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Department of Biochemistry & Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, USA
Robert Hromas
Department of Medicine, University of Florida and Shands Health Care System, Gainesville, FL, USA
Suk-Hee Lee*
Department of Biochemistry & Molecular Biology, Indiana University School of Medicine, Indianapolis, Indiana, USA
*Address all correspondence to:
1. Introduction
Ionizing radiation (IR) and various cytotoxic chemicals including reactive oxygen species (ROS) induce DNA double-strand breaks (DSBs) when they attack the phosphate backbones of the two DNA strands simultaneously. DSBs, once generated, not only cause a discontinuity in the genetic code, but also are vulnerable to further loss of DNA from a nuclease attack or the formation of abnormal DNA structures from chromosomal translocation, all of which can significantly increase genomic instability leading to cancer. Repair of DSB damage is therefore crucial for maintaining the physical and genetic integrity of the genome.
DNA damage sensors are the first responder to various types of DNA damages. Upon DSB damage, Mre11–Rad50–Nbs1 (MRN) complex initially recognizes DNA damage, and recruits and activates the ataxia-telangiectasia mutated (ATM) through protein interaction with Nbs1 (Fig. 1) [1, 2]. ATM is a member of the phosphoinositide 3-kinase (PI3K)-related protein kinase (PIKK) family of serine/threonine protein kinases that phosphorylates a number of target proteins containing conserved phosphorylation motif (SQ/TQ) in response to DNA damage [3] that include MRN complex, a histone variant, H2AX, a checkpoint mediator, MDC1, a checkpoint kinase, CHK2 and p53 [4]. Phosphorylations of MRN complex, H2AX and MDC1 are necessary for recruitment of the factors involved in signal transduction and homologous recombination (HR) to facilitate the repair process [5-9]. A marginal repair defect was observed in AT cells, which could be due to the reduced efficiency of homologous recombination [10]. Damage-induced phosphorylation of CHK2 and activation of p53 also induce the cell cycle arrest at the G1 phase [6, 11, 12].
Figure 1.
Three major DSB repair pathways in mammals.Following recognition of DSB damage by MRN complex and ATM, leading to phosphorylation of H2AX, DSB repair can occur through nonhomologous end joining (NHEJ), homologous recombination (HR), or microhomology-mediated end joining (MMEJ) repair pathways. The error-free pathway of HR in the late S- and G2-phases of the cell cycle requires a sister chromatid to restore broken DNA to its original sequence, whereas the error-prone pathway of NHEJ often processes the DNA by adding or deleting nucleotides before joining the ends. In some circumstances one or more of the broken ends is refractory to Ku mediated NHEJ. In this case, MMEJ can proceed by nucleolytic processing and resection of the 3’-end until a short region of complimentary bases is revealed. Pairing of this microhomology stabilizes the broken ends, displaced flaps are removed and ligation can occur. Although many of the proteins involved in these major DSB repair pathways have been identified, the precise mechanisms involved remain poorly understood.
In mammals, DSB damages are largely repaired by non-homologous end joining (NHEJ) pathway throughout the cell cycle that directly ligates the break ends without the need for a homologous template (Fig. 1), so NHEJ is an error-prone repair pathway. Microhomology-mediated end joining (MMEJ) shares the repair proteins with NHEJ pathway, except that it uses a short patch (5-25 base pairs) of homologous sequences to align the broken strands before joining (Fig. 1). When a break occurs a homology of 5-25 complementary base pairs on both strands is identified and used as a basis for which to align the strands with mismatched ends. Once aligned, any overhang or mismatched bases on both strands are removed and any missing nucleotides are inserted. MMEJ works by ligating the mismatched hanging strands of DNA, removing overhanging nucleotides and filling in the missing base pairs. MMEJ repair occurs during the S-phase of the cell cycle, as opposed to the G0/G1 and early S phases in NHEJ. MMEJ ligates the DNA strands without checking for consistency and causes deletions, since it removes base pairs (flaps) on the strand in order to align the two pieces; it is an error-prone repair pathway and results in deletion mutations. In most cases, a cell uses MMEJ only when the NHEJ repair is not available or unsuitable due to the disadvantage posed by introducing deletions into the genetic code. When a sister chromatid is available during late S- and G2-phases of the cell cycle, DSB damage can also be repaired by homology-directed repair, called homologous recombination (HR) (Fig. 1). This requires extensive 5’-3’ resection of DNA to generate a 3’ single-stranded tail. This is then displaced by the RAD51 recombinase, which forms a nucleoprotein filament which invades a homologous DNA duplex. This process named strand exchange forms a DNA crossover or Holliday junction which provides a primer to initiate new DNA synthesis. At this point there can be several outcomes. In synthesis dependent strand annealing the newly synthesized DNA reverts back to its original partner where it can be used as a template to complete repair. Alternatively for homologous recombination, the Holliday junction migrates away from the initial point of exchange (branch migration) until the junction is resolved by nucleolytic cleavage of either the crossed strands or non-crossed strands of the junction. Resolution of the two Holliday junctions in different orientations results in the exchange of flanking markers (crossover), whereas resolution in the same orientation does not result in exchange of flanking markers (non-crossover).
Since NHEJ repair involves a direct rejoining of the separated DNA ends, it requires the coordinated assembly of damage-responsive proteins at the damage site. DSB repair through NHEJ is initiated by binding Ku70-Ku80 complex to the DSB ends (Fig. 2). The Ku70/80 complex first binds to the DNA ends and recruits DNA-dependent protein kinase catalytic subunit (DNA-PKcs), a 465-kDa ser/thr kinase that mediates synapsis of the ends and then undergoes activation of its kinase. DNA-PKcs is a member of PIKK family [13], but its contribution to checkpoint response is insignificant. Kinase activity is required for NHEJ, but its function remains unclear. Rather, it phosphorylates multiple proteins involved in NHEJ [14]. Artemis, a nuclease, and PNK, a kinase/phosphatase, process the ends [15-17], and DNA ligase IV, a complex with XRCC4, ligates two DSB ends (Fig. 2) [18, 19]. The recruitment of the XRCC4-DNA ligase IV (Lig4) complex is essential for the final step of ligation. XLF (also known as Cernunnos) is known to stimulate Lig4 in vitrothrough its interaction with XRCC4. Although DNA end joining systems in mammals are dependent on above-mentioned factors (Ku70/80, DNA-PKcs, and XRCC4/Lig4), additional factors are required for end processing during NHEJ. Artemis exists in a complex with DNA-PKcs and has nuclease activity. Mre11 and Artemis possess 3’-5’ and 5’-3’ exonuclease, respectively, both of which may be involved in promoting the joining of noncomplementary ends via utilizing microhomologies near the ends of the DSB. The Werner syndrome protein (WRN) with its DNA cleavage activity stimulated by Ku complex is also a potential player in DNA end processing. Others implicated in DNA end processing include FEN-1, PNK, and DNA polymerases μ and λ. In addition, DNA polymerase(s) are also likely involved in the gap filling of NHEJ reaction. Metnase (also known as SETMAR) is a new comer in DSB repair pathways that not only methylates histone H3 lysine 36 at DSB sites but also plays several other roles in the joining of DSB damages. Although this review discussed current issues on DSB repair in general, it mainly focuses on the emerging roles of Metnase in DSB repair pathway.
Figure 2.
Nonhomologous end joining (NHEJ) repair pathway in mammals.When a DSB is introduced, Ku binds to the DNA because of its high affinity for DNA ends. The binding of Ku elicits conformational changes that allow it to bind DNA-PKcs. Ku may also serve as an alignment factor for the accuracy of NHEJ. Upon the assembly of DNA-PK on DNA breaks, this DNA repair complex activates its serine/threonine protein kinase activity and phosphorylates target substrates such as Artemis that colocalize at the ends of broken DNA prior to end processing and end joining events.
2. Human SET-Transposase chimeric protein in DSB repair
Transposases mediate DNA movement by recognizing both ends of transposon to excise the element from one site and insert it at other location in the genome, a process which can be repeated multiple times for a given segment [20-25]. It is likely that the ends are brought together and form a synaptic complex comprising two transposase molecules and the two ends of the corresponding element [25-28]. While transposase has played an important evolutionary role accounting for half of the present organization of the human genome [29], transposase activity was thought to be extinct in humans because unregulated DNA mobility could be highly deleterious in a long lived organism. To date, only one example of an intact copy of the Hsmar1 transposase domain has been identified within the human genome [30]. The Hsmar1 transposon, a class II transposable element, is an ancient element within the human genome introduced at least 50 million years ago in ancestral primates [23]. This “functional” Hsmar1 transposase domain exists as a chimeric fusion protein, Metnase (also known as SETMAR), which resulted from an insertion of the Hsmar1 transposon downstream of a SET gene (suppressor of variegation 3-9, enhancer-of-zeste, trithorax)1 encoding a histone lysine methyltransferase (HLMT), generating the SET and transposase fusion protein [23, 30]. Metnase is not found in prosimian monkeys or other mammals. Presumably this fusion event has conferred some evolutionary advantage to anthropoid primates as the activities of both the SET domain and transposase domain have largely been retained.
The Metnase-SET domain comprises pre-SET (aa 14-118), SET (aa 120-256), and post-SET (aa 273-302) domains (Fig. 3). The pre-SET domain contains a cysteine- and histidine-rich putative Zn++ binding motif, and the SET domain has the conserved the histone lysine methyltransferase motif shared with other SET proteins in humans [31, 32]. On the other hand, the Metnase-Transposase domain contains the conserved DNA binding and the catalytic motifs (Fig. 3). Potential DNA-binding motifs in the Metnase transposase were identified by comparative sequence analysis. These include a Nuclease-associated modular DNA-binding 1 (NUMOD1) motif, residues 417-434 representing a DNA binding helix-turn-helix based on its similarity to other families [33, 34], and a helix-turn-helix (HTH) motif, residues 347-381 (Fig. 3). Although Metnase cannot perform transposition, it has been shown to retain a number of activities associated with transposases including 5’-terminal inverted repeats (TIR)-specific DNA binding [23, 35-37], DNA looping activity [25], 5’-end processing activity [25, 35, 37], and promotion of integration at a TA dinucleotide target site [25, 38]. Recent structural analysis of the Metnase transposase domain has revealed features within the catalytic site that are distinct from those of related transposases and yet were likely present within the ancestral Hsmar1transposase. However, Metnase’s DNA cleavage activity, unlike other functionally active transposases, is not coupled to its TIR-specific DNA binding [35, 37].
Figure 3.
Schematic diagram of human Metnase (SETMAR).The Pre-SET domain contains cysteine- and histidine-rich putative Zn++ binding motif. The SET domain has the HLMT motif; transposase domain contains DNA binding motifs [helix-turn-helix (HTH) and NUMOD1] and a conserved DNA cleavage (DDE-like) motif.
Metnase is widely expressed in human tissues promotes NHEJ repair and mediates genomic integration of foreign DNA [32, 35, 39]. Metnase’s involvement in NHEJ repair came from an in vivo study showing that overexpression of Metnase increased NHEJ repair, while it did not produce any significant changes in HR repair [32]. Similarly, cells treated with Metnase-specific siRNA showed a significant reduction in NHEJ repair activity in vivo. Metnase overexpression resulted in a 3-fold survival advantage after IR treatment compared to a vector control [32], further evidence of a role for Metnase in NHEJ. Metnase is also involved in genomic integration of foreign DNA [40, 41] that depends on some of the other NHEJ factors [42, 43]. Earlier study showed that a deletion of either SET or the transposase domain abrogated Metnase’s function in DNA repair, indicating that both domains are required for this function [40]. Upon DNA damage, Metnase colocalizes with other DSB repair factors and has been shown to directly interact with Pso4 [34, 36], a human homolog of the 55-kDa protein encoded by the PSO4/PRP19 gene in Saccharomyces cerevisiaethat has pleiotropic functions in DNA recombination and error-prone repair [44-47]. Metnase-mediated stimulation of DNA end joining in vivo requires both histone methyltransferase and transposase-associated activities [32], indicating that it has multiple functions in NHEJ repair. The SET-transposase fusion protein not only promotes DSB repair, but also physically interacts with Topo IIα and enhances Topo II-mediated chromosomal decatenation [24, 39], both of which are crucial for controlling DSB damage. Metnase is widely expressed, and is located at chromosome 3p26, a region of frequent abnormalities in various cancers [23, 32]. Metnase is the only known example of a protein involved in DNA repair that includes a SET domain as well as the only intact and functional Hsmar1transposase within the human genome.
DSB damage induces post-translational modification of histone proteins at the DNA damage sites, which not only is necessary for DNA damage sensing but also promotes DNA repair [48-57]. H2AX, a member of the histone H2A family, is rapidly phosphorylated in response to ionizing radiation and DNA damaging drug, generating γH2AX [50, 53, 55-57]. Phosphorylation of the histone variant H2AX occurs at the conserved C-terminal phosphatidylinositol 3-OH-kinase-related kinase (PI3KK) motif, and likely play a key role in DDR and is required for the assembly of DNA repair proteins at the sites containing damaged chromatin as well as for activation of checkpoints proteins which arrest the cell cycle progression [58-61]. DSB damage also induces non-proteolytic ubiquitylation near DNA damage site on the chromatin. DSB-induced ubiquitination is mediated by the RNF8/RNF168 ubiquitin ligase cascade [60], and has emerged as a key mechanism for restoration of genome integrity by licensing the DSB-modified chromatin to recruit genome caretaker proteins such as 53BP1 and BRCA1 near the lesions. In parallel, Sumoylation of upstream DSB regulators is also required for execution of this ubiquitin-dependent chromatin response, although its molecular basis is not clear.
Histone methylation plays a key role in and as such regulates transcription, replication, cell differentiation, genome stability, and apoptosis [62-66]. Mounting evidence points to a role for histone lysine methylation in DSB repair [67-72]. In mammalian cells, H3K79 methylation is crucial for 53BP1 localization at DSB sites and interaction with p53 in damage checkpoint activation [73]. In S. cerevisiae, loss of H3K79 methylation inhibits Rad9-dependent activation of the checkpoint kinase Rad53 following DSB damage [74, 75], and in fission yeast controls the recruitment of the damage checkpoint adaptor protein, Crb2 [76-78]. A recent study showed that DSB damage induces dimethylation of histone H3 at lysine 36 (H3K36me2) in human cells [31, 32]. Chromatin immunoprecipitation (ChIP) and immunoblot analyses indicated that H3K36me2 is actually formed at DSB sites [31]. H3-K36 is associated with chromatin opening [79-84], which may also be a part of its DSB localization via chromatin modulation. In fact, mutations at known conserved SET domain amino acids (N210S, alteration at the NHSC at 210-213 to AAAA, and the YDY at 247-249 to AAA) significantly lowered DNA end joining [32]. Two conserved amino acid sequences (210-NHSCXPN-216 and 242-EEELXXXY-249) in the Metnase-SET domain are likely responsible for the interaction with SAM since a mutation at these sites failed to interact with 3H-labeled SAM [85-88]. Levels of DSB-induced H3K36me2 strongly correlate with Metnase expression and that the mutant (D248S) lacking HLMT activity fails to generate H3K36me2, suggesting that Metnase is responsible for the induction of H3K36me2 at DSB site [32]. Considering that the D248S mutant of Metnase fails to promote NHEJ repair, dimethylation of H3K36 is likely a major function of Metnase in promoting chromosomal DSB repair. Although the mechanism by which H3K36me2 promotes DSB repair is not clear, H3K36 methylation has been linked to chromatin opening accessible to transcription regulators and DNA repair proteins [89]. H3K36me2, once formed at DSB site, may create docking sites for other repair proteins, recruiting them for transcription and DNA repair. For example, H3K36 methylation attracts the histone deacetylase Rpd3S, which compact chromatin in the middle of transcribed genes, and inhibits false initiation of transcription during the elongation phase [90]. The methyltransferase Setd2 (also known as Set2) mediates trimethylation of H3K36me3 (H3K36me3), and it binds the phosphorylated tail of RNA polymerase II, implicating a role for H3K36me3 in transcription [49, 51, 91]. Setd2 mediates H3k36me3 in mammalian cells, but not di- or mono-methylation [92], raising a possibility that Metnase or other H3K36 dimethyltransferases may be necessary to generate H3K36me2 before Setd2 acts. In Drosophila, dimethylation of H3K36 peaks adjacent to promoters and requires distinct methyltransferases than those that mediate H3K36me3 [93]. The formation of H3K36me2 might also facilitate histone eviction at the DSB site, which then facilitates an access of the repair machineries to DNA damage site. This is supported by an observation that H3K36me2 enhances the presence of MRN complex and Ku70 at the DSB site [31]. These DNA repair proteins show an increased interaction with H3K36me2 after IR, and their presence at an induced DSB also correlated with Metnase expression levels. In addition, the chromatin immunoprecipitation study revealed that H3K36me2 not only enhances the rate of association of these repair proteins with the DSB but decreases their disassociation rates as well [31]. Because the MRN and Ku complexes can bind free DNA ends at a DSB in nonchromatinized DNA, the decreased rates of disassociation are likely the more important role of H3K36me2. This implies that the main benefit of H3K36me2 in DSB repair is more likely to stabilize the repair components at the DSB than to enhance their recruitment. It is possible that dimethylation of H3K36 at DSBs was an epiphenomenon and was not responsible for enhanced localization of early DSB repair components. On the other hand, when a point mutation at H3-K36 (K36R or K36A) caused a marked decrease in both the recruitment of NBS1 and Ku70 to the DSB and in DSB repair [31], indicating that H3K36me2 is required for efficient assembly and retention of repair components at DSBs and for optimum DSB repair. The identification of dimethylated H3K36 as a chromatin modification that enhances DSB repair by NHEJ places this modification alongside and ubiquitylated H2A as DNA damage-induced histone modifications that recruit repair components to DSBs and enhance repair [31, 94, 95]. In this regard, H3K36me2 by Metnase is consistent with an NHEJ histone code, as defined in the original histone code hypothesis for transcriptional regulation as histone modifications, acting in a combinatorial fashion on histones, which specify unique downstream functions [56]. Previous reports indicate that histone methylation may be important in DNA DSB repair by homologous recombination: The DSB repair component 53BP1, which is required for proper homologous recombination, is recruited to sites of damage by methylated histone H3 lysine 79 (H3K79) and histone H4 lysine 20 (H4K20) [76, 78, 96]. However, neither H3K79 nor H4K20 methylation is induced by DNA damage [96]. H3K36me2 is likely reserved for NHEJ repair pathway, because Ku70 and Metnase are involved in DSB repair by NHEJ rather than HR repair and because the latter requires complete histone eviction adjacent to the DSB. Human cancer cells that express Metnase at high levels display enhanced resistance to treatment with radiation or chemotherapy [32, 131, 132]. The resistance mediated by Metnase could reflect improved stabilization of the assembly of DSB repair components at DSB sites due to the generation of H3K36me2 at these sites. If so, a targeting of Metnase’s HLMT activity may improve the efficacy of common cancer therapies based on DNA damaging agents.
4. DNA endonuclease activity in the joining of DSB damage
IR induces DNA double strand breaks with different ends, most of which are not directly ligatable. Therefore, they need to be processed before end joining event in all three major DSB repair pathways, with the exception of adding nucleotides opposite to 5’-overhang by DNA polymerase [97-100]. DNA end processing can be divided into two types: ssDNA cleavage that removes either a 5’- or 3’-overhang to leave a blunt end, and nuclease activity producing a deletion that is consistent with alignment of the DNA ends by base pairing in region(s) of microhomology [17, 97, 99, 101]. Several endonucleases and their binding partners have been shown to participate in end processing during DSB repair. Mre11 and Artemis possess 3’-5’ exonuclease activity and ssDNA-specific 5’-3’ exonuclease, respectively, both of which may be involved in promoting the joining of noncomplementary ends via utilizing microhomologies near the ends of the DSB [17, 100, 102-106]. MRN’s exonuclease activity is for mismatched DNA ends and pauses at sites of microhomology [100], while its endonuclease is to open fully paired hairpin DNA [105]. Artemis possesses an endonuclease activity specific for hairpins and 5′- or 3′-overhangs following phosphorylation by DNA-PKcs [17, 106], suggesting that it plays a role in V(D)J recombination repair and perhaps in removing the 5′- and 3′-overhangs of non-compatible ends during NHEJ repair. Human CtIP physically and functionally interacting with MRN is another player in DNA end processing [107]. CtIP was originally identified as a binding partner for CtBP11 and the tumor suppressor proteins RB1 [108] and BRCA1 [86, 109], and is recruited to DNA damage and complexes with BRCA1 to control the G2/M DNA-damage checkpoint [110-112]. CtIP and the MRN complex promote ATR activation and HR through mediating DSB resection [107]. The Werner syndrome protein (WRN), a RecQ-like DNA helicase also possesses 3’-5’ exonuclease activity [42, 43, 113]. Considering that WRN is phosphorylated by DNA-PKcs [113], and its DNA cleavage activity is stimulated by Ku complex [114], WRN could play a role in DNA end processing. Other DNA helicases such as Bloom (BLM) and DNA2 may also play a role in DNA end processing [85]. These two DNA helicases physically interact to each other to resect DNA in a process that is ATP-dependent and requires BLM helicase and DNA2 nuclease functions [85]. RPA is essential for both DNA unwinding by BLM and enforcing 5′-3′ resection polarity by DNA2. MRN accelerates processing by recruiting BLM to the end. In the other, EXO1 resects the DNA and is stimulated by BLM, MRN, and RPA. BLM increases the affinity of EXO1 for ends, and MRN recruits and enhances the processivity of EXO1 [85].
Metnase possesses a unique endonuclease activity that preferentially acts on ssDNA overhang of a partial duplex DNA [35]. Cell extracts lacking Metnase exhibited significantly lowered end joining activity comparable to those seen in extracts lacking DNA-PKcs or Ku80 [35], whereas cell extracts over-expressing Metnase not only stimulated DNA end joining but also showed an enhanced end processing of non-compatible ends based on DNA sequencing analysis of end joining products [32, 35, 37]. Metnase has no hairpin or loop opening activity [35], indicating that it does not play a role in V(D)J recombination. Given that DNA end processing facilitates end joining by increasing the chance for partial annealing between two non-compatible ends, Metnase’s endonuclease activity may play a direct role in stimulating DNA end joining through processing of non-compatible ends. While Metnase contributes to DNA end joining through an enhanced processing of non-compatible ends, its DNA cleavage activity cannot explain Metnase’s stimulatory role in the joining of compatible ends [32, 35, 37]. Similar to DNA-PK- and Ku80-defective cells, cell extracts lacking Metnase failed to support joining of compatible ends [32], suggesting that Metnase also has a role in the joining of compatible ends, perhaps by promoting recruitment of the XRCC4-DNA ligase 4 (Lig4) complex [115], an essential player in the ligation step through a physical interaction upon DNA damage. The DNA binding property of Metnase may assist in the localization of DNA Lig4 at the free DNA ends. In this case, Metnase is epistatically above end-processing and subsequent joining, but perhaps below free end recognition and protection, in the NHEJ cascade.
One intriguing thing is how a transposase possesses ssDNA overhang cleavage activity in the absence of TIR sequence. The Metnase-transposase domain has a conserved DDE-like motif (D483, D575, and N610) that is crucial for DNA cleavage activity (Fig. 3) [35, 37, 116]. The function of residues in the DDE-motif includes coordination of a metal ion required for catalysis in other transposases. In addition to these residues, several other residues potentially play a role in the catalytic activity of the transposase domain [116]. Based on the crystal structure of the Metnase-transposase, the active sites of the two subunits that make up the dimer are distinctly different [116]; one subunit has bound metal in the active site and the other does not [116]. Metal is bound to the active site of one molecule comprising the dimer coordinated to Asp 483 and Asp 575. Residues K445, R578, and H580 within the catalytic pocket adopt different conformations in the metal-bound vs. non-metal bound active site structures and may also play important catalytic functions in ss-overhang cleavage activity. A loop within the active site of Metnase adopts two very different conformations resulting in a translation of a full residue when superimposed such that Arg 578 is located within the active site hydrogen-bonded to Glu 484 in the non-metal bound conformation and flipped out of the active site in the metal bound conformation. Similarly, the position of His 580 is quite different in each of the two different conformations in our structure. Interestingly, each conformation of His 580 is hydrogen-bonded to Glu 484. It remains to be seen what unique feature(s) of the catalytic domain with Metnase is directly linked to its role in DNA repair and replication fork arrest as compared to traditional transposase function.
Metnase is a DNA repair factor colocalized with MRN complex and other repair factors at the DNA damage sites [36]. On the other hand, it is a transposase that has a capacity to interact with thousands of potential binding sites (TIR) in human chromosomes [23, 25, 37]. Metnase binds to a specific 19 bp sequence within the consensus Hsmar1 TIR [23, 30, 38, 117]. Similar to other Mariner transposases, the Metnase Helix-Turn-Helix (HTH) motif accounts for this binding; specifically the R432 residue within the HTH region is essential for this binding [37]. In human genomes there are a large number of miniature inverted-repeat transposable elements (MITES). If the solo TIRs are added to the number of MITES, there are approximately 7,000 potential Metnase binding sites in human genome. How does a transposase with a sequence-specific DNA binding activity get localized at the DSB sites? A recent study identified Pso4 as a Metnase binding partner that seems to play a role in Metnase localization at DSB sites [36]. Although Pso4 is Metnase’s binding partner, coimmunoprecipitation of Metnase and Pso4 also pulled down the human homolog of Spf27, a member of the Prp19 core complex involved in pre-mRNA splicing [36]. Given that Pso4 is a part of the pre-mRNA splicing complex consisting of Pso4, Cdc5L, Plrg1, and Spf27 [118], the Metnase-Pso4 complex may be a part of the bigger complex including other members of the pre-mRNA splicing complex in vivo. Although the physiologic role of the Metnase-Pso4 interaction is still unclear, cells lacking Pso4 failed to show Metnase localization at the DSB sites [36], suggesting that Pso4 play a role in the recruitment of Metnase to the DSB sites. Upon DNA damage, Pso4 is induced [46] and formed a stable complex with Metnase [36]. A recent biochemical analysis suggested several interesting implications for the architecture of the Metnase-Pso4 complex on DNA. First, Metnase dimer forms a 1:1 stoichiometric complex with Pso4 on dsDNA [35, 36]. Although both Metnase and Pso4 can independently interact with TIR DNA, Pso4 is solely responsible for binding to dsDNA once the two proteins form a stable complex [35]. This claim is based on the findings that 1) the Metnase-Pso4 complex interacted with same stoichiometric amount of non-TIR DNA as the TIR DNA, 2) the Metnase-Pso4 complex interacted with same number of TIR molecules as Metnase or Pso4 alone did, and 3) formation of the Metnase-TIR complex was significantly inhibited by excess of TIR and not by non-TIR, whereas the Metnase-Pso4-TIR complexes were equally inhibited by both TIR and non-TIR DNA [35]. It is possible that Pso4, once forming a complex with Metnase, may directly interfere with Metnase’s DNA binding domain (helix-turn-helix motif) [37]. This notion is supported by findings that Metnase bound to TIR DNA went through a conformational change and was less effective than free Metnase in interacting with Pso4 [35]. Pso4 has 6 C-terminal WD-40 repeats [119], a module that is known to interact with post-translationally modified histone 3, including dimethylated-K4 [120]. Given that Metnase HLMT activity targets H3-K4 as well as H3-K36 [32], it is possible that chromatin association of Pso4 may occur via Metnase-mediated H3-K4 methylation, while Metnase requires Pso4 for its DSB localization. Since Pso4 is induced following IR treatment in vivo[36, 46, 121], formation of a stable Metnase-Pso4 complex likely occurs in response to DNA damage.The Pso4 also undergoes structural alterations in response to DNA damage [121]. The Metnase-Pso4 complex, once formed, likely goes to nonTIR sites such as DSB sites [36], since Pso4 is solely responsible for binding to DNA in forming the Metnase-Pso4-DNA complex. It would be interesting to see whether Pso4 also affects Metnase’s other biochemical functions such as DNA cleavage activity and HLMT activity. Further structural study would be necessary to clarify this intriguing issue.
Metnase also physically interacts with DNA ligase IV (Lig4), an essential DSB repair factor involved in the final end joining step in response to DNA damage [24], which supports the observations that Metnase promoted joining of both compatible and non-compatible ends [32, 35]. It remains to be seen whether Metnase plays a direct role in the recruitment of the XRCC4- Lig4 complex via its interaction with Lig4.
DNA double-strand breaks can be generated at the replication forks when the replication machinery encounters a single-strand break (SSB) or other types of DNA adducts. Attempted replication past a SSB can generate one-sided DSB which topologically differs from DSBs introduced by IR (Fig. 4). One-sided DSB is not a natural substrate for NHEJ, so these breaks can be repaired by homologous recombination repair pathway. Otherwise, it will remain unrepaired generating chromatid breaks, or it may ligate with a DSB in a different chromosome producing radial chromosomes. Stalled replication forks can also regress to generate a chicken-foot structure with a double stranded end (Fig. 4). Such a structure is topologically distinct from IR-induced DSBs in that it encompasses a single double-strand end rather than two double-strand ends.
Metnase possesses a distinct yet undefined role in the replication stress response [122]. Its role appears to be limited to restart of stalled and/or collapsed replication forks. DNA replication analyses indicated that Metnase promotes cell survival only when cells are subjected to replication stress such as hydroxyurea (HU), camptothecin (CT), or UV treatment [122]. Interestingly, when Metnase knockdown cells were treated with HU, the percentage of stopped forks greatly increased and there was a corresponding large decrease in the percentage of continuing forks, while new forks were extremely rare in both HU treated and untreated Metnase knockdown cells [122], indicating that Metnase plays a critical role in restarting stalled replication forks. It also suggests that Metnase may regulate new origin firing when cells experience replication stress. Metnase also regulates the efficiency of replication fork restart, and possibly initiation after replication stress, but it has no effect on the speed of ongoing forks [122].
Figure 4.
Generation of DSB damage and its repair during replication.Replication forks frequently encounter blocks to their progression including lesions such as single strand breaks. Structures such as a one sided DSB or a chicken-foot structure generated by fork regression can arise as a consequence of such replication stalling and the available evidence suggests a major function of HR is to repair or resolve such lesions.
Interestingly, a recent study with poly ADP-ribose polymerase 1 (PARP-1) revealed that it recruits MRE11 to stalled replication forks [123]. MRE11 with its endonuclease activity may play a role in processing stalled forks leading to RPA recruitment and eventual restart through HR . It is possible that Metnase promotes replication fork restart by promoting NHEJ [124]. NHEJ factors involved in NHEJ are known to promote cell survival after replication stress perhaps by facilitating rejoining of DSEs at collapsed forks [125, 126]. Since each collapsed fork produces only a single broken end that is not a natural substrate for NHEJ, however, it would be highly inaccurate producing radial chromosomes. Another possibility would be that NHEJ factors promote replication fork restart indirectly through interactions with HR factors [127]. When replication fork stalls, the initial cellular response is to stabilize the replisome to prevent fork collapse. Metnase does not appear to play a role in fork stabilization as similar fractions of cells with collapsed forks were observed regardless of Metnase expression level [122]. Another mechanism by which Metnase could promote fork restart is through its interactions with replisome factors including PCNA and RAD9. Although it is not yet known whether Metnase interacts directly with these proteins, the fact that the Metnase SET domain has a conserved PIP box is highly suggestive of direct interactions. Regardless, our results clearly place Metnase at stalled replication fork. The Metnase SET domain encodes a protein methylase, and Metnase is known to methylate histone H3 and itself [124, 128]. Metnase could regulate PCNA and/or RAD9 function through trans-methylation, or it could have a more general effect through chromatin modification. In particular, Metnase targets histone H3 lysines 4 and 36, which are associated with chromatin opening, these modifications could enhance repair factor recruitment to stalled or collapsed forks. Given the well-established role of RAD9 in the intra-S checkpoint response [129], Metnase could promote fork restart by influencing checkpoint activation or downstream checkpoint-dependent processes such as inhibition of origin firing. In addition, Metnase could affect replication fork restart through its direct interaction with Topoisomerase IIα (TopoIIα). TopoIIα is proposed to relax positive supercoils that form ahead of replication forks [130]. Currently there is no information about whether supercoils persist in front of stalled forks. However, when one of the replicative polymerases encounters a blocking lesion, the other polymerase can become uncoupled and progress for a distance, producing a single-stranded gap that is bound by RPA, triggering the intra-S checkpoint [129]. This uncoupled synthesis depends on continued DNA unwinding by the MCM helicase complex, thus positive supercoils will continue to accumulate. By promoting TopoIIα-dependent relaxation of these supercoils, Metnase could help create a favorable topological state that assists in fork restart. Conceivably, this could involve restart of stalled forks that are processed to a chicken-foot structure since the resolution of such structures is likely dependent on the topological context of the stalled fork. Alternatively, at collapsed forks, the required HR-mediated invasion of the DSE into the unbroken sister chromatid, require unwinding of the sister duplex and could similarly be affected by the local topological state. Metnase may play different roles depending on the particular state of the stalled or collapsed replication fork.
7. Abnormal expression of Metnase in tumor specimens
The Metnase gene has three exons spread over 13.8 kB located at 3p26, a region of frequent abnormalities in non-Hodgkin’s lymphoma, acute and chronic lymphocytic leukemia, myeloma, myelodysplasia, hereditary prostate cancer, and breast cancer (http://cgap.nci.nih.gov/Chromosomes/Mitelman). Metnase is expressed in all human tissues tested to various extents (32), with the highest expression in placenta and ovary and the lowest expression in skeletal muscle, which is reminiscent of expression patterns of other DNA repair proteins (131). Interestingly, different transcript variants were found in both normal and cancerous tissues (23), suggesting that Metnase is broadly expressed and has an important function in human. Metnase is frequently overexpressed in leukemia and breast cancer cell lines, and importantly, downregulating Metnase greatly enhances tumor cell sensitivity to common chemotherapeutics including epididophylotoxins and anthracyclines [132, 133]. Although the precise mechanism(s) by which Metnase promotes restart of the replication fork, Metnase may be a reasonable target for the therapeutic strategies that block DNA synthesis or take advantage of inherent defects of tumor cells in replication fork restart [134, 135].
While transposase accounts for half of the present organization of the human genome, transposase activity was thought to be extinct in humans probably because unregulated transposition would directly affect genomic stability, resulting in an unacceptably high rate of apoptosis or malignancy [29]. For this reason, transposase functions have been selected against the mammalian organisms [29], which lead to a generation of the SET-Transposase chimeric protein termed Metnase with novel functions in DSB repair, replication fork arrest, and chromosome decatenation that could actually defend the genome against improper DNA movement or DSB damage (Fig. 5).
Figure 5.
Metnase contributes to genome maintenance by promoting DSB repair and chromosome decatenation.
It should be pointed out that there are no other DNA repair proteins in which the DNA cleavage and histone lysine methyltransferase activities reside within the same protein. Although the role(s) of Metnase in DSB repair and other DNA metabolism are yet to be defined, a deletion of either the SET or the transposase domain abrogated its function in DNA repair [32], indicating that both domains are essential for this function. Histone lysine methyltransferases (HLMT) is a critical participant in chromatin integrity as evidenced by the number of human diseases including cancers associated with the aberrant expression of its family members [136]. Although the underlying mechanisms of tumorigenesis are still largely unknown, Metnase HLMT targeting of H3K36 dimethylation at DSB damage sites is not only crucial for damage recognition and the early stage of DSB repair, but is also of our interest in tumorigenesis [31]. Metnase may thus be a viable anticancer target for a wide variety of tumor types. Given that altered expression of Metnase affect joining of both compatible- and non-compatible ends [24, 32, 35, 37], Metnase likely have two separate functions in the joining of DSB damage: 1) the Metnase-Lig4 interaction [24] for joining of compatible ends by promoting recruitment of Lig4 complex to DSB sites, and 2) Metnase’s structure-specific endonuclease for joining of non-compatible ends by promoting end processing (Fig. 6). Further structure-function studies would be necessary to understand how a transposase becomes an endonuclease with ss-overhang cleavage in a TIR-independent manner.
Figure 6.
Proposed role(s) for Metnase in DSB repair and chromosome decatenation.Upon DSB damage, the Ku complex first binds to the DNA ends and recruits DNA-PKcs. Metnase binding partner, Pso4 is induced upon DSB damage, which, along with the Ku70/80, likely plays a crucial role in Metnase localization at DSB sites. Metnase’s interaction with Lig4 is also induced upon DSB damage, which promotes joining of compatible ends, while Metnase’s nuclease activity plays a role in joining of non-compatible ends.
This research was supported by grants from NIH (CA151367 and CA140422).
References
1.Carney JP, et al. The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of double-strand break repair to the cellular DNA damage response. Cell 1998;93(3) 477-486.
2.Falck J, Coates J, and Jackson SP. Conserved modes of recruitment of ATM, ATR and DNA-PKcs to sites of DNA damage. Nature 2005;434(7033) 605-611.
3.Savitsky K, et al. A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science 1995;268(5218) 1749-1753.
4.Matsuoka S, et al. ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science 2007;316(5828) 1160-1166.
5.Stewart GS, et al. MDC1 is a mediator of the mammalian DNA damage checkpoint. Nature 2003;421(6926) 961-966.
6.Lou Z, et al. MDC1 is coupled to activated CHK2 in mammalian DNA damage response pathways. Nature 2003;421(6926) 957-961.
7.Costanzo V, et al. Mre11 protein complex prevents double-strand break accumulation during chromosomal DNA replication. Mol Cell 2001;8(1) 137-147.
8.Burma S, et al. ATM phosphorylates histone H2AX in response to DNA double-strand breaks. J Biol Chem 2001;276(45) 42462-42467.
9.Goldberg M, et al. MDC1 is required for the intra-S-phase DNA damage checkpoint. Nature 2003;421(6926) 952-956.
10.Jeggo PA, Carr AM, and Lehmann AR. Splitting the ATM: distinct repair and checkpoint defects in ataxia-telangiectasia. Trends Genet 1998;14(8) 312-316.
11.Canman CE, et al. Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science 1998;281(5383) 1677-1679.
12.Kang J, et al. Functional interaction of H2AX, NBS1, and p53 in ATM-dependent DNA damage responses and tumor suppression. Mol Cell Biol 2005;25(2) 661-670.
13.Hartley KO, et al. DNA-dependent protein kinase catalytic subunit: a relative of phosphatidylinositol 3-kinase and the ataxia telangiectasia gene product. Cell 1995;82(5) 849-856.
14.Dobbs TA, Tainer JA, and Lees-Miller SP. A structural model for regulation of NHEJ by DNA-PKcs autophosphorylation. DNA Repair (Amst) 2010;9(12) 1307-1314.
16.Pannicke U, et al. Functional and biochemical dissection of the structure-specific nuclease ARTEMIS. EMBO J 2004;23(9) 1987-1997.
17.Ma Y, Schwarz K, and Lieber MR. The Artemis:DNA-PKcs endonuclease cleaves DNA loops, flaps, and gaps. DNA Repair (Amst) 2005;4(7) 845-851.
18.Grawunder U, et al. Activity of DNA ligase IV stimulated by complex formation with XRCC4 protein in mammalian cells. Nature 1997;388(6641) 492-495.
19.Critchlow SE, Bowater RP, and Jackson SP. Mammalian DNA double-strand break repair protein XRCC4 interacts with DNA ligase IV. Curr Biol 1997;7(8) 588-598.
20.Ivics Z, et al. The Sleeping Beauty transposable element: evolution, regulation and genetic applications. Curr Issues Mol Biol 2004;6(1) 43-55.
21.Miskey C, et al. DNA transposons in vertebrate functional genomics. Cell Mol Life Sci 2005;62(6) 629-641.
22.Brillet B, Bigot Y, and Auge-Gouillou C. Assembly of the Tc1 and mariner transposition initiation complexes depends on the origins of their transposase DNA binding domains. Genetica 2007;130(2) 105-120.
23.Cordaux R, et al. Birth of a chimeric primate gene by capture of the transposase gene from a mobile element. Proc Natl Acad Sci U S A 2006;103(21) 8101-8106.
24.Hromas R, et al. The human set and transposase domain protein Metnase interacts with DNA Ligase IV and enhances the efficiency and accuracy of non-homologous end-joining. DNA Repair (Amst) 2008;7(12) 1927-1937.
25.Liu D, et al. The human SETMAR protein preserves most of the activities of the ancestral Hsmar1 transposase. Mol Cell Biol 2007;27(3) 1125-1132.
26.Adams CD, et al. Tn5 transposase loops DNA in the absence of Tn5 transposon end sequences. Mol Microbiol 2006;62(6) 1558-1568.
27.Richardson JM, et al. Mechanism of Mos1 transposition: insights from structural analysis. EMBO J 2006;25(6) 1324-1334.
28.Crellin P, Sewitz S, and Chalmers R. DNA looping and catalysis; the IHF-folded arm of Tn10 promotes conformational changes and hairpin resolution. Mol Cell 2004;13(4) 537-547.
29.Lander ES, et al. Initial sequencing and analysis of the human genome. Nature 2001;409(6822) 860-921.
30.Robertson HM, Zumpano KL. Molecular evolution of an ancient mariner transposon, Hsmar1, in the human genome. Gene 1997;205(1-2) 203-217.
31.Fnu S, et al. Methylation of histone H3 lysine 36 enhances DNA repair by nonhomologous end- joining. Proc Natl Acad Sci U S A 2011;108(2) 540-545.
32.Lee SH, et al. The SET domain protein Metnase mediates foreign DNA integration and links integration to nonhomologous end-joining repair. Proc Natl Acad Sci U S A 2005;102(50) 18075-18080.
33.Sitbon E, and Pietrokovski S. New types of conserved sequence domains in DNA-binding regions of homing endonucleases. Trends Biochem Sci 2003;28(9) 473-477.
34.Beck BD, et al. Regulation of Metnase's TIR binding activity by its binding partner, Pso4. Arch Biochem Biophys 2010;498(2) 89-94.
35.Beck BD, et al. Biochemical Characterization of Metnase's Endonuclease Activity and Its Role in NHEJ Repair. Biochemistry 2011;50(20) 4360-4370.
36.Beck BD, et al. Human Pso4 is a metnase (SETMAR)-binding partner that regulates metnase function in DNA repair. J Biol Chem 2008;283(14) 9023-9030.
37.Roman Y, et al. Biochemical characterization of a SET and transposase fusion protein, Metnase: its DNA binding and DNA cleavage activity. Biochemistry 2007;46(40) 11369-11376.
38.Miskey C, et al. The ancient mariner sails again: transposition of the human Hsmar1 element by a reconstructed transposase and activities of the SETMAR protein on transposon ends. Mol Cell Biol 2007;27(12) 4589-4600.
39.Williamson EA, et al. The SET and transposase domain protein Metnase enhances chromosome decatenation: regulation by automethylation. Nucleic Acids Res 2008;36(18) 5822-5831.
40.Sekiguchi JM, and Ferguson DO. DNA double-strand break repair: a relentless hunt uncovers new prey. Cell 2006;124(2) 260-262.
41.Paull TT, and Gellert M. The 3' to 5' exonuclease activity of Mre 11 facilitates repair of DNA double-strand breaks. Mol Cell 1998;1(7) 969-979.
42.Kamath-Loeb AS, et al. Werner syndrome protein. II. Characterization of the integral 3'-5' DNA exonuclease. J Biol Chem 1998;273(51) 34145-34150.
43.Shen JC, et al. Werner syndrome protein. I. DNA helicase and dna exonuclease reside on the same polypeptide. J Biol Chem 1998;273(51) 34139-34144.
44.da Silva KV, de Morais Junior MA, and Henriques JA. The PSO4 gene of S. cerevisiae is important for sporulation and the meiotic DNA repair of photoactivated psoralen lesions. Curr Genet 1995;27(3) 207-212.
45.Henriques JA, et al. PSO4: a novel gene involved in error-prone repair in Saccharomyces cerevisiae. Mutat Res 1989;218(2) 111-124.
46.Mahajan KN, and Mitchell BS. Role of human Pso4 in mammalian DNA repair and association with terminal deoxynucleotidyl transferase. Proc Natl Acad Sci U S A 2003;100(19) 10746-10751.
47.Zhang N, et al. The Pso4 mRNA splicing and DNA repair complex interacts with WRN for processing of DNA interstrand cross-links. J Biol Chem 2005;280(49) 40559-40567.
48.Wang H, et al. mAM facilitates conversion by ESET of dimethyl to trimethyl lysine 9 of histone H3 to cause transcriptional repression. Mol Cell 2003;12(2) 475-487.
49.Sun J, and Li R. Human negative elongation factor activates transcription and regulates alternative transcription initiation. J Biol Chem 2010;285(9) 6443-6452.
50.Buro LJ, Chipumuro E, and Henriksen MA. Menin and RNF20 recruitment is associated with dynamic histone modifications that regulate signal transducer and activator of transcription 1 (STAT1)-activated transcription of the interferon regulatory factor 1 gene (IRF1). Epigenetics Chromatin 2010;3(1) 16.
51.Schwartz S, Meshorer E, and Ast G. Chromatin organization marks exon-intron structure. Nat Struct Mol Biol 2009;16(9) 990-995.
52.Yoh SM, Lucas JS, and Jones KA. The Iws1:Spt6:CTD complex controls cotranscriptional mRNA biosynthesis and HYPB/Setd2-mediated histone H3K36 methylation. Genes Dev 2008;22(24) 3422-3434.
53.Ahmad A, Zhang Y, and Cao XF. Decoding the epigenetic language of plant development. Mol Plant 2010;3(4) 719-728.
54.Hake SB, Xiao A. and Allis CD. Linking the epigenetic 'language' of covalent histone modifications to cancer. Br J Cancer 2007;96 Suppl R31-39.
55.Stucki M, and Jackson SP. gammaH2AX and MDC1: anchoring the DNA-damage-response machinery to broken chromosomes. DNA Repair (Amst) 2006;5(5) 534-543.
56.Strahl BD, and Allis CD. The language of covalent histone modifications. Nature 2000;403(6765) 41-45.
57.Fernandez-Capetillo O, et al. H2AX: the histone guardian of the genome. DNA Repair (Amst) 2004;3(8-9) 959-967.
58.Podhorecka M, Skladanowski A, and Bozko P. H2AX Phosphorylation: Its Role in DNA Damage Response and Cancer Therapy. J Nucleic Acids 2010;2010 1-9
59.Moon SH, et al. Dephosphorylation of gamma-H2AX by WIP1: an important homeostatic regulatory event in DNA repair and cell cycle control. Cell Cycle 2010;9(11) 2092-2096.
60.Yan J, and Jetten AM. RAP80 and RNF8, key players in the recruitment of repair proteins to DNA damage sites. Cancer Lett 2008;271(2) 179-190.
61.Downs JA. Chromatin structure and DNA double-strand break responses in cancer progression and therapy. Oncogene 2007;26(56) 7765-7772.
62.Hayashi M, Chin GM, and Villeneuve AM. C. elegans germ cells switch between distinct modes of double-strand break repair during meiotic prophase progression. PLoS Genet 2007;3(11) e191.
63.Reardon JT, Cheng Y, and Sancar A. Repair of DNA-protein cross-links in mammalian cells. Cell Cycle 2006;5(13) 1366-1370.
64.Pasierbek P, et al. A Caenorhabditis elegans cohesion protein with functions in meiotic chromosome pairing and disjunction. Genes Dev 2001;15(11) 1349-1360.
65.Moens PB, et al. Meiosis in a temperature-sensitive DNA-synthesis mutant and in an apomictic yeast strain (Saccharomyces cerevisiae). Philos Trans R Soc Lond B Biol Sci 1977;277(955) 351-358.
66.Rhodes MC, et al. Assessing a Theoretical Risk of Dolutegravir-Induced Developmental Immunotoxicity in Juvenile Rats. Toxicol Sci 2012.
67.Kassmeier MD, et al. VprBP binds full-length RAG1 and is required for B-cell development and V(D)J recombination fidelity. EMBO J 2012;31(4) 945-958.
68.Dalal I, et al. Novel mutations in RAG1/2 and ADA genes in Israeli patients presenting with T-B-SCID or Omenn syndrome. Clin Immunol 2011;140(3) 284-290.
69.Grundy GJ, Yang W, and Gellert M. Autoinhibition of DNA cleavage mediated by RAG1 and RAG2 is overcome by an epigenetic signal in V(D)J recombination. Proc Natl Acad Sci U S A 2010;107(52) 22487-22492.
70.Arnal SM, et al. Non-consensus heptamer sequences destabilize the RAG post-cleavage complex, making ends available to alternative DNA repair pathways. Nucleic Acids Res 2010;38(9) 2944-2954.
71.Grundy GJ, et al. Initial stages of V(D)J recombination: the organization of RAG1/2 and RSS DNA in the postcleavage complex. Mol Cell 2009;35(2) 217-227.
72.Zhang M, and Swanson PC. HMGB1/2 can target DNA for illegitimate cleavage by the RAG1/2 complex. BMC Mol Biol 2009;10 24.
73.Tsuji H, et al. Rag-dependent and Rag-independent mechanisms of Notch1 rearrangement in thymic lymphomas of Atm(-/-) and scid mice. Mutat Res 2009;660(1-2) 22-32.
74.Pavlicek JW, Lyubchenko YL, and Chang Y. Quantitative analyses of RAG-RSS interactions and conformations revealed by atomic force microscopy. Biochemistry 2008;47(43) 11204-11211.
75.Kriatchko AN, Bergeron S, and Swanson PC. HMG-box domain stimulation of RAG1/2 cleavage activity is metal ion dependent. BMC Mol Biol 2008;9 32.
76.Botuyan MV, et al. Structural basis for the methylation state-specific recognition of histone H4-K20 by 53BP1 and Crb2 in DNA repair. Cell 2006;127(7) 1361-1373.
77.Drejer-Teel AH, Fugmann SD, and Schatz DG. The beyond 12/23 restriction is imposed at the nicking and pairing steps of DNA cleavage during V(D)J recombination. Mol Cell Biol 2007;27(18) 6288-6299.
78.Sanders SL, et al. Methylation of histone H4 lysine 20 controls recruitment of Crb2 to sites of DNA damage. Cell 2004;119(5) 603-614.
79.Hah YS, Lee JH, and Kim DR. DNA-dependent protein kinase mediates V(D)J recombination via RAG2 phosphorylation. J Biochem Mol Biol 2007;40(3) 432-438.
80.Kato M, et al. Omenn syndrome--review of several phenotypes of Omenn syndrome and RAG1/RAG2 mutations in Japan. Allergol Int 2006;55(2) 115-119.
81.Messier TL, et al. In vivo transposition mediated by V(D)J recombinase in human T lymphocytes. EMBO J 2003;22(6) 1381-1388.
82.Ciubotaru M, et al. RAG1-DNA binding in V(D)J recombination. Specificity and DNA-induced conformational changes revealed by fluorescence and CD spectroscopy. J Biol Chem 2003;278(8) 5584-5596.
83.Jones JM, and Gellert M. Intermediates in V(D)J recombination: a stable RAG1/2 complex sequesters cleaved RSS ends. Proc Natl Acad Sci U S A 2001;98(23) 12926-12931.
84.Harfst E, et al. Normal V(D)J recombination in cells from patients with Nijmegen breakage syndrome. Mol Immunol 2000;37(15) 915-929.
85.Lin Y, et al. Detecting S-adenosyl-L-methionine-induced conformational change of a histone methyltransferase using a homogeneous time-resolved fluorescence-based binding assay. Anal Biochem 2012;423(1) 171-177.
86.Foreman KW, et al. Structural and functional profiling of the human histone methyltransferase SMYD3. PLoS One 2011;6(7) e22290.
87.Manzur KL, et al. A dimeric viral SET domain methyltransferase specific to Lys27 of histone H3. Nat Struct Biol 2003;10(3) 187-196.
88.Min J, et al. Structure of the SET domain histone lysine methyltransferase Clr4. Nat Struct Biol 2002;9(11) 828-832.
89.Bernstein BE, et al. Methylation of histone H3 Lys 4 in coding regions of active genes. Proc Natl Acad Sci U S A 2002;99(13) 8695-8700.
90.Carrozza MJ, et al. Histone H3 methylation by Set2 directs deacetylation of coding regions by Rpd3S to suppress spurious intragenic transcription. Cell 2005;123(4) 581-592.
91.Zentner GE, Tesar PJ, and Scacheri PC. Epigenetic signatures distinguish multiple classes of enhancers with distinct cellular functions. Genome Res 2011;21(8) 1273-1283.
92.Edmunds JW, Mahadevan LC, and Clayton AL. Dynamic histone H3 methylation during gene induction: HYPB/Setd2 mediates all H3K36 trimethylation. EMBO J 2008;27(2) 406-420.
93.Bell O, et al. Localized H3K36 methylation states define histone H4K16 acetylation during transcriptional elongation in Drosophila. EMBO J 2007;26(24) 4974-4984.
94.Paull TT, et al. A critical role for histone H2AX in recruitment of repair factors to nuclear foci after DNA damage. Curr Biol 2000;10(15) 886-895.
95.Bergink S, et al. DNA damage triggers nucleotide excision repair-dependent monoubiquitylation of histone H2A. Genes Dev 2006;20(10) 1343-1352.
96.Huyen Y, et al. Methylated lysine 79 of histone H3 targets 53BP1 to DNA double-strand breaks. Nature 2004;432(7015) 406-411.
97.Budman J. and Chu G. Processing of DNA for nonhomologous end-joining by cell-free extract. EMBO J 2005;24(4) 849-860.
98.Gellert M. V(D)J recombination: RAG proteins, repair factors, and regulation. Annu Rev Biochem 2002;71 101-132.
99.Lieber MR, et al. Mechanism and regulation of human non-homologous DNA end-joining. Nat Rev Mol Cell Biol 2003;4(9) 712-720.
100.Paull TT, and Gellert M. A mechanistic basis for Mre11-directed DNA joining at microhomologies. Proc Natl Acad Sci U S A 2000;97(12) 6409-6414.
101.Lieber MR, et al. The mechanism of vertebrate nonhomologous DNA end joining and its role in V(D)J recombination. DNA Repair (Amst) 2004;3(8-9) 817-826.
102.Maser RS, et al. Mre11 complex and DNA replication: linkage to E2F and sites of DNA synthesis. Mol Cell Biol 2001;21(17) 6006-6016.
103.Mirzoeva OK, Kawaguchi T, and Pieper RO. The Mre11/Rad50/Nbs1 complex interacts with the mismatch repair system and contributes to temozolomide-induced G2 arrest and cytotoxicity. Mol Cancer Ther 2006;5(11) 2757-2766.
104.Mirzoeva OK, and Petrini JH. DNA damage-dependent nuclear dynamics of the Mre11 complex. Mol Cell Biol 2001;21(1) 281-288.
105.Paull TT, and Gellert M. Nbs1 potentiates ATP-driven DNA unwinding and endonuclease cleavage by the Mre11/Rad50 complex. Genes Dev 1999;13(10) 1276-1288.
106.Ma Y, et al. Hairpin opening and overhang processing by an Artemis/DNA-dependent protein kinase complex in nonhomologous end joining and V(D)J recombination. Cell 2002;108(6) 781-794.
107.Li J, Wei H, and Zhou MM. Structure-guided design of a methyl donor cofactor that controls a viral histone H3 lysine 27 methyltransferase activity. J Med Chem 2011;54(21) 7734-7738.
108.Wang R, et al. Formulating a fluorogenic assay to evaluate S-adenosyl-L-methionine analogues as protein methyltransferase cofactors. Mol Biosyst 2011;7(11) 2970-2981.
109.Islam K, et al. Expanding cofactor repertoire of protein lysine methyltransferase for substrate labeling. ACS Chem Biol 2011;6(7) 679-684.
110.Binda O, et al. A chemical method for labeling lysine methyltransferase substrates. Chembiochem 2011;12(2) 330-334.
111.Cao F, et al. An Ash2L/RbBP5 heterodimer stimulates the MLL1 methyltransferase activity through coordinated substrate interactions with the MLL1 SET domain. PLoS One 2010;5(11) e14102.
112.Chen NC, et al. Regulation of homocysteine metabolism and methylation in human and mouse tissues. FASEB J 2010;24(8) 2804-2817.
113.Yannone SM, et al. Werner syndrome protein is regulated and phosphorylated by DNA-dependent protein kinase. J Biol Chem 2001;276(41) 38242-38248.
114.Cooper MP, et al. Ku complex interacts with and stimulates the Werner protein. Genes Dev 2000;14(8) 907-912.
115.Hromas R, et al. The human set and transposase domain protein Metnase interacts with DNA Ligase IV and enhances the efficiency and accuracy of non-homologous end-joining. DNA Repair (Amst) 2008;7(12) 1927-1937
116.Goodwin KD, et al. Crystal structure of the human Hsmar1-derived transposase domain in the DNA repair enzyme Metnase. Biochemistry 2010;49(27) 5705-5713.
117.Jordan IK. Evolutionary tinkering with transposable elements. Proc Natl Acad Sci U S A 2006;103(21) 7941-7942.
118.Ajuh P, et al. Functional analysis of the human CDC5L complex and identification of its components by mass spectrometry. EMBO J 2000;19(23) 6569-6581.
119.Vander Kooi CW, et al. The Prp19 U-box crystal structure suggests a common dimeric architecture for a class of oligomeric E3 ubiquitin ligases. Biochemistry 2006;45(1) 121-130.
120.Taverna SD, et al. How chromatin-binding modules interpret histone modifications: lessons from professional pocket pickers. Nat Struct Mol Biol 2007;14(11) 1025-1040.
121.Lu X, and Legerski RJ. The Prp19/Pso4 core complex undergoes ubiquitylation and structural alterations in response to DNA damage. Biochem Biophys Res Commun 2007;354(4) 968-974.
122.De Haro LP, et al. Metnase promotes restart and repair of stalled and collapsed replication forks. Nucleic Acids Res 2010;38(17) 5681-5691.
123.Bryant HE, et al. PARP is activated at stalled forks to mediate Mre11-dependent replication restart and recombination. EMBO J 2009;28(17) 2601-2615.
124.Lee SH, et al. The SET domain protein Metnase mediates foreign DNA integration and links integration to nonhomologous end-joining repair. Proc Natl Acad Sci U S A 2005;102(50) 18075-18080.
125.Lundin C, et al. Different roles for nonhomologous end joining and homologous recombination following replication arrest in mammalian cells. Mol Cell Biol 2002;22(16) 5869-5878.
126.Arnaudeau C, Lundin C, and Helleday T. DNA double-strand breaks associated with replication forks are predominantly repaired by homologous recombination involving an exchange mechanism in mammalian cells. J Mol Biol 2001;307(5) 1235-1245.
127.Shrivastav M, et al. DNA-PKcs and ATM co-regulate DNA double-strand break repair. DNA Repair (Amst) 2009;8(8) 920-929.
128.Williamson EA, et al. The SET and transposase domain protein Metnase enhances chromosome decatenation: regulation by automethylation. Nucleic Acids Res 2008;36(18) 5822-5831.
129.Budzowska M, and Kanaar R. Mechanisms of dealing with DNA damage-induced replication problems. Cell Biochem Biophys 2009;53(1) 17-31.
130.McClendon AK, Rodriguez AC, and Osheroff N. Human topoisomerase IIalpha rapidly relaxes positively supercoiled DNA: implications for enzyme action ahead of replication forks. J Biol Chem 2005;280(47) 39337-39345.
131.Moll U, et al. DNA-PK, the DNA-activated protein kinase, is differentially expressed in normal and malignant human tissues. Oncogene 1999;18(20) 3114–3126.
132.Wray J, et al. Metnase mediates resistance to topoisomerase II inhibitors in breast cancer cells. PLoS One 2009;4(4) e5323.
133.Wray J, et al. Metnase mediates chromosome decatenation in acute leukemia cells. Blood 2009;114(9) 1852-1858.
134.Bryant HE, et al. Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature 2005;434(7035) 913-917.
135.Farmer H, et al. Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature 2005;434(7035) 917-921.
136.Sims RJ 3rd, Nishioka K, and Reinberg D. Histone lysine methylation: a signature for chromatin function. Trends Genet 2003;19(11) 629-639.
Written By
Hyun Suk Kim, Robert Hromas and Suk-Hee Lee
Submitted: April 24th, 2012Reviewed: September 26th, 2012Published: May 22nd, 2013
Open access peer-reviewed chapter
