FRINK Linked Data Fragments server

Matches in SemOpenAlex for { <https://semopenalex.org/work/W2061545358> ?p ?o ?g. }

• W2061545358 endingPage "905" @default.
• W2061545358 startingPage "895" @default.
• W2061545358 abstract "Article17 February 2005free access Structure of an XPF endonuclease with and without DNA suggests a model for substrate recognition Matthew Newman Matthew Newman Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Judith Murray-Rust Judith Murray-Rust Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author John Lally John Lally Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Jana Rudolf Jana Rudolf Centre for Biomolecular Sciences, University of St Andrews, Fife, UK Search for more papers by this author Andrew Fadden Andrew Fadden Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Philip P Knowles Philip P Knowles Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Malcolm F White Malcolm F White Centre for Biomolecular Sciences, University of St Andrews, Fife, UK Search for more papers by this author Neil Q McDonald Corresponding Author Neil Q McDonald Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK School of Crystallography, Birkbeck College, London, UK Search for more papers by this author Matthew Newman Matthew Newman Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Judith Murray-Rust Judith Murray-Rust Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author John Lally John Lally Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Jana Rudolf Jana Rudolf Centre for Biomolecular Sciences, University of St Andrews, Fife, UK Search for more papers by this author Andrew Fadden Andrew Fadden Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Philip P Knowles Philip P Knowles Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK Search for more papers by this author Malcolm F White Malcolm F White Centre for Biomolecular Sciences, University of St Andrews, Fife, UK Search for more papers by this author Neil Q McDonald Corresponding Author Neil Q McDonald Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK School of Crystallography, Birkbeck College, London, UK Search for more papers by this author Author Information Matthew Newman1, Judith Murray-Rust1, John Lally1, Jana Rudolf2, Andrew Fadden1, Philip P Knowles1, Malcolm F White2 and Neil Q McDonald 1,3 1Structural Biology Laboratory, London Research Institute, Cancer Research UK, London, UK 2Centre for Biomolecular Sciences, University of St Andrews, Fife, UK 3School of Crystallography, Birkbeck College, London, UK *Corresponding author. Structural Biology Laboratory, London Research Institute, Cancer Research UK, 44 Lincoln's Inn Fields, London WC2A 3PX, UK. Tel.: +44 207 269 3259; Fax: +44 207 269 3258; E-mail: [email protected] The EMBO Journal (2005)24:895-905https://doi.org/10.1038/sj.emboj.7600581 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The XPF/Mus81 structure-specific endonucleases cleave double-stranded DNA (dsDNA) within asymmetric branched DNA substrates and play an essential role in nucleotide excision repair, recombination and genome integrity. We report the structure of an archaeal XPF homodimer alone and bound to dsDNA. Superposition of these structures reveals a large domain movement upon binding DNA, indicating how the (HhH)2 domain and the nuclease domain are coupled to allow the recognition of double-stranded/single-stranded DNA junctions. We identify two nonequivalent DNA-binding sites and propose a model in which XPF distorts the 3′ flap substrate in order to engage both binding sites and promote strand cleavage. The model rationalises published biochemical data and implies a novel role for the ERCC1 subunit of eukaryotic XPF complexes. Introduction Nucleotide excision repair (NER) is a highly conserved DNA repair pathway able to detect and remove a variety of bulky DNA lesions caused by UV light and environmental mutagens and thereby contribute to the genomic integrity of an organism (Sancar, 1996; Lindahl and Wood, 1999). Defects in NER are associated with three inherited human diseases—xeroderma pigmentosum (XP), trichothiodystrophy (TTD) and Cockayne syndrome (CS)—all of which have severe clinical consequences (reviewed in Lehmann, 2003). NER in higher eukaryotes involves the coordinated assembly of a large number of proteins, including the core NER factors XPC-HR23B, TFIIH, XPA, replication protein A (RPA) and two endonucleases XPG and ERCC1-XPF (Aboussekhra et al, 1995). Together these proteins cooperate to recognise, unravel and excise a 24–32-mer oligonucleotide bearing the DNA lesion prior to filling in the missing gap (Araujo and Wood, 1999). Although prokaryotes have a similar overall repair strategy, a much simpler and structurally unrelated multiprotein complex known as UvrABC carries out the same task (Petit and Sancar, 1999). The availability of fully sequenced archaeal genomes has revealed that many archaea have proteins related to eukaryotic NER factors, including the endonucleases XPF and XPG, rather than to the UvrABC bacterial repair system (White, 2003). XPF (xeroderma pigmentosa complementation group F, also known as ERCC4) associates with a noncatalytic partner ERCC1 (excision repair cross complementarity group 1) to form a structure-specific endonuclease that preferentially cleaves DNA duplexes adjacent to a 3′ single-stranded flap (Figure 1A and B). Such double-stranded (ds)/single-stranded (ss) DNA junctions are found in bubbles, simple Y structures and hairpins and are generated on the 5′ side of bulky DNA lesions (Sijbers et al, 1996a, 1996b). All of these DNA structures can be cleaved by ERCC1-XPF (de Laat et al, 1998). The ERCC1-XPF 3′ flap substrate polarity complements the unrelated XPG endonuclease, which cleaves duplex DNA next to a 5′ single-stranded flap and together these endonucleases are responsible for the dual incision that eliminates the lesion-bearing oligonucleotide (Araujo et al, 2000). More recently, the XPF paralogue, Mus81, was shown to function as a 3′ flap endonuclease when associated with its noncatalytic partner Eme1/Mms4 (Boddy et al, 2001; Kaliraman et al, 2001). Mus81-Eme1 cleaves a different set of branched DNA substrates from eukaryotic XPF, including stalled replication forks, nicked Holliday junctions and D-loops (reviewed in Hollingsworth and Brill, 2004). It has a strict requirement for a 5′ DNA end near the flap junction and generates a nicked DNA product with a gap of five unpaired nucleotides (Bastin-Shanower et al, 2003; Osman et al, 2003). For both XPF and Mus81, the major cleavage site always lies within the upstream duplex present in all the branched DNA structures they recognise (Figure 1B). Figure 1.(A) Domain structures of the XPF superfamily. Sequence numbering corresponds to the human ERCC1-XPF heterodimer, ApeXPF and the human Mus81-Eme1 heterodimer. The red box indicates the catalytic ERK motif within the nuclease domain (blue). HhH motifs are shown in green. Cyan indicates a ‘nuclease-like’ domain lacking catalytic residues. (B) Optimal substrates for crenarchaeal XPF and ERCC1-XPF. SsoXPF has a preference for substrates with dsDNA both upstream and downstream from the cleavage site (i.e. 3′ flap), while eukaryotic ERCC1-XPF prefers splayed duplex substrates. (C) Sequence alignment of selected XPF homologues. Representative sequences are shown for the nuclease and (HhH)2 domains from the euryarchaeal (18 sequences) and eukaryotic (18 sequences) XPFs, and for ERCC1 (18 sequences). Invariant residues within the XPF family are shown in red. Observed secondary structure of ApeXPF is indicated above the sequences. Residues involved in the nuclease–(HhH)2 interface are indicated by cyan triangles. Residues within the (HhH)2 dimer interface are indicated by pink ellipses, and those within the (HhH)2 domain hydrophobic core by pink stars. Mutated residues are indicated by dark blue triangles. Invariant residues conserved within the (HhH)2 domain of individual XPF subgroups are highlighted in green. Download figure Download PowerPoint XPF family members have a catalytic domain followed by a DNA-binding domain containing two consecutive helix–hairpin–helix HhH motifs (Thayer et al, 1995;Aravind et al, 1999; Shao and Grishin, 2000). The catalytic domain contains the active site motif GDXnERKx3D related to prokaryotic endonucleases, while the two HhH motifs form a compact (HhH)2 domain that has been shown to contribute to XPF dimer formation and binds duplex DNA in a sequence-independent manner (Nishino et al, 2003). Mus81 also contains a similar nuclease domain, but is flanked by single HhH motifs. Eukaryotic XPFs have an N-terminal SF2-like helicase domain that apparently lacks essential catalytic residues for ATPase activity (Sgouros et al, 1999). A similar ‘long’ form of XPF is present in most euryarchaea, one example being Hef (Helicase-associated endonuclease for fork-structured DNA) from Pyrococcus furiosus, which has an active helicase domain (Komori et al, 2002). Crenarchaea have a ‘short’ form of XPF that lacks the helicase-like domain and whose catalytic activity is regulated by interaction with PCNA (Figure 1A) (Roberts et al, 2003). These differences in domain architecture may reflect differences in the recruitment of XPFs to branched DNA structures (Komori et al, 2002; Roberts et al, 2003). All XPF family proteins require divalent cations for nuclease activity (Sijbers et al, 1996a, 1996b; Nishino et al, 2003). They can either form heterodimers with much smaller but related partners, ERCC1 (eukaryotic XPF) or Eme1/Mms4 (Mus81), or form homodimers (archaea) (Sijbers et al, 1996a, 1996b; Nishino et al, 2003). This dimeric organisation is critical for stability and catalytic activity of XPF (Gaillard and Wood, 2001; Nishino et al, 2003). Additional functions for the noncatalytic partners of XPF have been proposed, for example, ERCC1 targets XPF to sites of DNA damage through its interaction with the DNA-binding protein XPA (Li et al, 1995). To understand more about XPF architecture and the basis of ds/ssDNA substrate recognition, we have determined the structure of an essentially intact crenarchaeal XPF from Aeropyrum pernix (ApeXPF) in the presence and absence of dsDNA. Analysis of these structures has revealed for the first time a large domain movement on binding dsDNA and has led us to suggest a model for XPF substrate recognition involving distortion of branched and nicked DNA substrates. Results Structure determination For crystallisation purposes, we used a truncated form of ApeXPF comprising residues 19–231 (denoted as ApeXPF throughout). In solution, ApeXPF forms homodimers as does the closely related Sulfolobus solfataricus XPF (SsoXPF) (Lally et al, 2004; Roberts et al, 2004). ApeXPF can be stimulated by a heterotrimeric PCNA to cleave 3′ flap structures; however, SsoXPF has a more robust activity in a nuclease assay and we therefore used SsoXPF for the mutational analysis (Supplementary Figure S1). We tried to crystallise ApeXPF in the presence of DNA with a wide range of 3′ flaps, hairpins and Y structures, and in some cases obtained crystals but none with good diffraction properties. Trigonal crystals were eventually obtained using a 15-mer DNA duplex; they diffracted to 2.8 Å resolution and contain a single ApeXPF dimer (Table I). The structure was solved by molecular replacement using as a search model the ApeXPF nuclease domain derived from a partially refined structure of residues 19–150 (Lally et al, 2004; Materials and methods). The initial electron density maps phased only on the nuclease domains showed good density for the (HhH)2 domain and the duplex DNA (Supplementary Figure S2) and allowed tracing of the entire molecule apart from the interdomain linker of one protomer, which is disordered. The final structure was refined to an R-factor of 23.0% and R-free of 29.0% at 2.8 Å resolution (Table I). Table 1. Data collection and refinement statistics Protein ApeXPF ApeXPF ApeXPF Form Native 1 mM K2OsO4 Native+dsDNA X-ray source Station 14.2, SRS Station 14.2, SRS Station 9.6, SRS Wavelength (Å) 1.0000 1.0000 0.9780 Space group C2 C2 P3221 Unit cell parameters (Å) a=210, b=42.7, c=118.7 a=209, b=42.8, c=119.9 a=b=141.3, c=85.3 β=121.4° β=121.4° Za 4 4 2 No. of measurements 52 273 (7746) 33 163 (4880) 125 841 (8403) No. of unique reflections 14 887 (2159) 9471 (1414) 21 619 (2463) Resolution limit (Å) 3.2 (3.37–3.2) 3.6 (3.79–3.6) 2.8 (2.95–2.8) Completeness 99.9 (100) 98.5 (99.9) 97.5 (92.8) Rsym 6.2 (30.8) 5.0 (32.1) 8.3 (39.9) Average I/σ(I) 10.6 (2.4) 13.1 (2.3) 13.7 (2.8) Ranomalous — 5.0 (23.1) — Rderivative — 26.0 (31.2) — R-factor 0.24 — 0.23 R-free 0.31 — 0.29 Model 6516 protein atoms — 3357 protein atoms, 608 DNA atoms, 6 sulphate ions, 1 Mg ion, 7 H2O R.m.s. bond lengths (Å) 0.02 — 0.021 R.m.s. bond angles (deg) 1.9 — 2.2 Ramachandran plota 86.9%/12.6% 90.0%/8.9% Values in parentheses refer to the highest resolution shell indicated in the row for resolution limit. a Percentage of residues located in the most favourable/additionally allowed regions of the Ramachandran plot. Monoclinic crystals of ApeXPF grown without DNA contain two ApeXPF dimers and diffracted to 3.2 Å resolution. This structure was solved by molecular replacement using the two domains of the DNA-bound structure separately as search models and guided by heavy atoms sites identified from an osmate derivative (see Materials and methods). Two of the four protomers had continuous density including the linker sequence, while the other two had disordered interdomain linkers. This structure was refined to an R-factor of 24.0% and R-free of 31.0% at 3.2 Å resolution (Table I). Structure of apo-ApeXPF ApeXPF 19–231 has two structured domains, an N-terminal nuclease domain (residues 19–148) and an (HhH)2 domain (residues 165–226) connected by a 15-residue linker (Figures 1C and 2A). As expected by analogy with Hef (Nishino et al, 2003), both the nuclease and (HhH)2 domains form independent, tightly associated dimers with an equivalent domain from the second protomer. In this arrangement, the domains are uncoupled. The α/β nuclease domain structure is closely related to the Hef nuclease domain and the protomers make similar dimer contacts via two helices (α4 and α5) and an edge β-strand (β6) (Nishino et al, 2003). The active site lies in a large cleft bounded by the structural elements α1′, α2, α3 and a loop between strands β2 and β3 (Figure 1C). This cleft is lined with the overwhelming majority of XPF conserved residues including the metal-binding and catalytic residues from the signature sequence E-R-K (E62-R63-K64) that runs along the base of the cleft as part of strand β4. There is density at a site equivalent to the metal site observed in Hef (Nishino et al, 2003). Similar density is present in the ApeXPF–DNA complex, which due to its higher resolution shows more precisely that the coordination geometry and distance to side chains of D52 and E62 are consistent with it being a metal ion, presumably magnesium. Figure 2.(A) Structure of the apo form of the ApeXPF dimer. Protein domains are coloured to correspond with Figure 1A. The local two-fold axes relating the nuclease and (HhH)2 domains are shown as black and magenta lines, respectively. (B) Two views of the ApeXPF–DNA complex. Protein domains are coloured to correspond with Figure 1A. The carbon atoms of the DNA C strand are yellow and those of the DNA D strand are grey. (C) Domain rearrangement between ApeXPF with and without DNA shown by superposing the nuclease domains of protomer A from the two structures. The ‘open’ apo structure is on the left, the ‘closed’ DNA-bound structure on the right and the two superposed in the middle. Download figure Download PowerPoint Each (HhH)2 domain forms an integral five-helical domain bearing two functional helix–hairpin–helix motifs related by pseudo-two-fold symmetry (Thayer et al, 1995; Doherty et al, 1996; Shao and Grishin, 2000). A similar domain is present in RuvA (PDB code 1C7Y) (Ariyoshi et al, 2000). The RuvA (HhH)2 domain has an r.m.s. difference of 1.55 Å (54 C-alpha atoms) and 1.7 Å (53 C-alpha atoms) with the (HhH)2 domains from the two protomers of the ApeXPF–DNA structure. ApeXPF has two G-I-G hairpins at residues 179–181 and 211–213. Although the (HhH)2 domain fold has been observed previously (Thayer et al, 1995; Rafferty et al, 1996; Singh et al, 2002), this is the first structure of a dimer of (HhH)2 domains. There are three major contacts in this predominantly hydrophobic dimer interface (Figure 1C and Supplementary Figure S3). These centre around the connecting helix that links the two HhH motifs, residues from the first helix of the first HhH motif and residues C-terminal to the second HhH motif. A C-terminal extension to the domain contributes the highly conserved Y228 to the dimer interface. The two independent dimers in the asymmetric unit of the apo ApeXPF crystals are very similar and both have the linker region of one protomer fully extended. The local two-fold axes relating the protomers of the nuclease and (HhH)2 domain dimers respectively are not coincident so that overall the ApeXPF dimer does not obey two-fold symmetry (Figure 2A). Each of the independent apo ApeXPF dimers is in close contact with another identical dimer related by operation of a crystallographic two-fold axis generating a tetramer. The contact involves two equivalent linker regions that form a two-stranded antiparallel sheet across the two-fold axis (Supplementary Figure S4). Comparison of both tetrameric arrangements indicates an r.m.s. difference of 3.4 Å on 802 C-alpha atoms. Structure of an ApeXPF–dsDNA complex The ApeXPF–dsDNA structure comprises two ApeXPF protomers (denoted A and B), together with one DNA duplex (strands denoted C and D) (Figure 2B). The DNA duplex is bound to the nuclease and (HhH)2 domains of protomer A, and this we define as site I. Overall, the ApeXPF in the complex forms an asymmetric dimer in which protomer A is more compact and has an ordered interdomain linker (Figure 2B). Comparing the A chains of the DNA-bound and DNA-free forms shows that when their nuclease domains are superimposed, the two orientations of the (HhH)2 domains are related by a rotation of 95°, which is primarily a closure rather than twist (Figure 2C). This shifts the (HhH)2 domains by over 30 Å. The linker forms a complex hinge region with the main flexures at residues 151–155 and 164. In the DNA-bound form, it contributes L163 to the interface between the nuclease and (HhH)2 domains (Figure 1C and Supplementary Figure S5). The DNA-bound ApeXPF structure is dominated by the interaction of the (HhH)2 domain of protomer A with dsDNA. Interdomain interactions between the nuclease and (HhH)2 domains of the A protomer further stabilise the ApeXPF conformation and bury several aromatic residues at the interface close to W169 (Figure 1C and Supplementary Figure S5). Protomer B makes almost no contact with the DNA at site I, and its nuclease and (HhH)2 domain do not interact; its interdomain linker is not seen in the electron density maps, but must adopt an extended conformation because protomer B residues 148 and 160 are 29 Å apart. The interdomain linker has been suggested to be sensitive to proteolysis due to conformational flexibility (Nishino et al, 2003). We therefore dissolved ApeXPF–dsDNA crystals and confirmed by SDS–PAGE that no proteolytic cleavage had occurred, thus confirming the integrity of both linkers (data not shown). Site I recognition involves binding of a six base-pair stretch of minor groove by the (HhH)2 domain of protomer A and the interaction of a blunt end (representing a ds/ss discontinuity) with the catalytic domain of protomer A. The stretch of minor groove recognised by the (HhH)2 domain is just two base pairs from the end of the duplex and this ensures that when the nuclease and (HhH)2 domains become coupled, the duplex DNA is oriented with the 3′ end of one strand adjacent to Y123 (Figure 3A and B). The 5′ end of strand D lies adjacent to a crystallisation-derived sulphate ion bound to R126 from both protomers A and B. Binding DNA with an opposite polarity would not be possible, as the HhH hairpin footprint could not accommodate the much wider major groove. The G-I-G hairpin from one HhH motif binds two backbone phosphates from T13 and G14 of DNA strand C, while the other hairpin binds phosphates from T7 and C8 of the opposite sense strand D (Figure 3A). Several positively charged residues immediately after the hairpins (R182, R183 and R187 of the first HhH motif and K215 and R216 of the second HhH motif) also contribute to the interaction with the minor groove. Of these, sequence-independent contacts include R187 interaction with the C12 phosphate backbone of the C strand, and R216 penetration into the minor groove to bind T7 (D strand) backbone phosphate. The unique orientation of dsDNA is assisted by a sequence-dependent contact from R182 of the first HhH motif to guanine-14 of strand C. Protomer A's nuclease domain contacts a blunt end of the DNA duplex through hydrophobic interactions (residues A86, Y123 and G124) with the two end bases A15 (strand C) and T1 (strand D) (Figure 3). The blunt end is not a feature of preferred XPF substrates but here represents a ds/ss discontinuity and lies adjacent to a shallow hydrophobic groove (Figure 3B and C) that may contact ssDNA (see below). Figure 3.(A) Schematic of ApeXPF interaction with the 15-mer dsDNA. Hydrogen bonds to DNA involving protein side chains are shown as magenta arrows, and those involving protein backbone are shown as blue arrows. Hydrophobic interactions are represented by dashed lines. Blue and cyan boxes around residue numbers represent nuclease domains from protomers A and B, respectively, and green boxes indicate the (HhH)2 domain contacts. A small contact between DNA and a symmetry-related protein molecule is indicated for G1–C15. (B) The hydrophobic groove on the nuclease domain of protomer A. All side chains are shown as sticks. This is a possible ssDNA-binding site, and aromatic residues that might stack with or intercalate between DNA bases are labelled. The strip is bordered by positively charged side chains, some of which could interact with the phosphate backbone. (C) Protein electrostatic surface in the ApeXPF–DNA complex. Red indicates acidic regions and blue indicates basic regions. Only protein atoms were included in the calculation of the electrostatic potential with APBS (Baker et al, 2001). Download figure Download PowerPoint A two-site model for ds/ssDNA junction recognition by XPF The (HhH)2 domain of protomer A is a major component of site I, but the (HhH)2 domain of protomer B is also potentially capable of binding DNA. We note that the two-fold axis relating the (HhH)2 domains of protomers A and B makes an angle close to 45° with the helical axis of the dsDNA. Crystal contacts prevent the protomer B (HhH)2 domain from engaging dsDNA in the trigonal crystal form. Instead, a sulphate ion occupies a position equivalent to the phosphate backbone position of the DNA bound to protomer A's first HhH motif. We therefore modelled how protomer B's (HhH)2 domain might bind DNA. As the (HhH)2 domain dimer two-fold axis is at 45° to the helical axis of the dsDNA, rotating the dsDNA (strands C and D) about this two-fold axis generates a second putative dsDNA molecule (strands C′ and D′) at 90° to the experimentally observed dsDNA (Figure 4A). Inspection of the modelled DNA in the context of the crystallographically observed complex shows one end of the C′D′ duplex close to the active site of the nuclease domain of protomer A (Figure 4B). A second potential DNA-binding site (denoted site II) therefore exists, which comprises the (HhH)2 domain of protomer B and the active site of protomer A, although it is unoccupied in our structure. Figure 4.(A) Interaction of DNA with site I, and modelling of DNA bound to site II. Site I-bound dsDNA is shown in stick representation, and the GIG-containing hairpin regions of the protomer A (HhH)2 domain are highlighted in red. The two-fold axis between the (HhH)2 domains of protomers A and B is indicated by a magenta line, and the approximate direction of the DNA helical axis is shown by a black line. The modelled DNA generated by rotation of the observed DNA around the (HhH)2 two-fold axis (backbone representation) is in the correct orientation to interact with the two hairpin motifs of protomer B. (B) A structural model for DNA substrate binding to ApeXPF. (HhH)2 domains and DNA are drawn as in (A), but nuclease domains are also included. In our model, the observed DNA at site I would be downstream of the ds/ss junction and the modelled DNA at site II would be upstream. The sulphate ions found in the structure (shown in red/yellow) mimic the phosphate backbone of site II DNA. The binding sites are also shown schematically with DNA strands and protein chains coloured according to Figures 1B and 5C, and 2, respectively. (C) Substrate preferences for SsoXPF as determined by Roberts et al (2004). Distinct features of the preferred 3′ flap substrate are highlighted. A double greater than sign indicates at least a 10-fold improvement in the rate of cleavage by SsoXPF, whereas a single greater than sign indicates one- to three-fold improvement. (D) Stereo view of the protomer A active site. The ERK active site motif has light red side chains, and other key conserved side chains are labelled. Sulphate ions are shown where they may mimic phosphate backbone binding sites. Also shown are the side chain of R126 and the adjacent sulphate ion, which lies close to the 5′ end of strand D. (E) Identical stereo view of the protomer A active site to (A) but including the modelled second duplex DNA at site II generated by rotation about the (HhH)2 domain dimer dyad axis as described in the text. Download figure Download PowerPoint We also assume that ApeXPF has similar substrate preferences to SsoXPF, which cleaves 3′ flaps and nicked duplexes preferentially over splayed duplexes (Figure 4C; Roberts et al, 2004). Both these preferred substrates have two segments of contiguous dsDNA and therefore the observed crystallographic DNA duplex at site I could represent either the upstream or downstream duplex of such substrates. Since XPF is known to cleave only within the upstream duplex of a 3′ flap (Sijbers et al, 1996a, 1996b; de Laat et al, 1998; Bastin-Shanower et al, 2003; Osman et al, 2003; Roberts et al, 2003), and the 3′ end of the C strand at site I is 17 Å from the active site of protomer A (and 35 Å from the active site of protomer B), we consider it more likely that dsDNA bound at site I represents the downstream part of such a substrate. Strand C would therefore be part of the continuous strand (or undamaged strand) (Figure 4B). This would imply that site II binds the upstream duplex, which would be processed by the active site of protomer A. This arrangement would allow for the XPF cleavage sites being upstream of the ds/ssDNA junction whether the duplex traverses the active site or the strands are separated. It is evident that in order for both (HhH)2 domains to engage DNA substrate simultaneously, a 3′ flap substrate would have to bend by almost 90° (Figure 4B). The 3′ end of the modelled C′ strand lies close to the catalytic centre of protomer A and adjacent to the absolutely conserved residues solvent accessible R26 and R77, just outside of the catalytic cleft (Figure 4D and E). The C′ strand is therefore positioned with the expected polarity for the cleaved strand and with dsDNA adjacent to the active site, in agreement with experimental data. The polarity of the C′ strand (5′-3′) and the presence of two sulphate ions adjacent to the invariant Q81 and R61 suggest how the 3′ flap is led into the active site (Figure 4D and E). A closer view of how the cleaved strand is positioned within the active site cleft requires structures of XPF with longer and branched DNA substrates, a goal we are working towards. The 3′ end of strand C at site I abuts a shallow hydrophobic groove in the nuclease domain extending between helices α3 and α4 (Figure 3B and C). The groove is lined with several conserved aromatic residues centred on F79 whose spacings approximate to base separation, and this is suggestive of an ssDNA-binding site. Variable length gapped products with a short single-stranded region in the continuous DNA strand are generated by SsoXPF, ERCC1-XPF and Mus81-Eme1 as the cleavage site is anywhere from 2 to 8 nucleotides 5′ of the ds/ssDNA junction (Figure 1B) (Sijbers et al, 1996a, 1996b; de Laat et al, 1998; Bastin-Shanower et al, 2003; Osman et al, 2003; Roberts et al, 2003). The hydrophobic groove may tether such a region extending from the 5′ end of the modelled strand D′ (and thus representing the continuous, u" @default.
• W2061545358 created "2016-06-24" @default.
• W2061545358 creator A5009496843 @default.
• W2061545358 creator A5015511982 @default.
• W2061545358 creator A5029677401 @default.
• W2061545358 creator A5032323043 @default.
• W2061545358 creator A5046171490 @default.
• W2061545358 creator A5050051838 @default.
• W2061545358 creator A5055052454 @default.
• W2061545358 creator A5057021821 @default.
• W2061545358 date "2005-02-17" @default.
• W2061545358 modified "2023-09-23" @default.
• W2061545358 title "Structure of an XPF endonuclease with and without DNA suggests a model for substrate recognition" @default.
• W2061545358 cites W136476865 @default.
• W2061545358 cites W1511719954 @default.
• W2061545358 cites W1608467849 @default.
• W2061545358 cites W1886853229 @default.
• W2061545358 cites W1953968639 @default.
• W2061545358 cites W1964714687 @default.
• W2061545358 cites W1981680259 @default.
• W2061545358 cites W1990849368 @default.
• W2061545358 cites W1992082799 @default.
• W2061545358 cites W1994561103 @default.
• W2061545358 cites W1994608549 @default.
• W2061545358 cites W1994875540 @default.
• W2061545358 cites W1996372461 @default.
• W2061545358 cites W1999590520 @default.
• W2061545358 cites W2001167026 @default.
• W2061545358 cites W2001641653 @default.
• W2061545358 cites W2013083986 @default.
• W2061545358 cites W2021084223 @default.
• W2061545358 cites W2028774533 @default.
• W2061545358 cites W2029582401 @default.
• W2061545358 cites W2036103078 @default.
• W2061545358 cites W2051031816 @default.
• W2061545358 cites W2056474830 @default.
• W2061545358 cites W2064577894 @default.
• W2061545358 cites W2071106544 @default.
• W2061545358 cites W2078983417 @default.
• W2061545358 cites W2080605611 @default.
• W2061545358 cites W2082398105 @default.
• W2061545358 cites W2090441795 @default.
• W2061545358 cites W2102265131 @default.
• W2061545358 cites W2108572433 @default.
• W2061545358 cites W2109113345 @default.
• W2061545358 cites W2112200653 @default.
• W2061545358 cites W2113769029 @default.
• W2061545358 cites W2117538545 @default.
• W2061545358 cites W2118726693 @default.
• W2061545358 cites W2125489655 @default.
• W2061545358 cites W2130460588 @default.
• W2061545358 cites W2142341469 @default.
• W2061545358 cites W2143694829 @default.
• W2061545358 cites W2150072036 @default.
• W2061545358 cites W2155447972 @default.
• W2061545358 cites W2156716526 @default.
• W2061545358 cites W2160781488 @default.
• W2061545358 cites W2162551655 @default.
• W2061545358 cites W2165616356 @default.
• W2061545358 cites W2166424758 @default.
• W2061545358 cites W2172916891 @default.
• W2061545358 cites W4237568215 @default.
• W2061545358 doi "https://doi.org/10.1038/sj.emboj.7600581" @default.
• W2061545358 hasPubMedCentralId "https://www.ncbi.nlm.nih.gov/pmc/articles/554130" @default.
• W2061545358 hasPubMedId "https://pubmed.ncbi.nlm.nih.gov/15719018" @default.
• W2061545358 hasPublicationYear "2005" @default.
• W2061545358 type Work @default.
• W2061545358 sameAs 2061545358 @default.
• W2061545358 citedByCount "109" @default.
• W2061545358 countsByYear W20615453582012 @default.
• W2061545358 countsByYear W20615453582013 @default.
• W2061545358 countsByYear W20615453582014 @default.
• W2061545358 countsByYear W20615453582015 @default.
• W2061545358 countsByYear W20615453582016 @default.
• W2061545358 countsByYear W20615453582017 @default.
• W2061545358 countsByYear W20615453582018 @default.
• W2061545358 countsByYear W20615453582019 @default.
• W2061545358 countsByYear W20615453582020 @default.
• W2061545358 countsByYear W20615453582021 @default.
• W2061545358 countsByYear W20615453582022 @default.
• W2061545358 crossrefType "journal-article" @default.
• W2061545358 hasAuthorship W2061545358A5009496843 @default.
• W2061545358 hasAuthorship W2061545358A5015511982 @default.
• W2061545358 hasAuthorship W2061545358A5029677401 @default.
• W2061545358 hasAuthorship W2061545358A5032323043 @default.
• W2061545358 hasAuthorship W2061545358A5046171490 @default.
• W2061545358 hasAuthorship W2061545358A5050051838 @default.
• W2061545358 hasAuthorship W2061545358A5055052454 @default.
• W2061545358 hasAuthorship W2061545358A5057021821 @default.
• W2061545358 hasBestOaLocation W20615453581 @default.
• W2061545358 hasConcept C181199279 @default.
• W2061545358 hasConcept C18903297 @default.
• W2061545358 hasConcept C2777028655 @default.
• W2061545358 hasConcept C2777289219 @default.
• W2061545358 hasConcept C2994592520 @default.
• W2061545358 hasConcept C54355233 @default.
• W2061545358 hasConcept C552990157 @default.
• W2061545358 hasConcept C55493867 @default.

• next