Structural and functional relationships of FAN1

DNA Repair xxx (xxxx) xxx–xxx

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Review

Structural and functional relationships of FAN1 Hyeonseok Jin, Yunje Cho

⁎

Department of Life Science, Pohang University of Science and Technology, Pohang, South Korea

A R T I C L E I N F O

A B S T R A C T

Keywords: FAN1 DNA repair Interstrand cross-link Chromosome maintenance Stalled-replication fork Cancer Fanconi anemia Genome instability Kidney interstitial nephritis FANCD2 Structure-specific nuclease

FANCD2/FANCI-associated nuclease (FAN1) is a 5′ flap structure-specific endonuclease and 5′ to 3′ exonuclease. This nuclease can resolve interstrand cross-links (ICLs) independently of the Fanconi anemia (FA) pathway and controls the progression of stalled replication forks in an FA-dependent manner, thereby maintaining chromosomal stability. Several FAN1 mutations are observed in various cancers and degenerative diseases. Recently, several crystal structures of the FAN1-DNA complexes have been reported, and to date, these represent the only structures for a DNA bound ICL-repair nuclease. Puzzlingly, human FAN1 forms two different quaternary structures with different DNA binding modes, and based on these structures, two ICL-repair mechanisms have been proposed. In one mechanism, monomeric FAN1 recognizes the 5′ flap terminal phosphate via a basic pocket and successively cleaves at every third nucleotide of the DNA substrates. In the other mechanism, dimeric FAN1 scans, latches, and unwinds the postnick duplex of the substrate DNA to direct the scissile phosphodiester group to the active site. In this review, we discuss the structures, function, and proposed mechanisms of FAN1 nuclease, and provide the insights into its role in ICL repair and in processing of stalled replication forks.

1. Introduction A group of anti-cancer drugs including nitrogen mustards, mitomycin C, platinum derivatives, and psoralens can cross-link the bases from complementary strands of DNA [1,2]. These interstrand crosslinks (ICLs) are extremely toxic because they block the progression of the replication and transcription machineries. If not properly repaired, these ICLs cause replication fork collapse and the accumulation of double strand breaks (DSBs), leading to chromosomal destabilization [3,4]. To avoid the damages induced by ICLs, cells must activate a system to repair ICLs [5,6]. To date, three models for the repair of ICLs have been proposed [7–10]. Since it is likely that different ICL-inducing agents generate different local DNA duplex structures, and different assays have demonstrated particular pathways for detecting and repairing these alterations, it remains to be determined how each model reflects a different biological system [11,12]. Regardless, each system utilizes the existing cellular components reflecting in vivo activities, and it is likely that the cells employ one or more of these ICL repair systems that could work together in a coordinated manner. The first model is referred to as replication-coupled ICL repair, in which the Fanconi anemia (FA)-dependent incision of the ICL is a key

event [13–16]. This model requires four groups of proteins: structurespecific nucleases, translesion synthesis (TLS) DNA polymerases, as well as proteins in the homologous recombination (HR) and the FA pathways [17,18]. The model can be further divided depending on how forks encounter the ICL: the single fork convergence model when the fork collides at a single side of the ICL, and the dual fork convergence model when the forks are stalled on both sides of the ICL. Replicationcoupled ICL repair initiates when a single replication fork collides with an ICL. The FANCM-FAAP24-MHF complex recognizes the stalled fork structure and recruits the FA core complex, formed by eight different FA proteins, to the damaged site [19,20]. The FANCM complex activates the ataxia telangiectasia and rad3–related (ATR) kinase, which phosphorylates the FA core complex and the FANCD2-FANCI (ID2) complex. Subsequently, the FA core ubiquitinates the ID2 complex, which then recruits the structure-specific nucleases via its ubiquitin domain, allowing them to unhook the cross-link [21–23]. A nuclease complex containing SLX1, SLX4, XPF, and ERCC1 is considered the most likely nuclease responsible for this activity [24,25]. The unhooked ICL is then bypassed by TLS DNA polymerases and the replication fork is restored through strand invasion and resolution via the homologous recombination process [1,26,27]. In the dual fork convergence model,

Abbreviations: FA, FANC, Fanconi anemia; ICL, interstrand cross-link; DSB, double strand break; TLS, translesion synthesis; HR, homologous recombination; ID2, FANCD2/FANCI; AP, apurinic/apyrimidinic; UBZ, ubiquitin-binding zinc finger; Ub, ubiquitin or ubiquitinated; SAP, SAF-A/B, Acinus, and PIAS; TPR, tetratricorepeat; VRR-nuc, virus-type replication repair nuclease; WHD, winged-helix domain; HD, helical domain; MMC, mitomycin C; nd, nuclease dead; HU, Hydroxyurea; MMS, methylmethanesulfonate; APH, aphidicolin; KIN, karyomegalic interstitial nephritis; NTD, N-terminal domain; CTD, C-terminal domain; Pa, Pseudomonas aeruginosa; hFAN1M, monomeric human FAN1; hFAN1D, dimeric human FAN1; HJ, holliday junction; Pf, Pyrococcus furiosus; Hjc, holliday junction resolvase; BER, base excision repair; H2TH, helix-two turn-helix; HhH, helix-hairpin-helix ⁎ Corresponding author. E-mail address: [email protected] (Y. Cho). http://dx.doi.org/10.1016/j.dnarep.2017.06.016

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Please cite this article as: Jin, H., DNA Repair (2017), http://dx.doi.org/10.1016/j.dnarep.2017.06.016

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the two converged forks initially stall at −20 to −40 nt from the lesion owing to steric hindrance by the replicative CDC45/MCM2-7/GINS (CMG) helicases [28]. The FANCM complex binds to the DNA and initiates activation of the FA pathway. After the CMG helicase is released from the ICL, one leading strand is extended to within 1 nt of the ICL. By the time the fork reaches this point, the FA core ubiquitinates the ID2 complex and the fork is restored by unhooking, lesion bypass, and homologous recombination [29,30]. These models are more completely described in a number of excellent reviews [13,17,27,31] The second model involves a repair-independent replication process [10]. In this replication-traverse model, replication forks directly bypass the ICL without lesion repair via a mechanism mediated by the FANCM/MHF complex translocase. The FANCM-MHF complex is directly recruited to the stalled replication fork, translocates the stalled fork past the ICL, and restarts the replication on the distal side of the ICL [32]. The third model involves the DNA glycosylase NEIL3, a bifunctional enzyme that possesses both DNA glycosylase and abasic site (AP) lyase activities [33,34]. This model is specific for the ICLs induced by psoralen or abasic sites, which are formed by the attack of an isomerized aldehyde by a base from the opposite strand. In this model, the ICL is resolved by cleavage of one of the two N-glycosidic bonds by the NEIL3 glycosylase. After unhooking, the gaps are filled by TLS DNA polymerases. In this mechanism, double strand break (DSB) formation is avoided, and thus, gross chromosomal rearrangements can be minimized. When the N-glycosidic bond cleavage is blocked, the unhooking occurs via the FA-dependent incision mechanism [33,34] FAN1 was initially identified as one of the nucleases that resolve the ICL in the FA-dependent incision repair pathway [31,35–39]. Recent data, however, suggest that FAN1 unhooks ICLs independently of the FA pathway [40,41] (Fig. 1A). Although the biological function of FAN1 is still unclear, the ICL repair activity of FAN1 is important for the maintenance of chromosome stability. Furthermore, FAN1 plays a critical role in protecting the stalled replication fork in response to replication stress in an FA-dependent manner [40] (Fig. 1B). A number of FAN1-DNA complex structures have recently been reported; these are currently the only available structures for an ICL repair nuclease bound to DNA [42–44]. However, despite the availability of these reported structures, the mechanism for FAN1-mediated ICL repair remains unclear, as different structures and mechanisms have been proposed by different groups. In this review, we describe the current understanding of the structure, function, and mechanism of FAN1.

mono-ubiquitinated (Ub-) FANCD2, and thus named FAN1 [35–38,45]. Interaction between the UBZ domain and the Ub-ID2 (FANCI-FANCD2) complex has been shown to be essential for the localization of the nuclease to ICL-induced DNA damages and the UBZ-Ub-ID2 interaction appears to be essential for resistance to ICL-inducing agents. As a result, FAN1 was originally thought to function in the Fanconi anemia (FA) pathway. FAN1 is a highly conserved nuclease that consists of SAF-A/B, Acinus, and PIAS (SAP), tetratricopeptide repeat (TPR), and virus-type replication repair nuclease (VRR_NUC) domains [37,38,42–44,46]. Interestingly, bacterial and unicellular eukaryotic FAN1s are devoid of the UBZ domain [37,42,45]. The C-terminal VRR_NUC domain contains the catalytic PD-(D/E)XK motif that endows the protein with endonuclease and 5′ to 3′ exonuclease activities [35–39,47]. FAN1 recognizes the ss/dsDNA junction of 5′ flap or nicked DNA (or 5′ phosphate of the flap, see below) and makes an incision 2–4 nt into the dsDNA from the junction. The substrate specificity of FAN1 varies across different species, but the nuclease can efficiently make incisions in DNA substrates with a 5′ flap, a replication fork, duplex DNA with a nick or a gap, a splayed arm, as well as linear duplex DNA and single stranded DNA [35–39]. The most specific binding can be achieved in the presence of a 3′ flap of 4–8 nt and a 5′ flap of 1 or 2 nt or a nick to the flap structured DNA [43]. However, the nuclease can also efficiently cleave DNA molecules with a long 5′ flap [37,39,48]. Furthermore, FAN1 promotes the cleavage of a 5′ flap bound to the RPA proteins, which suggest that FAN1 can still cleave the 5′ flap DNA even in the presence of bulky adducts on ssDNA [49]. For efficient cleavage, postnick duplex with a minimal length of 10 base pairs (bp) is required [50]. FAN1 efficiently makes dual incisions on ICLs at various positions, which can vary from 2 to 6 nt away from the ss/dsDNA junction [43,50]. The diverse nuclease activity of FAN1 against various substrates implies that the nuclease can participate in several different events in DNA repair such as ICL unhooking, trimming of the unhooked oligonucleotide, and D-loop incision during homologous recombination [35,50–52]. ICL repair by FAN1 is important as cells devoid of FAN1 exhibit hypersensitivity in response to mitomycin C (MMC) [35,41,53–55], and this ICL hypersensitivity can be rescued by wildtype FAN1 but not by a mutant FAN1 lacking nuclease activity [31,40,41].

2. Biochemical features of FAN1

Although FAN1 was identified as a nuclease capable of binding to the ID2 complex in ICL repair, and, as described above, its nuclease activity is important in ICL repair, other lines of evidence reveal that FAN1 repairs ICLs independently of the FA pathway (Fig. 1A). First, analyses of FAN1-deficient patients reveal that the phenotype is dissimilar to the phenotype of FA patients lacking one of the FANC proteins, which raises the question of whether the ICL repair function of

3. FA-independent and −dependent functions of FAN1

FAN1 was discovered by four separate groups using bioinformatics, proteomics, or the genome-wide small hairpin RNA screening approach [35–39]. These analyses revealed a nuclease with an ubiquitin-binding zinc finger (UBZ) domain at its N-terminus. This nuclease is recruited to stalled replication forks via interaction between its UBZ domain and

A UBZ

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Fig. 1. The FA-independent and −dependent function of FAN1. (A) FAN1 can repair ICLs (red polygonal line) in the absence of FA components. However, it is possible that FAN1 can be recruited to the ICL via unknown cellular proteins. (B) FAN1 is recruited to a stalled replication fork by Ub-FANCD2 and controls its progression independently of the ICL.

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to five bp of postnick duplex can bind to the CTD, consistent with the biochemical data showing that 10 bp of duplex is needed for efficient binding and cleavage by hFAN1 [50]. In addition, the hFAN1M has several key features important in recognizing specific substrates. First, hFAN1M possesses a basic pocket at the bottom of the VRR_NUC domain. The pocket contains four basic residues (Arg706, Arg952, His742, and Lys986) and accommodates the terminal phosphate of a 1 nt 5′ flap (4rib.pdb) or a nick (4ria.pdb) (Fig. 3A). This interaction is critical for an optimal interaction between the hFAN1M and DNA substrates with a 5′ flap of 1 nt or a nick, as all structures of FAN1 complexed with DNA with a nick or 5′ flap of 1 nt showed binding of the 5′ terminal phosphate to this pocket. Second, the active site is located three nt away from the 5′ flap phosphate. In the active site, a Ca2+ ion that mimics the catalytic metal ion interacts with the oxygen of a phosphodiester group of the third nucleotide from the 5′ flap end. The Ca2+ ion is coordinated by Asp960, Glu815, Glu975, and Val976. Third, 5′ flap DNA binding is augmented in the presence of a 3′ flap. Two nucleotides of the 3′ flap extend away from the prenick duplex and interact with Tyr374, Val577, and Arg581 at the surface of the NTD (helical domain) via their base and ribose moieties and through their phosphate moiety. Although only two nucleotides are visible in the structure, the extension of 3′ flap by 8 nt significantly increases the affinity. The geometry of the active site and the basic pocket in hFAN1M implies that the nuclease makes successive cuts at every third nucleotide of the flap DNA (Fig. 3B). Successive incisions on the same strand of DNA would generate a gap containing ssDNA. Enzymatically, hFAN1 can make about four cuts to generate a 12 nt gap, after which the efficiency decreases significantly. Thus, the “3 nt exonuclease” mechanism may provide a basis by which FAN1 makes incisions at the sides flanking the ICLs. In principle, a 1 nt processive exonuclease mechanism could unhook an ICL. If this is the case, then why does FAN1 employ a 3 nt exonuclease mechanism? The answer may be that since an ICL distorts the local DNA structure near the lesion, a 3 nt exonuclease mechanism may be more efficient for the unhooking of the distorted ICL substrates. Data from other groups have shown that FAN1 also can process long flap DNA independent of other nucleases [35,37,39,48]. Furthermore, FAN1 can process DNA in which the flap is bound by RPA [49]. The hFAN1 structure with a 5′ flap of 2 nt (4ric.pdb) showed that the trajectory of the 5′ flap is different from those with a 1 nt flap or a nicked DNA (Fig. 3C and D). Instead of interacting with the basic residues in the pocket, the two nucleotides after the terminal phosphate extend away and bypass the basic pocket from the duplex in a direction different from those with a 1 nt flap or a nicked DNA. Although the terminal phosphate in this structure is located near Arg668, Arg672, Leu612, and Ser616 and its path is blocked, slight movement of the 5′ flap end and local structural changes in hFAN1 could allow a path for a longer 5′ flap segment or a flap with a bulky group. Although FAN1M preferably makes an incision every 3 nt, the nuclease can also cleave the phosphodiester bond between each third nt. Consistently, the crystal structures with a 5′ flap of 1 nt (either without a 3′ flap or with a 3′ flap of 8 nt, 4ri9.pdb) revealed that the first nucleotide after the 5′ terminal phosphate loops out to interact with Arg668 and Arg672, while the 5′ terminal phosphate binds to the basic pocket (Fig. 3C). Such looping shifts the register of the scissile phosphodiester groups at the active site and locates the fourth nt after the 5′ flap phosphate at the active site. In the FAN1-nicked DNA structure (4ria.pdb), the 5′ terminal phosphate of a nick also binds to the basic pocket, but the following phosphate backbone is extended with a trajectory different from that of a 5′ flap of 1 nt, locating the second nt after the 5′ terminal phosphate at the active site. These structures therefore explain an observation that while FAN1 may show the most efficient cleavage at every third nt from the 5′ phosphate of the flap, the nuclease can also make cuts at the second and fourth phosphodiester bonds from the terminal phosphate.

FAN1 is epistatic with the FA pathway [56,57]. Second, UBZ-deficient or −mutated FAN1 that is unable to interact with Ub-FANCD2 fully rescues the ICL repair defects in cells completely lacking the FAN1 protein, suggesting that recruitment of FAN1 by Ub-FANCD2 is dispensable for ICL repair [40,41]. Third, studies in the chicken DT40 cell line showed that FAN1−/−/FANCC−/− double-knockout cells are more sensitive to cisplatin than wild-type, FAN1−/−, or FANCC−/− singleknockout cells [39]. Fourth, FAN1 alone can efficiently unhook ICLs in vitro [43,50]. Also, FAN1 can be directly recruited to DNA via its SAP domain [41]. Fifth, unhooking of a cisplatin-induced ICL was shown to be unaffected in FAN1-immunodepleted Xenopus egg extracts [12]. Although FAN1-mediated ICL repair is FA-independent, studies suggest that association of FAN1 to FANCD2 via UBZ-Ub interaction is crucial for the protection of genomic stability [58,59] in response to replication stress, and this function is independent of ICL repair (Fig. 1B). In DNA fiber analyses, cells with FAN1nd/nd exhibited longer replication tracks in cells treated with the fork-stalling agent hydroxyurea (HU) or with methylmethanesulfonate (MMS), which can be restored to normal by the wild-type FAN1 but not by the FAN1 mutant with non-functional UBZ or with impaired nuclease activity. This result demonstrates that both FAN1 nuclease activity and UBZ-dependent binding to FANCD2 are required for normal progression of a stalled fork. However, the molecular mechanism by which FAN1 controls processing of the stalled forks is unclear. FAN1 also can be recruited to chromatin via the FANCD2-BLM complex independent of the UBZ domain and promote the restart of aphidicolin (APH)-stalled replication forks, and ultimately suppress firing of new origins [60]. These data suggest that FAN1 plays an important role in controlling the restart of the stalled replication fork. Presumably because of its role in maintaining chromosomal integrity, mutation or deficiency of FAN1 could be associated with various degenerative diseases and cancers [15,26,57,61–63] (see below). 4. Structure and mechanism of FAN1 4.1. FAN1 monomer − DNA: 3 nucleotide exonuclease model In all reported structures of the FAN1–DNA complexes [42–44], FAN1 lacking the UBZ domain and 5′ flap DNA were used for structure determination. With the exception of one bacterial homolog (Pseudomonas aeruginosa FAN1, PaFAN1), all the reported FAN1 structures were determined using human FAN1. The structures are largely divided into two groups. One group of structures has a FAN1 monomer-DNA complex, and the other group has a FAN1 dimer-DNA complex [43,44] (Fig. 2A–C). Although the overall structures of the FAN1 monomer are very similar in both groups, the quaternary structures, DNA recognition modes, and substrate cleavage mechanisms are markedly different between the monomeric and dimeric forms. Human FAN1 monomer (hFAN1M, 4ri8.pdb) forms a bi-lobed structure in which the N-terminal domain (NTD) and the C-terminal domain (CTD) are positioned nearly orthogonal to one another (Fig. 2A). From the junction between the NTD and the CTD, a helical domain, a winged helix (WH) domain, and the SAP domain are sequentially ordered to form the NTD. The CTD comprises the TPR domain on one side and the VRR domain on the other side. The NTD recognizes the prenick duplex and 3′ flap, whereas the CTD interacts with the postnick duplex and the 5′ flap. These interactions provide the primary feature for the substrate recognition by hFAN1M. All domains are extensively involved in binding to DNA. In the described structure, the DNA forms a V-shaped structure with a 70° kink at the junction between the pre- and postnick duplexes. The kink is achieved through the use of a hydrophobic, helical wedge as has been observed in other flap structure-specific nucleases [64–66]. The NTD makes contact with nine nucleotides through its continuous surface, whereas the CTD interacts with five nucleotides via a deep groove. However, judging from the PaFAN1-DNA structure, an additional four 3

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Fig. 2. Overall structure of FAN1 monomer and dimer. (A) hFAN1 consists of NTD and CTD domains, which recognize the pre- and postnick duplexes of a flap DNA, respectively (PDB ID: d4ri8). Each omain is labelled and colored as shown in 2D. (B) Bacterial FAN1 (PaFAN1) interacts with DNA in a mode similar to that of hFAN1 (4r89.pdb). (C) FAN1 dimer interacts with DNA via NTD and CTD of first FAN1 (bottom molecule) and NTD of second FAN1 (top molecule, yellow; 4rea.pdb). The figure shown here represents the “substrate-unwinding” state of hFAN1 dimer. The metal ion in the active site is represented as a blue sphere or an asterisk (for the expected metal ion). (D) Domain architecture of FAN1. The N-terminal UBZ domain is omitted.

DNA in the presence and absence of the metal ion is different. Aligning the structures of the NTDs of PaFAN1 in the absence and presence of metal ions, the CTD shifts by 10° and shifts the scissile phosphate toward the active site. This conformational change may provide a basis for the processive exonuclease activity by PaFAN1 (Fig. 3F).

4.2. Comparison with bacterial FAN1 The bacterial FAN1 homolog, PaFAN1 also interacts with the 5′ flap DNA as a monomer [42] (4r8a.pdb). Although the angle between the NTD and CTD of PaFAN1 slightly differs from that in hFANM, the overall structure of bacterial FAN1 is very similar (Fig. 2B). Recognition of the pre- and postnick duplex by the NTD and CTD, respectively, and DNA kinking at the nicked junction by the helical wedge are also observed in the PaFAN1 structure, suggesting the highly conserved nature of this nuclease across the species. Despite these overall similarities, there are, however, several notable differences between the structures. First, the basic pocket in hFAN1M is not conserved in PaFAN1. No basic charged residues are present at, or near, the region equivalent to the basic pocket of hFAN1 (Fig. 3E). Instead, a different pocket is formed at the junction of the VRR_NUC, the TPR, and the helical domains. The 5′ flap ssDNA and the junction between the prenick and postnick segments are accommodated by this pocket. In the pocket, the 5′ flap ssDNA in PaFAN1 initially points to the pocket and then turns back towards the outside of the pocket, adopting a U shaped structure. This conformation explains the ability of PaFAN1 to cleave long 5′ flap strands, as the long 5′ flap ssDNA extends away from the surface of PaFAN1. Presumably due to the lack of the basic pocket, bacterial FAN processively incises one nucleotide after an initial endonucleolytic cut (Fig. 3F). The second major difference is that the 3′ flap binding residues in hFAN1M are not conserved in PaFAN1, in which the longer 3′ flap can extend away from the duplex without interference. However, further studies will be needed to determine whether a 3′ flap augments the binding of a substrate to PaFAN1. A third difference is that the catalytic metal ion is located around the fourth and fifth phosphates downstream from the junction between the prenick and postnick duplexes in PaFAN1 (4r89.pdb), whereas hFAN1 makes a cut at third nt from the junction (Fig. 3B and F). The fifth phosphate in PaFAN1 is bound by two Mn2+ ions and Lys524. The first metal is coordinated with Asp507, Glu522, and Val523 (main chain oxygen) and the phosphate oxygen, while the second metal is coordinated with Glu386, Asp507, and the phosphate oxygen. In the PaFAN1, the incision site is determined by the position of the 3′ end of the prenick segment. One notable feature of PaFAN1 is that the conformation of FAN1-

4.3. FAN1 dimer − DNA: substrate-scanning and −unwinding model A critical observation is that hFAN1 assembles as a dimer in the presence of DNA [44]. PaFAN1 lacks a basic motif in helix α9, which is crucial for the formation of a dimer in hFAN1. Consistent with this observation, Xiong’s group reported three different dimeric FAN1-DNA structures (2.2–4.2 Å resolution) that were determined in the presence of a 5′ flap DNA of one nt. In the dimeric structure (hFAN1D), FAN1 forms a ‘head to tail’ dimer through the TPR and VRR domains of the first FAN1 molecule interacting with the SAP domain of second FAN1 (Fig. 2C). The first FAN1 molecule appears to play a primary role in catalysis by engaging the DNA in its active site and the second FAN1 molecule plays an auxiliary role, facilitating substrate orientation and flap unwinding. Each of the reported FAN1D structures is proposed to represent a different state of FAN1D: namely the ‘substrate-scanning (4rec.pdb)’, ‘substrate-latching (4reb.pdb)’, and ‘substrate-unwinding (4rea.pdb)’ forms. In each structure, DNA binds to the FAN1 dimer in a different manner. Although whether dimeric FAN1 interacts with DNA with higher affinity than the FAN1 monomer is unclear, both pre- and postnick DNA bind tightly to all domains except the TPR domain in the ‘substrate-unwinding’ form. In the “substrate-scanning” form, the primary FAN1 recognizes 10 nt of prenick duplex and 3 nt of postnick duplex via its NTD. The DNA-binding surface in the NTD of the dimer is very similar to that in the NTD of hFAN1M, as the entire prenick and part of the postnick DNA interact with the continuous surface formed by the helical domain, the winged-helix, and the SAP domain. Although only one FAN1 molecule has been proposed to interact with DNA in the ‘substrate-scanning model’, the crystal structure shows that the asymmetric unit contains one FAN1-DNA complex, indicating that a FAN1 dimer interacts with two DNA molecules. It is possible that each FAN1 molecule initially binds to a DNA molecule and one of the proteins is released after the scanning process. In the ‘substrate-latching’ form, nine nt of the postnick duplex contacts the NTD in the primary FAN1, 4

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A

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Fig. 3. The ICL repair mechanism of FAN1. The basic pocket of hFAN1M with 3 different substrates, (A) 5′ flap of 1 nt and 3′ flap of 4 nt (4ri8.pdb); (C) 5′ flap of 1 nt and 3′ flap of 8 nt (4ri9.pdb); (D) 5′ flap of 2 nt (4ric.pdbb); the DNA schematics are shown below each structure. Phosphate ions in DNA backbone, Ca2+ ions, Ba2+ ions, and Mn2+ ions are represented as spheres (red, dark pink, light pink, and yellow, respectively). (B) The “3 nt exonuclease” mechanism by FAN1M. Binding of the 5′ terminal phosphate of a flap or a nick to the basic pocket of FAN1M locates the scissile phosphate group of every third nt after the terminal phosphate at the active site. Terminal phosphates, Ca2+ ions, and disordered second metal ions are represented as filled circles (red and green) and empty circles, respectively. The number of the phosphate groups was counted from the junction. (E-F) PaFAN1 (4r89.pdb) possesses a pocket dissimilar to the basic pocket (flap binding pocket). PaFAN1 Exhibits 1 nt processive exonuclease activity. (G) hFAN1D processes the flap or nicked DNA via substrate scanning (4rec.pdb), latching (4reb.pdb), and unwinding (4rea.pdb) processes.

(Fig. 3G). In three FAN1D-DNA structures, the auxiliary FAN1 is rotated by 10–15° relative to the primary FAN1 through a conformational change of helices α7 and α5β1. However, differences in the resolution of the structures limits accurate information about their movement. It has been proposed that the Mre11 dimer melts DNA via rigid body rotation of its capping domain, or of each Mre11 subunit [67–69]. The dimeric structure of hFAN1 provides a plausible mechanism for the cleavage of long 5′ flap DNA substrates as well as those DNA substrates with the 5′ flap bound with bulky adducts such as RPA.

whereas one nt of the prenick duplex contacts the helical domain of the NTD in the auxiliary FAN1. In this state, the positions of the pre- and postnick segments were reversed from those in the “substrate-scanning” state, which suggests that FAN1 may scan through the entire DNA. In the ‘substrate-unwinding’ form, four nt of postnick duplex is unwound and is directed to the primary FAN1 active site. Although the distance between the active site residues and the 5′ terminal phosphate of the flap is more than 11 Å away, the structure suggests that DNA may be melted by FAN1 and the resulting ssDNA directed to the active site 5

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However, even in the “substrate-unwinding” state, the 5′ terminal phosphate of the flap is located far away from the active site. Thus, it would be difficult to explain the endonucleolytic incision of a DNA with a short 5′ flap. Furthermore, the ‘substrate-scanning and −unwinding’ model has a limitation in explaining unhooking of the ICL near the ss/ dsDNA junction. For instance, ICL induced by psoralen will block replication fork progression 2 nt ahead of an ICL. In addition to a substantial conformational change of the structure of postnick DNA, the primary and/or auxiliary FAN1 must undergo gross structural changes to locate the scissile phosphate group of an ICL near the junction at the active site.

duplexes is achieved by the separate motifs or domains. However, FAN1M does not undergo any noticeable conformational changes upon binding DNA and accommodates the DNA to its preformed binding site [43,46], whereas FEN1 undergoes a substrate-induced disorder-toorder transition for the recognition of 3′ and 5′ flaps. Such structural transition creates a deep binding pocket for the 3′ flap, which in turn triggers the formation of the active site and helical gateway, where the 5′ flap binds and passes through. The active site, together with the K+/ H2TH (helix two turn helix) motif, which is one helical turn apart from the active site, provides a platform for the binding of the postnick duplex [66]. MUS81-EME1 is a 3′ nuclease that resolves the nicked HJ in homologous recombination processes. MUS81-EME1 (4p0r.pdb) uses two repeats of helix-hairpin-helix (2(HhH)2) domains to separately recognize pre- and postnick duplexes [64] (Fig. 4C). While the postnick segment is recognized by the 2(HTH)2 domain of MUS81 and the nuclease-like domain of EME1, the prenick duplex is bound by the 2(HTH)2 domain of EME1 and the nuclease domain of MUS81. MUS81EME1 initially recognizes the 5′ flap, which triggers the disorder-toorder transition for the formation of the 3′ flap binding surface. DNA binding by the FAN1 dimer is similar to that of the Mre11 nuclease (4tug.pdb and 3dsd.pdb), which functions as a homodimer (Fig. 4D and E). The Mre11 nuclease possesses endonuclease activity and 3′ to 5′ exonuclease activity [69,73,74]. Although there is no structural similarity between the DNA binding motif of Mre11 and that of FAN1, both nucleases interact with DNA via backbone phosphate interactions. At present, the exact mechanism by which Mre11 achieves endonucleolytic cleavage is unclear since no structure shows that the 5′ scissile phosphate is located at the active site. ATP hydrolysis-induced disengagement of the Rad50 dimer might melt the duplex DNA to allow its access to the active site [68]. For the exonuclease mechanism, it has been proposed that either rotation of the capping domain or a subunit alters the position of the “recognition loop” to wedge the bases in the minor groove in the DNA melting process [67–69]. It remains to be solved how FAN1 scans the substrate to select the incision point.

5. Evolutionary relationship between FAN1 and junction resolvase Similarities in the structure and substrate specificities between the PaFAN1 and hFAN1 imply that FAN1 was probably present in early stages of evolution [42–44,46]. The VRR_NUC domain of FAN1 forms an α/β fold with its central five-stranded sheet flanked by a helix and a bundle of helices on each side [70]. This domain is most similar to the Holliday junction (HJ) cleavage enzyme of Pyrococcus furiosus (PfHjc, 1gef.pdb), that of type II restriction enzymes, and the single-domain VRR_NUC proteins from phage or bacteria that form a dimer (4qbo.pdb and 5fdk.pdb) and possess HJ resolvase activities [48,71,72]. The VRR_NUC domain of FAN1 contains a six-helix bundle inserted between the second and third strand of the β sheet, at the equivalent position of an opposing molecule in the dimeric HJ resolvase (PfHjc). The insertion of the helix bundle prevents the FAN1 dimer formation through the VRR_NUC domain. This unique feature of the VRR_NUC domain in FAN1endows it with strong 5′ flap nuclease activity but prevents it from resolving intact HJs [48]. We speculate that the dimeric Hjc enzyme may have evolved to disassemble into monomers through acquisition of helical insertions. This modification may then have altered the substrate specificity of the nuclease. The VRR_NUC domain of FAN1 has a dual role: 5′ flap recognition by the six-helix bundle and active site formation by the central α/β catalytic subdomain. The 5′ flap binding site is not conserved between PaFAN1 and hFAN1. Addition of the TPR domain to the VRR_NUC domain forms a groove in which the postnick duplex can fit. FAN1 can then recognize and incise the linear duplex DNA [35,37]. In addition, FAN1 traverses the ICL in linear duplex DNA [50]. Because linear duplex DNA cannot make a sharp kink as the 5′ flap DNA does, it is likely that linear duplex only interacts with the CTD. Thus, we speculate that the CTD may be an earlier form of FAN1, which acquired the NTD at a later stage to interact with flapped substrates.

7. FAN1 and disease Deficiency of FAN1 in humans has recently been shown to cause the rare hereditary kidney disease karyomegalic interstitial nephritis (KIN), characterized by chronic renal fibrosis, tubular degeneration, and polyploidy in multiple tissues [57]. FAN1-deficient mice also exhibited polyploidy and karyomegaly in the kidney and liver with age [40,41]. Mutations in FAN1 are also detected in patients suffering from various degenerative and neurological diseases including schizophrenia or autism. [61,63]. Presumably, chromosomal abnormalities caused by a failure of replication fork protection may underscore some of these diseases. Disease-associated FAN1 mutations can be mapped onto the FAN1-DNA structures, providing a framework to understand the basis for FAN1-linked diseases at the molecular level (Fig. 5). Overall, the mutations can be classified into three groups. The first group of mutations is clustered near the active site and expected to abolish the nuclease activity of FAN1. These mutations include missense mutations K794R and D960N, and truncation mutations R536*, R679Tfs*5 (a frameshift that stops at the 5th residue after Arg679), W707*, R710*, R749*, D873Tfs*17, L925Pfs*25, and R952* and were found in patients with KIN, schizophrenia, autism, pancreatic or colorectal cancer, indicating that the nuclease activity of FAN1 appears to be linked with all FAN1 associated-diseases. A second group of mutations is distributed over all the domains and may contribute to alterations in structural stability and the DNA interaction activity. Most of the missense mutations, R377W, M393V or M393T, L395P, R591W, R658Q, and C871R are located in the interface between the helical domain and CTD, strongly suggesting that these residues are involved in the correct conformational integrity of NTD and/or CTD. Mutations including M50R, E437G, R507H, P894S, Q929P, and G937D are classified into

6. Comparison between FAN1 and other structure-specific nucleases Because hFAN1 interacts with DNA as a monomer and a dimer, each mode of DNA binding by hFAN1 resembles different structure-specific nucleases. In the “3 nt exonuclease” mechanism, FAN1M primarily recognizes the substrate via separate NTD and CTD domains (Fig. 4A). Additional local interactions of the 3′ and 5′ flap by FAN1 further specify the binding of the DNA substrate for optimal cleavage. Recognition of pre- and postnick flap substrates by a discontinuous surface is also observed in other 5′ nucleases including flap endonuclease 1 (FEN1, 3q8l.pdb) and exonuclease 1 (Exo1, 3qea.pdb). (Fig. 4B). FEN1 recognizes 5′ flap structures formed during Okazaki fragment processing or during long patch base excision repair (BER), and incises 1 nt into dsDNA from the junction of a short 5′ flap. All these nucleases kink their substrates at the junction between pre- and postnick segments via the hydrophobic helical wedge, which facilitates the binding and positioning of the scissile phosphate at the active site [65,66]. Furthermore, although these nucleases interact with the substrate through different DNA binding domains, the recognition of pre- and postnick 6

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A

B

M

hFAN1, 4ri8.pdb Wedge

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hMUS81-EME1, 4p0r.pdb Wedge

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N

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C 2HhH2 C

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E

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hFAN1, 4rec.pdb

MjMre11, 4tug.pdb

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Prenick

Fig. 4. Structural comparison between FAN1 and other structural-specific nucleases. (A–C) The DNA binding mode of hFAN1M is similar to those of other 5′ or 3′ nucleases. Pre- and postnick duplexes are recognized by independent surfaces, and the DNA bends at the nick by helical wedge (magenta). The metal ion is shown as a red sphere. (A) Structure of the FAN1 monomer (4ri8.pdb). The NTD is shown in gray and the CTD is shown in white. (B) hFEN1 (3q8l.pdb). (C) hMUS81-EME1 (4p0r.pdb). The nuclease, nuclease-like, and the two HhH2 domains are labeled. The expected metal ion is marked using an asterisk. (D, E) Comparison of the structures between hFAN1D and archaeal Mre11 dimer. (D) hFAN1 dimer (4rec.pdb). Only the primary FAN1 (gray) interacts with the DNA in this scanning mode. Subunit rotation may facilitate the melting of DNA. The expected metal ion is marked using an asterisk. (E) Methanococcus janaschii (Mj) Mre11 dimer (4tug.pdb). Domain rotation or subunit of the Mre11 dimer is expected to melt DNA by the wedge loop (magenta). The metal ions are shown as red spheres.

*

TPR

on the roles of the FAN1 nuclease in both FA-dependent and −independent pathways. FAN1 can unhook ICL in the absence of other factors. FAN1 can also be recruited to FANCD2 via a UBZ-Ub interaction and controls the processing of the stalled replication fork independent of ICL repair. The role of FAN1 in ICL repair and replication fork progression, and its association with various diseases illustrate the significance of FAN1 in the maintenance of chromosomal integrity. One of the outstanding questions that remain to be answered is that of how FAN1 protects the stalled forks from replication stress. Because FAN1 nuclease activity is required for the restart of stalled forks induced by specific agents and prevents new origin firing, it is possible that FAN1 acts as a safeguard that controls replication re-initiation during specific replication stresses. In the FA pathway, the ID2 complex is believed to recruit the nuclease complex containing SLX1-SLX4-XPF for ICL repair, whereas the ID2-FAN1 binding mediated via UBZ-Ub interaction is also important for stalled replication fork progression. It is unlikely that the ID2 complex recruits these nucleases at the same time. The question then is to determine the signals by which the ID2 complex selects these nucleases. Both the SLX1-SLX4-XPF complex and FAN1 can make dual incisions on locally distorted DNA, such as cross-linked DNA. However, the size of the SLX1/SLX4/XPF complex and the range and structure of the damages repaired by this nuclease complex are likely to be different from those by FAN1. While the DNA structures on which FAN1 operates in cells are not yet clear, its ability to process ICLs as well as a wide variety of substrates with different structures suggests that the nuclease can make incisions on forks stalled on severely distorted DNA strands by specific agents. It is also possible that FAN1 functions together with other nucleases in resolving stalled forks. FAN1 can interact with FANCD2 independent of the UBZ domain and protects the APH-stalled replication forks together with Mre11. If the FAN1 nuclease activity is too low, the replication fork is stalled, and if the activity is too high, the nuclease causes fork degradation and initiates origin firing. Thus, failure of regulation of the FAN1 nuclease activity may lead to genomic instability. Further studies are required to explain how interactions

Postnick Prenick R507H

R679Tfs*5

R749* R710* W707* K794R

VRR-NUC

SAP

D960N R952* R658Q

D873Tfs*17 C871R G937D

P654L

R536*

L395P

M393V or M393T L925Pfs*25

WHD

R591W P894S

E437G Q929P

R377W

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Fig. 5. Structural mapping of the FAN1-assocaited disease mutation. FAN1-associated mutations involved in autism and schizophrenia (dark pink), KIN (yellow), and pancreatic or colorectal cancers (cyan) are mapped onto the hFAN1M structure (4ri8.pdb). Missense mutations are shown as spheres and truncation mutations are shown as empty triangles.

the last group and most of them are exposed to the surface of the protein, except for M50R. These mutations are likely to be important for protein–protein interactions. The E437G mutation is located in the dimeric interface of the ‘substrate-unwinding’ model of hFAN1D and possibly affects the domain rotation of FAN1 during the substrate unwinding process. In general, the KIN-associated mutations are clustered in the CTD domain. Also, FAN1 mutations observed in pancreatic and colorectal cancers affect either the UBZ–Ub interaction or nuclease activity, which suggest that the FA-dependent control of a stalled fork progression by FAN1 in response to replication stress might be associated with development of specific types of cancers.

8. Outlooks Structural and functional analyses of FAN1 have provided insights 7

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(2013) a012732. [6] J.P. Duxin, J.C. Walter, What is the DNA repair defect underlying Fanconi anemia? Curr. Opin. Cell Biol. 37 (2015) 49–60. [7] M. Raschle, P. Knipscheer, M. Enoiu, T. Angelov, J. Sun, J.D. Griffith, T.E. Ellenberger, O.D. Scharer, J.C. Walter, Mechanism of replication-coupled DNA interstrand crosslink repair, Cell 134 (2008) 969–980. [8] J. Zhang, J.M. Dewar, M. Budzowska, A. Motnenko, M.A. Cohn, J.C. Walter, DNA interstrand cross-link repair requires replication-fork convergence, Nat. Struct. Mol. Biol. 22 (2015) 242–247. [9] W. Wang, Emergence of a DNA-damage response network consisting of Fanconi anaemia and BRCA proteins, Nat. Rev. Genet. 8 (2007) 735–748. [10] J. Huang, S. Liu, M.A. Bellani, A.K. Thazhathveetil, C. Ling, J.P. de Winter, Y. Wang, W. Wang, M.M. Seidman, The DNA translocase FANCM/MHF promotes replication traverse of DNA interstrand crosslinks, Mol. Cell 52 (2013) 434–446. [11] M. Ben-Yehoyada, L.C. Wang, I.D. Kozekov, C.J. Rizzo, M.E. Gottesman, J. Gautier, Checkpoint signaling from a single DNA interstrand crosslink, Mol. Cell 35 (2009) 704–715. [12] D. Klein Douwel, R.A. Boonen, D.T. Long, A.A. Szypowska, M. Raschle, J.C. Walter, P. Knipscheer, XPF-ERCC1 acts in Unhooking DNA interstrand crosslinks in cooperation with FANCD2 and FANCP/SLX4, Mol. Cell 54 (2014) 460–471. [13] Y. Kee, A.D. D'Andrea, Expanded roles of the Fanconi anemia pathway in preserving genomic stability, Genes Dev. 24 (2010) 1680–1694. [14] M.C. Kottemann, A. Smogorzewska, Fanconi anaemia and the repair of Watson and Crick DNA crosslinks, Nature 493 (2013) 356–363. [15] K.A. Rickman, F.P. Lach, A. Abhyankar, F.X. Donovan, E.M. Sanborn, J.A. Kennedy, C. Sougnez, S.B. Gabriel, O. Elemento, S.C. Chandrasekharappa, D. Schindler, A.D. Auerbach, A. Smogorzewska, Deficiency of UBE2T, the E2 ubiquitin ligase necessary for FANCD2 and FANCI ubiquitination causes FA-T subtype of fanconi anemia, Cell Rep. 12 (2015) 35–41. [16] L.H. Thompson, J.M. Hinz, Cellular and molecular consequences of defective Fanconi anemia proteins in replication-coupled DNA repair: mechanistic insights, Mutat. Res. 668 (2009) 54–72. [17] H. Kim, A.D. D’Andrea, Regulation of DNA cross-link repair by the Fanconi anemia/ BRCA pathway, Genes Dev. 26 (2012) 1393–1408. [18] K.D. Mirchandani, A.D. D'Andrea, The Fanconi anemia/BRCA pathway: a coordinator of cross-link repair, Exp. Cell Res. 312 (2006) 2647–2653. [19] A. Ciccia, C. Ling, R. Coulthard, Z. Yan, Y. Xue, A.R. Meetei, H. Laghmani el, H. Joenje, N. McDonald, J.P. de Winter, W. Wang, S.C. West, Identification of FAAP24, a Fanconi anemia core complex protein that interacts with FANCM, Mol. Cell 25 (2007) 331–343. [20] M. Huang, J.M. Kim, B. Shiotani, K. Yang, L. Zou, A.D. D’Andrea, The FANCM/ FAAP24 complex is required for the DNA inter-strand crosslink-induced checkpoint response, Mol. Cell 39 (2010) 259–268. [21] R.A. Schwab, A.N. Blackford, W. Niedzwiedz, ATR activation and replication fork restart are defective in FANCM-deficient cells, The EMBO Journal 29 (2010) 806–818. [22] A. Smogorzewska, S. Matsuoka, P. Vinciguerra, E.R. McDonald 3rd, K.E. Hurov, J. Luo, B.A. Ballif, S.P. Gygi, K. Hofmann, A.D. D’Andrea, S.J. Elledge, Identification of the FANCI protein, a monoubiquitinated FANCD2 paralog required for DNA repair, Cell 129 (2007) 289–301. [23] S.J. Collis, A. Ciccia, A.J. Deans, Z. Horejsi, J.S. Martin, S.L. Maslen, J.M. Skehel, S.J. Elledge, S.C. West, S.J. Boulton, FANCM and FAAP24 function in ATR-mediated checkpoint signaling independently of the Fanconi anemia core complex, Mol. Cell 32 (2008) 313–324. [24] M.R. Hodskinson, J. Silhan, G.P. Crossan, J.I. Garaycoechea, S. Mukherjee, C.M. Johnson, O.D. Scharer, K.J. Patel, Mouse SLX4 is a tumor suppressor that stimulates the activity of the nuclease XPF-ERCC1 in DNA crosslink repair, Mol. Cell 54 (2014) 472–484. [25] N. Bhagwat, A.L. Olsen, A.T. Wang, K. Hanada, P. Stuckert, R. Kanaar, A. D'Andrea, L.J. Niedernhofer, P.J. McHugh, XPF-ERCC1 participates in the Fanconi anemia pathway of cross-link repair, Mol. Cell. Biol. 29 (2009) 6427–6437. [26] A.T. Wang, T. Kim, J.E. Wagner, B.A. Conti, F.P. Lach, A.L. Huang, H. Molina, E.M. Sanborn, H. Zierhut, B.K. Cornes, A. Abhyankar, C. Sougnez, S.B. Gabriel, A.D. Auerbach, S.C. Kowalczykowski, A. Smogorzewska, A dominant mutation in human RAD51 reveals its function in DNA interstrand crosslink repair independent of homologous recombination, Mol. Cell 59 (2015) 478–490. [27] J. Zhang, J.C. Walter, Mechanism and regulation of incisions during DNA interstrand cross-link repair, DNA Repair (Amst) 19 (2014) 135–142. [28] Y.V. Fu, H. Yardimci, D.T. Long, T.V. Ho, A. Guainazzi, V.P. Bermudez, J. Hurwitz, A. van Oijen, O.D. Schärer, J.C. Walter, Selective bypass of a lagging strand roadblock by the eukaryotic replicative DNA helicase, Cell 146 (2011) 931–941. [29] P. Knipscheer, M. Raschle, A. Smogorzewska, M. Enoiu, T.V. Ho, O.D. Scharer, S.J. Elledge, J.C. Walter, The Fanconi anemia pathway promotes replication-dependent DNA interstrand cross-link repair, Science 326 (2009) 1698–1701. [30] D.T. Long, M. Raschle, V. Joukov, J.C. Walter, Mechanism of RAD51-dependent DNA interstrand cross-link repair, Science 333 (2011) 84–87. [31] M. Huang, A.D. D'Andrea, A new nuclease member of the FAN club, Nat. Struct. Mol. Biol. 17 (2010) 926–928. [32] A.R. Meetei, A.L. Medhurst, C. Ling, Y. Xue, T.R. Singh, P. Bier, J. Steltenpool, S. Stone, I. Dokal, C.G. Mathew, M. Hoatlin, H. Joenje, J.P. de Winter, W. Wang, A human ortholog of archaeal DNA repair protein Hef is defective in Fanconi anemia complementation group M, Nat. Genet. 37 (2005) 958–963. [33] V. Rolseth, S.Z. Krokeide, D. Kunke, C.G. Neurauter, R. Suganthan, Y. Sejersted, G.A. Hildrestrand, M. Bjørås, L. Luna, Loss of Neil3 the major DNA glycosylase activity for removal of hydantoins in single stranded DNA, reduces cellular proliferation and sensitizes cells to genotoxic stress, Biochimica et Biophysica Acta

between FAN1 and other cellular proteins regulate the function of FAN1. Puzzlingly, there are two different structures proposed for FAN1, each of which is supported by biochemical analyses. Although we cannot rule out either of the proposed structures and associated mechanisms, each model has the advantage of being able to explain the processing of different substrates. While the “3 nt exonuclease” mechanism is suitable for short flaps with one or two nt or a nick, the “substrate scanning and unwinding” mechanism provides a persuasive explanation for incision of long flapped DNA. In fact, the quaternary structure of FAN1 may change in the presence of different DNA structures. For instance, the FAN1 monomer interacts with specific DNA structures with short 3′ and 5′ flaps, and then the nuclease may assemble into a dimer to process DNA with long 5′ flaps or with 5′ flaps where the bulky adducts or proteins are bound. However, it is interesting that the bacterial FAN1 homolog functions as a monomer and incises most of the same substrates. There are several examples where the enzyme activities can be regulated by alteration of oligomeric states [75]. The paradox for the “3 nt exonuclease” and “substrate scanning and unwinding” mechanisms can be solved by multidisciplinary approaches including single-molecule FRET, biochemical, and biophysical analyses. FAN1 nuclease is an attractive target for designing anti-cancer drugs. Recently, two studies showed that FANCD2 is essential in fork protection by regulating fork arrest and restart in BRCA1 or BRCA2 deficient cells [76–78]. Cancer cells resulting from BRCA1 or BRCA2 mutation cannot tolerate loss of fancd2, exhibiting chromosomal instability and increased cell death. BRCA1 and BRCA2 are critical proteins that cooperate the Fanconi Anemia and BRCA pathways [79]. Because FAN1 is a downstream nuclease of FANCD2, a synthetic lethality approach to target BRCA1 or BRCA2-deficient cancers by inhibiting FAN1 activity may selectively kill tumor cells lacking BRCA1 and BRCA2 [76]. Targeting the basic pocket where the 5′ phosphate of the flap binds or the dimerization interface may be an efficient way to design small molecules that abrogate FAN1. Many qualitative studies have provided important information about FAN1 in a relatively short time since its first discovery in 2010. Nevertheless, there are many questions about FAN1 that remain to be resolved, in addition to its structure and mechanism, its regulation, the interaction network with other proteins, its function in the replication process and in other DNA repair pathways, and identification of other FAN1-linked diseases and their basis. Additional studies using integrated approaches will provide clues on these questions. Conflict of interest The authors declare no conflict of interest. Acknowledgements We thank Youngran Kim for helpful discussions and preparation of figures. This work was supported by grants from the National Research Foundation of Korea (NRF) funded by the Korea government (MEST, No. 2015R1A2A1A05001694 and NRF-2013M3A6A4044580) and BK21 program (Ministry of Education). References [1] A.J. Deans, S.C. West, DNA interstrand crosslink repair and cancer, Nat. Rev. Cancer 11 (2011) 467–480. [2] Y. Huang, L. Li, DNA crosslinking damage and cancer – a tale of friend and foe, Transl. Cancer Res. 2 (2013) 144–154. [3] D.M. Noll, T.M. Mason, P.S. Miller, Formation and repair of interstrand cross-links in DNA, Chem. Rev. 106 (2006) 277–301. [4] K.M. McCabe, S.B. Olson, R.E. Moses, DNA interstrand crosslink repair in mammalian cells, J. Cell. Physiol. 220 (2009) 569–573. [5] C. Clauson, O.D. Scharer, L. Niedernhofer, Advances in understanding the complex mechanisms of DNA interstrand cross-link repair, Cold Spring Harb. Perspect Biol. 5

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H. Jin, Y. Cho (BBA) − Mol. Cell Res. 1833 (2013) 1157–1164. [34] S. Wang, K. Liu, L. Xiao, L. Yang, H. Li, F. Zhang, L. Lei, S. Li, X. Feng, A. Li, J. He, Characterization of a novel DNA glycosylase from S. sahachiroi involved in the reduction and repair of azinomycin B induced DNA damage, Nucleic Acids Res. 44 (2016) 187–197. [35] K. Kratz, B. Schopf, S. Kaden, A. Sendoel, R. Eberhard, C. Lademann, E. Cannavo, A.A. Sartori, M.O. Hengartner, J. Jiricny, Deficiency of FANCD2-associated nuclease KIAA1018/FAN1 sensitizes cells to interstrand crosslinking agents, Cell 142 (2010) 77–88. [36] T. Liu, G. Ghosal, J. Yuan, J. Chen, J. Huang, FAN1 acts with FANCI-FANCD2 to promote DNA interstrand cross-link repair, Science 329 (2010) 693–696. [37] C. MacKay, A.C. Declais, C. Lundin, A. Agostinho, A.J. Deans, T.J. MacArtney, K. Hofmann, A. Gartner, S.C. West, T. Helleday, D.M. Lilley, J. Rouse, Identification of KIAA1018/FAN1, a DNA repair nuclease recruited to DNA damage by monoubiquitinated FANCD2, Cell 142 (2010) 65–76. [38] A. Smogorzewska, R. Desetty, T.T. Saito, M. Schlabach, F.P. Lach, M.E. Sowa, A.B. Clark, T.A. Kunkel, J.W. Harper, M.P. Colaiacovo, S.J. Elledge, A genetic screen identifies FAN1, a Fanconi anemia-associated nuclease necessary for DNA interstrand crosslink repair, Mol. Cell 39 (2010) 36–47. [39] K. Yoshikiyo, K. Kratz, K. Hirota, K. Nishihara, M. Takata, H. Kurumizaka, S. Horimoto, S. Takeda, J. Jiricny, KIAA1018/FAN1 nuclease protects cells against genomic instability induced by interstrand cross-linking agents, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 21553–21557. [40] C. Lachaud, A. Moreno, F. Marchesi, R. Toth, J.J. Blow, J. Rouse, Ubiquitinated Fancd2 recruits Fan1 to stalled replication forks to prevent genome instability, Science 351 (2016) 846–849. [41] S. Thongthip, M. Bellani, S.Q. Gregg, S. Sridhar, B.A. Conti, Y. Chen, M.M. Seidman, A. Smogorzewska, Fan1 deficiency results in DNA interstrand cross-link repair defects, enhanced tissue karyomegaly, and organ dysfunction, Genes Dev. 30 (2016) 645–659. [42] G.H. Gwon, Y. Kim, Y. Liu, A.T. Watson, A. Jo, T.J. Etheridge, F. Yuan, Y. Zhang, Y. Kim, A.M. Carr, Y. Cho, Crystal structure of a Fanconi anemia-associated nuclease homolog bound to 5' flap DNA: basis of interstrand cross-link repair by FAN1, Genes Dev. 28 (2014) 2276–2290. [43] R. Wang, N.S. Persky, B. Yoo, O. Ouerfelli, A. Smogorzewska, S.J. Elledge, N.P. Pavletich, DNA repair. Mechanism of DNA interstrand cross-link processing by repair nuclease FAN1, Science 346 (2014) 1127–1130. [44] Q. Zhao, X. Xue, S. Longerich, P. Sung, Y. Xiong, Structural insights into 5' flap DNA unwinding and incision by the human FAN1 dimer, Nat. Commun. 5 (2014) 5726. [45] R.D. Shereda, Y. Machida, Y.J. Machida, Human KIAA1018/FAN1 localizes to stalled replication forks via its ubiquitin-binding domain, ABBV Cell Cycle 9 (2010) 3977–3983. [46] P.X. Yan, Y.G. Huo, T. Jiang, Crystal structure of human Fanconi-associated nuclease 1, Protein Cell 6 (2015) 225–228. [47] A. Pingoud, M. Fuxreiter, V. Pingoud, W. Wende, Type II restriction endonucleases: structure and mechanism, Cell. Mol. Life Sci.: CMLS 62 (2005) 685–707. [48] S. Pennell, A.C. Declais, J. Li, L.F. Haire, W. Berg, J.W. Saldanha, I.A. Taylor, J. Rouse, D.M. Lilley, S.J. Smerdon, FAN1 activity on asymmetric repair intermediates is mediated by an atypical monomeric virus-type replication-repair nuclease domain, Cell Rep. 8 (2014) 84–93. [49] D. Takahashi, K. Sato, E. Hirayama, M. Takata, H. Kurumizaka, Human FAN1 promotes strand incision in 5'-flapped DNA complexed with RPA, J. Biochem. 158 (2015) 263–270. [50] J. Pizzolato, S. Mukherjee, O.D. Scharer, J. Jiricny, FANCD2-associated nuclease 1 but not exonuclease 1 or flap endonuclease 1, is able to unhook DNA interstrand cross-links in vitro, J. Biol. Chem. 290 (2015) 22602–22611. [51] M. Castella, T. Taniguchi, The role of FAN1 nuclease in the Fanconi anemia pathway, ABBV Cell Cycle 9 (2010) 4259–4260. [52] L. O'Donnell, D. Durocher, DNA repair has a new FAN1 club, Mol. Cell 39 (2010) 167–169. [53] A. Smogorzewska, M. Kottemann, B. Conti, S. Thongthip, K. Rickman, F. Lach, Molecular mechanism of DNA interstrand crosslink repair, FASEB J. 30 (2016) (500.502). [54] J.P. Trujillo, L.B. Mina, R. Pujol, M. Bogliolo, J. Andrieux, M. Holder, B. Schuster, D. Schindler, J. Surralles, On the role of FAN1 in Fanconi anemia, Blood 120 (2012) 86–89. [55] Y. Fontebasso, T.J. Etheridge, A.W. Oliver, J.M. Murray, A.M. Carr, The conserved Fanconi anemia nuclease Fan1 and the SUMO E3 ligase Pli1 act in two novel Pso2independent pathways of DNA interstrand crosslink repair in yeast, DNA Repair (Amst) 12 (2013) 1011–1023. [56] H.L. Williams, M.E. Gottesman, J. Gautier, The differences between ICL repair during and outside of S phase, Trends Biochem. Sci. 38 (2013) 386–393. [57] W. Zhou, E.A. Otto, A. Cluckey, R. Airik, T.W. Hurd, M. Chaki, K. Diaz, F.P. Lach, G.R. Bennett, H.Y. Gee, A.K. Ghosh, S. Natarajan, S. Thongthip, U. Veturi, S.J. Allen, S. Janssen, G. Ramaswami, J. Dixon, F. Burkhalter, M. Spoendlin, H. Moch, M.J. Mihatsch, J. Verine, R. Reade, H. Soliman, M. Godin, D. Kiss, G. Monga, G. Mazzucco, K. Amann, F. Artunc, R.C. Newland, T. Wiech, S. Zschiedrich, T.B. Huber, A. Friedl, G.G. Slaats, J.A. Joles, R. Goldschmeding, J. Washburn, R.H. Giles, S. Levy, A. Smogorzewska, F. Hildebrandt, FAN1 mutations cause

[58]

[59]

[60]

[61]

[62]

[63]

[64]

[65]

[66]

[67]

[68]

[69]

[70]

[71]

[72]

[73]

[74] [75]

[76] [77]

[78]

[79]

9

karyomegalic interstitial nephritis, linking chronic kidney failure to defective DNA damage repair, Nat. Genet. 44 (2012) 910–915. Y.H. Chen, M.J. Jones, Y. Yin, S.B. Crist, L. Colnaghi, R.J. Sims 3rd, E. Rothenberg, P.V. Jallepalli, T.T. Huang, ATR-mediated phosphorylation of FANCI regulates dormant origin firing in response to replication stress, Mol. Cell. 58 (2015) 323–338. K. Schlacher, H. Wu, M. Jasin, A distinct replication fork protection pathway connects Fanconi anemia tumor suppressors to RAD51-BRCA1/2, Cancer Cell 22 (2012) 106–116. I. Chaudhury, D.R. Stroik, A. Sobeck, FANCD2-controlled chromatin access of the Fanconi-associated nuclease FAN1 is crucial for the recovery of stalled replication forks, Mol. Cell. Biol. 34 (2014) 3939–3954. N. Segui, L.B. Mina, C. Lazaro, R. Sanz-Pamplona, T. Pons, M. Navarro, F. Bellido, A. Lopez-Doriga, R. Valdes-Mas, M. Pineda, E. Guino, A. Vidal, J.L. Soto, T. Caldes, M. Duran, M. Urioste, D. Rueda, J. Brunet, M. Balbin, P. Blay, S. Iglesias, P. Garre, E. Lastra, A.B. Sanchez-Heras, A. Valencia, V. Moreno, M.A. Pujana, A. Villanueva, I. Blanco, G. Capella, J. Surralles, X.S. Puente, L. Valle, Germline mutations in FAN1 cause hereditary colorectal cancer by impairing DNA repair, Gastroenterology 149 (2015) 563–566. A.L. Smith, N. Alirezaie, A. Connor, M. Chan-Seng-Yue, R. Grant, I. Selander, C. Bascunana, A. Borgida, A. Hall, T. Whelan, S. Holter, T. McPherson, S. Cleary, G.M. Petersen, A. Omeroglu, E. Saloustros, J. McPherson, L.D. Stein, W.D. Foulkes, J. Majewski, S. Gallinger, G. Zogopoulos, Candidate DNA repair susceptibility genes identified by exome sequencing in high-risk pancreatic cancer, Cancer Lett. 370 (2016) 302–312. I. Ionita-Laza, B. Xu, V. Makarov, J.D. Buxbaum, J.L. Roos, J.A. Gogos, M. Karayiorgou, Scan statistic-based analysis of exome sequencing data identifies FAN1 at 15q13.3 as a susceptibility gene for schizophrenia and autism, Proc. Natl. Acad. Sci. U. S. A. 111 (2014) 343–348. G.H. Gwon, A. Jo, K. Baek, K.S. Jin, Y. Fu, J.B. Lee, Y. Kim, Y. Cho, Crystal structures of the structure-selective nuclease Mus81-Eme1 bound to flap DNA substrates, EMBO J. 33 (2014) 1061–1072. J. Orans, E.A. McSweeney, R.R. Iyer, M.A. Hast, H.W. Hellinga, P. Modrich, L.S. Beese, Structures of human exonuclease 1 DNA complexes suggest a unified mechanism for nuclease family, Cell 145 (2011) 212–223. S.E. Tsutakawa, S. Classen, B.R. Chapados, A.S. Arvai, L.D. Finger, G. Guenther, C.G. Tomlinson, P. Thompson, A.H. Sarker, B. Shen, P.K. Cooper, J.A. Grasby, J.A. Tainer, Human flap endonuclease structures, DNA double-base flipping, and a unified understanding of the FEN1 superfamily, Cell 145 (2011) 198–211. S. Sung, F. Li, Y.B. Park, J.S. Kim, A.K. Kim, J. O. k. Song, J. Kim, S.E. Che, Y. Lee, D.N.A. Cho, end recognition by the Mre11 nuclease dimer: insights into resection and repair of damaged DNA, EMBO J. 33 (2014) 2422–2435. Y. Liu, S. Sung, Y. Kim, F. Li, G. Gwon, A. Jo, A.K. Kim, T. Kim, O.K. Song, S.E. Lee, Y. Cho, ATP-dependent DNA binding, unwinding, and resection by the Mre11/ Rad50 complex, EMBO J. 35 (2016) 743–758. R.S. Williams, G. Moncalian, J.S. Williams, Y. Yamada, O. Limbo, D.S. Shin, L.M. Groocock, D. Cahill, C. Hitomi, G. Guenther, D. Moiani, J.P. Carney, P. Russell, J.A. Tainer, Mre11 dimers coordinate DNA end bridging and nuclease processing in double-strand-break repair, Cell 135 (2008) 97–109. J. Kosinski, M. Feder, J.M. Bujnicki, The PD-(D/E)XK superfamily revisited: identification of new members among proteins involved in DNA metabolism and functional predictions for domains of (hitherto) unknown function, BMC Bioinf. 6 (2005) 172. N. McGregor, S. Ayora, S. Sedelnikova, B. Carrasco, J.C. Alonso, P. Thaw, J. Rafferty, The structure of Bacillus subtilis RecU Holliday junction resolvase and its role in substrate selection and sequence-specific cleavage, Structure (London, England: 1993) 13 (2005) 1341–1351. T. Nishino, K. Komori, D. Tsuchiya, Y. Ishino, K. Morikawa, Crystal structure of the archaeal holliday junction resolvase Hjc and implications for DNA recognition, Structure (London, England: 1993) 9 (2001) 197–204. K.P. Hopfner, A. Karcher, D.S. Shin, L. Craig, L.M. Arthur, J.P. Carney, J.A. Tainer, Structural biology of Rad50 ATPase: ATP-driven conformational control in DNA double-strand break repair and the ABC-ATPase superfamily, Cell 101 (2000) 789–800. T.T. Paull, M. Gellert, The 3' to 5' exonuclease activity of Mre 11 facilitates repair of DNA double-strand breaks, Mol. Cell 1 (1998) 969–979. P. Turner, C. Nilsson, D. Svensson, O. Holst, L. Gorton, E.N. Karlsson, Monomeric and dimeric cyclomaltodextrinases reveal different modes of substrate degradation, Biol Brat 60 (2005) 79–87. C. Lachaud, J. Rouse, A route to new cancer therapies: the FA pathway is essential in BRCA1- or BRCA2-deficient cells, Nat. Struct. Mol. Biol. 23 (2016) 701–703. J. Michl, J. Zimmer, F.M. Buffa, U. McDermott, M. Tarsounas, FANCD2 limits replication stress and genome instability in cells lacking BRCA2, Nat. Struct. Mol. Biol. 23 (2016) 755–757. Z. Kais, B. Rondinelli, A. Holmes, C. O'Leary, D. Kozono, A.D. D'Andrea, R. Ceccaldi, FANCD2 maintains fork stability in BRCA1/2-deficient tumors and promotes alternative end-joining DNA repair, Cell Rep. 15 (2016) 2488–2499. A. D’Andrea, BRCA1: a missing link in the fanconi Anemia/BRCA pathway, Cancer Discov. 3 (2013) 376–378.