g). It forms the α8/β8 barrel structure, where a barrel-like parallel β-sheet is surrounded by eight α-helices. In this fold, the key residues for the enzymatic activity are usually located on the C-terminal side of the barrel. The structure of E. coli endoIV was the first DNA repair enzyme structure with the TIM barrel (Hosfield et al., 1999). The active site contains a cluster of three zinc ions coordinated by histidines and aspartates. The endoIV-DNA complex structure revealed how these zinc ions coordinate the cleaved AP site.

His-Me finger endonuclease fold

T4 endonuclease VII (T4 endoVII) and several other nucleases, such as the colicin nucleases, Serratia nuclease and I-PpoI intein, contain this folding motif (Figure 3h). It is usually embedded as a constituent of larger architectures. The core fold is a β-hairpin flanked by two helices. Within the hairpin, several histidines and acidic residues form a cluster and coordinate a catalytically important divalent metal. In the case of T4 endoVII, a single metal ion is coordinated to aspartate, glutamate, and asparagines (Raaijmakers et al., 1999). The I-PpoI-DNA complex structure revealed that a histidine lies within the distance of hydrogen-bond from the scissile phosphate group in the metal-containing active site (Galburt et al., 1999).

DNA recognition by DNA repair nuclease

The binding modes of DNA nucleases are roughly divided into two categories, corresponding to non-specific and specific associations. Both modes are important for efficient and accurate recognition between enzymes and DNA. Non-specific DNA binding allows enzymes to scan for target sequences or damage by a rapid diffusion process along the DNA. Once the nuclease finds its proper target, specific interactions are made to dock the active site residues correctly to the chemical groups within the DNA for cleavage. These two binding modes have been visualized within the crystal structures of the type II restriction endonucleases (reviewed in Pingoud and Jeltsch, 2001). In the cases of EcoRV, BamHI, and PvuII, the non-specific binding involves a weak association, which is contributed by an electrostatic interaction between the minimum surface area of the protein and the DNA, and the overall shape of the DNA remains in the canonical B-form, without serious deformations. By contrast, in the specific complex, the DNA is buried within the deep cleft of the protein in a sequence-specific manner, accompanied by the remarkable deformation of the DNA duplex, which is required for the cleavage by the enzyme.

This scheme can be generally applied to DNA repair nucleases as well. The nuclease surfaces are rich in basic residues, which form positive surfaces competent for electrostatic interactions with DNA. Some nucleases, such as MutH or Vsr, which both share the restriction endonuclease fold, possess partial competences for sequence-specific recognition, just like restriction endonucleases. However, most DNA repair nucleases recognize certain mismatches, forms of damages, or particular backbone structures of DNA. Therefore, they require additional and unique binding mechanisms for specific interactions with DNA. Although the information available for DNA repair nuclease-DNA complexes is limited, they can still provide considerable insights into such recognition mechanisms.

Base flipping out

Base flipping out has been observed in many DNA glycosylases and methyltransferases (reviewed in Roberts and Cheng, 1998; Vassylyev and Morikawa, 1997; Mol et al., 2000a; Parikh et al., 2000). The flipping-out of a base is defined as the local conformational change of a DNA duplex, where a base is swung out from inside of the helix into an extrahelical position and is usually inserted into the binding pocket of the protein. The space created by this process of base pair disruption is occupied by protein atoms, which are often involved in catalytic reactions. This mechanism is observed in the two crystal structures of the AP endonucleases, Ape1 and EndoIV (Figure 4a,b), which were both complexed with DNA duplexes containing an AP site in the middle. These two structures showed a similar base flipping, but different fitting modes, between the DNA and the proteins.

Figure 4
figure4

DNA recognition by DNA repair nuclease. Ribbon diagram of the DNA repair nuclease. (Left panel) Overall structure of the protein-DNA complex. (Right panel) Close-up view of the boxed region. Proteins are represented by a yellow ribbon diagram, and the side chains involved in DNA recognition are displayed and numbered with a stick model. The bound DNA is shown as a white stick model, and the flipped out nucleotides are colored red. The observed metals are shown as spheres. Blue, zinc; light blue, manganese; red, magnesium; gray, calcium. (a) endoIV (b) Ape1 (c) Vsr (d) RB69 polymerase exonuclease domain

In the Ape1-DNA complex, the abasic nucleotide was flipped out into the enzyme pocket (Mol et al., 2000b) (Figure 4a). The gap was filled on the minor groove side by two methionines (Met270, Met271) and on the major groove side by arginine (Arg177). These insertions generate a sharp kink of the DNA duplex at the abasic site. The comparison of the free form with the complex revealed a small difference, suggesting that the surface of the enzyme contains a preformed pocket to be filled by the flipped out base. Thus, it is likely that Ape1 searches its target by scanning for a possible base flipping site. Once Ape1 finds the target, the base flips out into the enzyme pocket, and the remaining gap is occupied by the inserted arginine to stabilize the protein-DNA complex. Biochemical experiments confirmed the role of this arginine, which when mutated to alanine, resulted in elevated enzyme turnover. (Mol et al., 2000b).

In the endoIV-DNA complex, an abasic site is similarly flipped out into the protein pocket (Hosfield et al., 1999) (Figure 4b). However, the conformation of the DNA duplex is drastically different from that of Ape1-DNA. The orphan base opposite the abasic site also occupies an extrahelical position. Consequently, the DNA duplex is sharply bent (90°) at the abasic site. The gap made by both flipped out nucleotides is filled by arginine (Arg37), tyrosine (Tyr72), and leucine (Leu73) inserted from the minor groove. In contrast to the preformed pocket of Ape1, the recognition loops of endoIV undergo a drastic conformational change upon DNA binding. The residues involved in base flipping are located in this loop. It is likely that endoIV scans the DNA duplex on the minor groove side by this DNA recognition loop. Once the enzyme finds the target, it inserts all of the DNA-penetrating residues, and flips the two bases into extrahelical positions.

Insertion of aromatic side chains

Another important factor in the recognition between repair enzymes and DNA is the insertion of aromatic amino acids into DNA duplexes. This is different from the insertion of amino acid side chains, which fill up the gap created by a base-flipping out. A representative case was observed in the Vsr-DNA complex (Tsutakawa et al., 1999a) (Figure 4c). Vsr recognizes a TG wobble mismatch base pair located in a five base pair long recognition sequence. In the close vicinity of the mismatch, Vsr intercalates three conserved aromatic amino acids (Phe67, Trp68, Trp86) from the major groove. In addition to the inserted helix from the minor groove, this insertion expands the space between the TG mismatch and the adjacent base pair, while the base pair itself is not disrupted. A similar insertion of aromatic residues was observed in the MutS-DNA complex, where the aromatic side chain of a conserved phenylalanine was inserted next to a mismatched or gapped base pair (Lamers et al., 2000; Obmolova et al., 2000).

The exonuclease domain of DNA polymerase uses aromatic residues for the correct positioning of the nucleotides (Figure 4d). In the editing complex of RB69 DNA polymerase with its substrate DNA, two single-stranded nucleotides are located in the groove of the exonuclease domain (Shamoo and Steitz, 1999). One of the nucleotides is held by forming a hydrogen bond with the side chain of Arg260. Another nucleotide, whose backbone is cleaved, is located more deeply within the exonuclease pocket, and is segregated from the remaining region by the insertion of two aromatic side chains (Phe123 and Phe221) to separate the two nucleotides. These phenylalanines create a wall, and thus the base is correctly positioned within the active site pocket.

In vivo and in vitro experiments, measuring the UV sensitivity and probing with potassium permanganate, have demonstrated that in E. coli RuvC, the aromatic side chain of Phe69 plays a crucial role in specific recognition with the Holliday junction (Yoshikawa et al., 2001). Phe69 lies in the protruding loop and directs its side chain into the catalytic cleft, which accommodates one of the DNA duplexes. A similar residue is also present in the yeast structural homolog, Ydc2, whereas it is absent in another yeast homolog, Cce1. Consequently, the detailed structural view of recognition mechanism between RuvC and the junction DNA is required to solve the complex directly.

Active site environments of DNA repair nucleases

All nucleases cleave the same phosphodiester bond, to leave 5′-phosphate and 3′-OH groups at the produced segments. Similar reactions are conducted by phosphatases and ribozymes, although their catalytic mechanisms have not been clarified yet. The overall aspect of this enzymatic scheme is that the attacking water is activated by a general base in the nuclease active center, which usually bears a metal cofactor. This activation is performed by protein side chains or divalent metals. The activated water is converted to a hydroxide, which attacks the phosphate, thus forming the transition state intermediate. There are two modes for this nucleophilic substitution: associative and dissociative. The associative mechanism involves the formation of a pentacovalent intermediate with a hydroxide, followed by the release of a leaving group. In this mechanism, a general base is required to generate the hydroxide, and a general acid is needed to stabilize the leaving group. The dissociative mechanism, on the other hand, does not require this general acid and general base, and they form a metaphosphate intermediate, which requires more stabilization of the transition intermediate. Many nucleases are assumed to follow the associative mechanism, while alkaline phosphatase uses the dissociative mechanism.

A large number of nucleases utilize metal cofactors for the hydrolytic reaction. They are proposed to play any one or a combination of the following roles (Figure 5) (Jencks, 1969): (1) positioning the substrate and/or the attacking nucleophile; (2) enhancing the nucleophilicity of the phosphate at the scissile bond; (3) activating the nucleophile; (4) neutralizing the negative charge in the transition state; (5) facilitating the departure of the leaving group. To examine these roles, various metals are recruited to the nuclease active sites. While the utilized metal may differ, depending upon the nuclease, magnesium or manganese is the most common metal for catalysis, and in rare cases, zinc is used. The magnesium ion appears to be transiently recruited to the active sites, whereas zinc and manganese are more tightly bound to the catalytic centers.

Figure 5
figure5

Schematic diagram of cleavage by DNA repair nucleases. X, Y, Z-H denote general base, Lewis acid, and general acid, respectively. Numbers in circles indicate reaction steps where the metal cofactors may be involved

EndoIV contains three zincs, which are coordinated by five histidines, two glutamates, and two aspartates, in addition to two water molecules (Figure 6a). These metals are so tightly coordinated to the enzyme that even EDTA cannot chelate them (Levin et al., 1991). Two of the three zinc atoms are likely to be involved in generating the attacking nucleophile, in cooperation with the carboxyl side chain of Glu261. Furthermore, all three of the metals coordinate the phosphate moiety after cleavage (Hosfield et al., 1999).

Figure 6
figure6

Active site of DNA repair nuclease. Close-up view of the nuclease active site, shown in a stereo diagram. Residues involved in the nuclease activity and the metal coordination are drawn in stick models. The coloring scheme is same as in Figure 4. (a) endoIV (b) Mre11 (c) Ape1 (d) Vsr (e) RB69 polymerase exonuclease domain

Mre11 coordinates two manganese ions through five histidines, two aspartates, and one asparagine (Figure 6b) (Hopfner et al., 2001). The two manganese ions directly coordinate the phosphate moiety of the dAMP. When magnesium is substituted for manganese, they can only occupy one of the two metal binding sites, and the nuclease is inactive. This indicates that both metals are required for the nuclease activity.

As for the nucleases that require magnesium cations for catalyis, the number of metals and their positions in the active sites are more ambiguous. They are coordinated with protein atoms in a more transient manner. This relatively weak binding, and the fact that the electron number of the magnesium cation is comparable to a water molecule, make the clear identification of the metal positions more difficult. In addition, the number of bound metals may change, depending upon different crystallization conditions. The free form structure of the Ape1 crystal, obtained under acidic conditions, revealed a single, bound metal (Samarium) (Gorman et al., 1997), whereas the crystal obtained at a neutral pH contained two metals (Lead) in the active site (Beernink et al., 2001). These Ape1 data indicate that the metals occupy multiple sites, which are affected by the protonation of the acidic residues. It appears that two metals are required for catalysis, since Ape1 is only active at a neutral pH. However, the actual numbers and the role of each metal cannot be clarified at the moment, because the structure of the Ape1-DNA complex was obtained under acidic conditions, and only one manganese ion is bound to the product DNA cleaved at the abasic site (Figure 6c) (Mol et al., 2000b). Similar ambiguity with respect to the number of metals was reported for RNaseHI, such as one magnesium (Katayanagi et al., 1993) and two manganeses (Goedken and Marqusee, 2001). In the Vsr-DNA complex structure, two magnesium ions are clearly observed in the active site, with one of the metals holding both the 5′ phosphate and 3′ OH groups (Figure 6d, Tsutakawa et al., 1999a). Two metals are also found in the exonuclease domain of polymerases (Calcium) (Figure 6e) (Shamoo and Steitz, 1999) and in T7 endoI (Manganese) (Hadden et al., 2002), although the two sites are not equivalent between the two enzymes, and one of the two metals shows partial occupancy.

Future perspectives

With the rapid accumulation of metal binding site information, various catalytic mechanisms have been proposed, including the classical two metal binding mechanism (Beese and Steitz, 1991). However, it appears to us that the actual numbers and positions of the metals involved in catalysis are too broadly varied from enzyme to enzyme to describe their hydrolytic mechanisms by a unified catalytic scheme. More detailed structural information, hopefully combined with biochemical data, is essential to obtain clear insights into the metal dependent nuclease mechanisms. Meanwhile, the large diversity in nuclease architectures suggests that they can specifically recognize DNA substrates by virtue of the large variety of surface properties, which were adopted through selection over an extremely long period. In particular, the nucleases involved in DNA repair have acquired a special damage recognition system. At present, much of the structural information is based on that of prokaryotic and archaeal proteins. Eukaryotic nucleases obviously hold more complicated structures and properties, because they must bear eukaryote-specific regulatory mechanisms involving protein–protein and protein-DNA interactions. Further 3D structural characterizations of the eukaryotic DNA repair nucleases should provide additional variations or conserved architectures of protein folding, while structural analyses of their complexes with DNA substrates will clarify the recognition mechanisms.

References

  1. Aravind L, Koonin EV . 1998a Trends Biochem. Sci. 23: 17–19

  2. Aravind L, Koonin EV . 1998b Nucleic Acids Res. 26: 3746–3752

  3. Aravind L, Walker DR, Koonin EV . 1999 Nucleic Acids Res. 27: 1223–1242

  4. Beernink PT, Segelke BW, Hadi MZ, Erzberger JP, Wilson III DM, Rupp B . 2001 J. Mol. Biol. 307: 1023–1034

  5. Beese LS, Steitz TA . 1991 EMBO J. 10: 25–33

  6. Blunt T, Finnie NJ, Taccioli GE, Smith GC, Demengeot J, Gottlieb TM, Mizuta R, Varghese AJ, Alt FW, Jeggo PA . 1995 Cell, 80: 813–823

  7. Boddy MN, Gaillard PH, McDonald WH, Shanahan P, Yates III JR, Russell P . 2001 Cell 107: 537–548

  8. Boddy MN, Lopez-Girona A, Shanahan P, Interthal H, Heyer WD, Russell P . 2000 Mol. Cell. Biol. 20: 8758–8766

  9. D'Amours D, Jackson SP . 2002 Nat. Rev. Mol. Cell. Biol. 3: 317–327

  10. de Boer J, Hoeijmakers JH . 2000 Carcinogenesis 21: 453–460

  11. Farber GK, Petsko GA . 1990 Trends Biochem. Sci. 15: 228–234

  12. Fijalkowska IJ, Schaaper RM . 1996 Proc. Natl. Acad. Sci. USA 93: 2856–2861

  13. Galburt EA, Chevalier B, Tang W, Jurica MS, Flick KE, Monnat Jr RJ, Stoddard BL . 1999 Nat. Struct. Biol. 6: 1096–1099

  14. Goedken ER, Marqusee S . 2001 J. Biol. Chem. 276: 7266–7271

  15. Goldsby RE, Lawrence NA, Hays LE, Olmsted EA, Chen X, Singh M, Preston BD . 2001 Nat. Med. 7: 638–639

  16. Gorman MA, Morera S, Rothwell DG, de La Fortelle E, Mol CD, Tainer JA, Hickson ID, Freemont PS . 1997 EMBO J. 16: 6548–6558

  17. Griffith JP, Kim JL, Kim EE, Sintchak MD, Thomson JA, Fitzgibbon MJ, Fleming MA, Caron PR, Hsiao K, Navia MA . 1995 Cell 82: 507–522

  18. Hadden JM, Declais AC, Phillips SE, Lilley DM . 2002 EMBO J. 21: 3505–3515

  19. Hamdan S, Carr PD, Brown SE, Ollis DL, Dixon NE . 2002 Structure (Camb) 10: 535–546

  20. Hopfner KP, Karcher A, Craig L, Woo TT, Carney JP, Tainer JA . 2001 Cell 105: 473–485

  21. Hosfield DJ, Mol CD, Shen B, Tainer JA . 1998 Cell 95: 135–146

  22. Hosfield DJ, Guan Y, Haas BJ, Cunningham RP, Tainer JA . 1999 Cell 98: 397–408

  23. Hwang KY, Baek K, Kim HY, Cho Y . 1998 Nat. Struct. Biol. 5: 707–713

  24. Interthal H, Heyer WD . 2000 Mol. Gen. Genet. 263: 812–827

  25. Jencks WP . 1969 Catalysis in Chemistry and Enzymology New York: McGraw Hill pp 111–115

    Google Scholar 

  26. Kaliraman V, Mullen JR, Fricke WM, Bastin-Shanower SA, Brill SJ . 2001 Genes Dev. 15: 2730–2740

  27. Katayanagi K, Okumura M, Morikawa K . 1993 Proteins 17: 337–346

  28. Kirchgessner CU, Patil CK, Evans JW, Cuomo CA, Fried LM, Carter T, Oettinger MA, Brown JM . 1995 Science 267: 1178–1183

  29. Lamers MH, Perrakis A, Enzlin JH, Winterwerp HH, de Wind N, Sixma TK . 2000 Nature 407: 711–717

  30. Levin JD, Shapiro R, Demple B . 1991 J. Biol. Chem. 266: 22893–22898

  31. Lieber MR . 1997 Bioessays 19: 233–240

  32. Ma Y, Pannicke U, Schwarz K, Lieber MR . 2002 Cell 108: 781–794

  33. Modrich P, Lahue R . 1996 Annu. Rev. Biochem. 65: 101–133

  34. Mol CD, Hosfield DJ, Tainer JA . 2000a Mutat. Res. 460: 211–229

  35. Mol CD, Izumi T, Mitra S, Tainer JA . 2000b Nature 403: 451–456

  36. Mol CD, Kuo CF, Thayer MM, Cunningham RP, Tainer JA . 1995 Nature 374: 381–386

  37. Morrison A, Johnson AL, Johnston LH, Sugino A . 1993 EMBO J. 12: 1467–1473

  38. Moshous D, Callebaut I, de Chasseval R, Corneo B, Cavazzana-Calvo M, Le Deist F, Tezcan I, Sanal O, Bertrand Y, Philippe N, Fischer A, de Villartay JP . 2001 Cell 105: 177–186

  39. Mullen JR, Kaliraman V, Ibrahim SS, Brill SJ . 2001 Genetics 157: 103–118

  40. Murzin AG, Brenner SE, Hubbard T, Chothia C . 1995 J. Mol. Biol. 247: 536–540

  41. Obmolova G, Ban C, Hsieh P, Yang W . 2000 Nature 407: 703–710

  42. Parikh SS, Putnam CD, Tainer JA . 2000 Mutat. Res. 460: 183–199

  43. Petit C, Sancar A . 1999 Biochimie 81: 15–25

  44. Pingoud A, Jeltsch A . 2001 Nucleic Acids Res. 29: 3705–3727

  45. Prakash S, Prakash L . 2000 Mutat. Res. 451: 13–24

  46. Raaijmakers H, Vix O, Toro I, Golz S, Kemper B, Suck D . 1999 EMBO J. 18: 1447–1458

  47. Ramotar D, Popoff SC, Gralla EB, Demple B . 1991 Mol. Cell. Biol. 11: 4537–4544

  48. Roberts RJ, Cheng X . 1998 Annu. Rev. Biochem. 67: 181–198

  49. Shamoo Y, Steitz TA . 1999 Cell 99: 155–166

  50. Sharples GJ . 2001 Mol. Microbiol. 39: 823–834

  51. Shevelev IV, Hubscher U . 2002 Nat. Rev. Mol. Cell. Biol. 3: 364–376

  52. Stewart GS, Maser RS, Stankovic T, Bressan DA, Kaplan MI, Jaspers NG, Raams A, Byrd PJ, Petrini JH, Taylor AM . 1999 Cell 99: 577–587

  53. Suck D, Lahm A, Oefner C . 1988 Nature 332: 464–468

  54. Tsutakawa SE, Jingami H, Morikawa K . 1999a Cell 99: 615–623

  55. Tsutakawa SE, Muto T, Kawate T, Jingami H, Kunishima N, Ariyoshi M, Kohda D, Nakagawa M, Morikawa K . 1999b Mol. Cell 3: 621–628

  56. Tsutakawa SE, Morikawa K . 2001 Nucleic Acids Res. 19: 3775–3783

  57. Vassylyev D, Morikawa K . 1997 Curr. Opin. Struct. Biol. 7: 103–109

  58. Wilson III DM, Thompson LH . 1997 Proc. Natl. Acad. Sci. USA 94: 12754–12757

  59. Xanthoudakis S, Miao G, Wang F, Pan YC, Curran T . 1992 EMBO J. 11: 3323–3335

  60. Yamagata A, Kakuta Y, Masui R, Fukuyama K . 2002 Proc. Natl. Acad. Sci. USA 99: 5908–5912

  61. Yang W . 2000 Mutat. Res. 460: 245–256

  62. Yoshikawa M, Iwasaki H, Shinagawa H . 2001 J. Biol. Chem. 276: 10432–10436

Download references

Acknowledgements

We regret that the limit of space may have not allowed us to site all works in the field. We thank Kayoko Komori for critical reading of the manuscript and helpful comments. T Nishino is a research fellow of the Japan society for the promotion of sciences. This research was partly supported by NEDO (New Energy and Industrial Technology Development Organization).

Author information

Affiliations

Authors

Corresponding author

Correspondence to Kosuke Morikawa.

Rights and permissions

Reprints and Permissions

About this article

Cite this article

Nishino, T., Morikawa, K. Structure and function of nucleases in DNA repair: shape, grip and blade of the DNA scissors. Oncogene 21, 9022–9032 (2002). https://doi.org/10.1038/sj.onc.1206135

Download citation

Keywords

  • DNA repair
  • nuclease
  • metal-dependent cleavage
  • protein-DNA interaction
  • structure-function relationships

Further reading

Search

Quick links