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• W1844128776 abstract "Article2 September 2011free access Crystal structures of an archaeal class II DNA photolyase and its complex with UV-damaged duplex DNA Stephan Kiontke Stephan Kiontke Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany Search for more papers by this author Yann Geisselbrecht Yann Geisselbrecht Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany Search for more papers by this author Richard Pokorny Richard Pokorny Faculty of Biology, Department of Plant Physiology and Photobiology, Philipps University, Marburg, Germany Search for more papers by this author Thomas Carell Thomas Carell Center of Integrative Protein Science (CiPSM) at the Department of Chemistry and Biochemistry, Ludwig-Maximilians University, Munich, Germany Search for more papers by this author Alfred Batschauer Corresponding Author Alfred Batschauer Faculty of Biology, Department of Plant Physiology and Photobiology, Philipps University, Marburg, Germany Search for more papers by this author Lars-Oliver Essen Corresponding Author Lars-Oliver Essen Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany Search for more papers by this author Stephan Kiontke Stephan Kiontke Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany Search for more papers by this author Yann Geisselbrecht Yann Geisselbrecht Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany Search for more papers by this author Richard Pokorny Richard Pokorny Faculty of Biology, Department of Plant Physiology and Photobiology, Philipps University, Marburg, Germany Search for more papers by this author Thomas Carell Thomas Carell Center of Integrative Protein Science (CiPSM) at the Department of Chemistry and Biochemistry, Ludwig-Maximilians University, Munich, Germany Search for more papers by this author Alfred Batschauer Corresponding Author Alfred Batschauer Faculty of Biology, Department of Plant Physiology and Photobiology, Philipps University, Marburg, Germany Search for more papers by this author Lars-Oliver Essen Corresponding Author Lars-Oliver Essen Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany Search for more papers by this author Author Information Stephan Kiontke1, Yann Geisselbrecht1, Richard Pokorny2, Thomas Carell3, Alfred Batschauer 2 and Lars-Oliver Essen 1 1Faculty of Chemistry, Department of Biochemistry, Philipps University, Marburg, Germany 2Faculty of Biology, Department of Plant Physiology and Photobiology, Philipps University, Marburg, Germany 3Center of Integrative Protein Science (CiPSM) at the Department of Chemistry and Biochemistry, Ludwig-Maximilians University, Munich, Germany *Corresponding authors: Faculty of Biology, Department of Plant Physiology and Photobiology, Philipps University, Karl-von-Frisch-Strasse 8, 35032 Marburg, Germany. Tel.:+49 6421 282 7064; Fax: +49 6421 282 1545; E-mail: [email protected] of Chemistry, Department of Biochemistry, Philipps University, Hans-Meerwein-Strasse, 35032 Marburg, Germany. Tel.: +49 6 421 282 2032; Fax: +49 6 421 282 2191; E-mail: [email protected] The EMBO Journal (2011)30:4437-4449https://doi.org/10.1038/emboj.2011.313 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions Figures & Info Class II photolyases ubiquitously occur in plants, animals, prokaryotes and some viruses. Like the distantly related microbial class I photolyases, these enzymes repair UV-induced cyclobutane pyrimidine dimer (CPD) lesions within duplex DNA using blue/near-UV light. Methanosarcina mazei Mm0852 is a class II photolyase of the archaeal order of Methanosarcinales, and is closely related to plant and metazoan counterparts. Mm0852 catalyses light-driven DNA repair and photoreduction, but in contrast to class I enzymes lacks a high degree of binding discrimination between UV-damaged and intact duplex DNA. We solved crystal structures of Mm0852, the first one for a class II photolyase, alone and in complex with CPD lesion-containing duplex DNA. The lesion-binding mode differs from other photolyases by a larger DNA-binding site, and an unrepaired CPD lesion is found flipped into the active site and recognized by a cluster of five water molecules next to the bound 3′-thymine base. Different from other members of the photolyase-cryptochrome family, class II photolyases appear to utilize an unusual, conserved tryptophane dyad as electron transfer pathway to the catalytic FAD cofactor. Introduction UV-induced lesions in B-type duplex DNA are either the pyrimidine-pyrimidone (6-4) photoproduct or the predominant cyclobutane pyrimidine dimer (CPD) in form of its cis-syn isomer (Heelis et al, 1993). These UV lesions are repaired by light-dependent DNA-repair enzymes called DNA photolyases, which are members of the structurally related photolyase-cryptochrome family that also comprises cryptochromes and DASH cryptochromes. Cryptochromes act as blue-light photoreceptors and exert various physiological functions like regulation of the circadian clock in animals and plants. Unlike photolyases they generally lack any kind of DNA-repair activity (Lin and Todo, 2005), whereas DNA photolyases can be specified according to their substrate specificity as (6-4) or CPD photolyases, respectively. The photolyase-cryptochrome family is present in all three domains of life, that is, archaea, eubacteria and eukaryotes, and hence has arisen very early during evolution to protect genomes against the genotoxic effects of ultraviolet light originating from the sun. However, their evolution and the phylogenetic relationship of its members and subfamilies have been controversially discussed (Kanai et al, 1997; Falciatore and Bowler, 2005; Ozturk et al, 2008; Lucas-Lledo and Lynch, 2009). Based on sequence analyses, CPD photolyases have been initially subdivided into just two classes: class I enzymes occurring exclusively in microbes and class II photolyases mostly restricted to higher, multicellular eukaryotes. Only recently, cryptochromes of the DASH type have been recognized to catalyse light-driven CPD-repair activity in single-stranded and loop-structured duplex DNA as well (Selby and Sancar, 2006; Pokorny et al, 2008). Accordingly, other but distant subfamilies like class III photolyases present in some eubacteria (Ozturk et al, 2008) or a novel type of cryptochromes occurring in proteobacterial species (Hendrischk et al, 2009) have been discovered. Overall, this implies that the ancient photolyase-cryptochrome family is highly diversified and that the insights derived so far from the well-characterized class I CPD and (6-4) photolyases are not necessarily applicable to other subfamilies. For example, the cryptochrome subfamilies from plants and animals as well as the DASH cryptochromes have apparently branched off from class I CPD and (6-4) photolyases, respectively, but not from class II enzymes or other subfamilies. Both photolyases and cryptochromes have a bilobal architecture consisting of two domains: an N-terminal domain that may contain a light-harvesting antenna chromophore to additionally broaden their activity spectra and a C-terminal α-helical catalytic domain comprising the light-sensitive FAD cofactor. This architecture is preserved in the structurally characterized class I photolyases (Park et al, 1995; Tamada et al, 1997; Komori et al, 2001; Fujihashi et al, 2007), (6-4) photolyases (Maul et al, 2008; Hitomi et al, 2009), plant cryptochromes (Brautigam et al, 2004) as well as DASH cryptochromes (Brudler et al, 2003; Klar et al, 2007), although some differences have been described. For example, the catalytic domain of the Thermus thermophilus CPD photolyase is C-terminally truncated by ∼20 residues as compared with other class I photolyases (Komori et al, 2001), whereas mature cry3 from Arabidopsis thaliana bears an N-terminal extension of 39 residues (Klar et al, 2007) that is present only in some plant but not in cyanobacterial orthologues (Brudler et al, 2003). Diverse classes of antenna chromophores like 5,10-methenyltetrahydrofolate (MTHF), 8-hydroxydeazaflavin, FMN or FAD have been identified in some photolyases/cryptochromes to broaden their activity spectra, whereas many others apparently lack any bound antenna chromophores. In DNA photolyases, binding and repair of UV-damaged DNA is the defining function of the catalytic domain. The low intrinsic affinity of class I CPD photolyases to intact DNA in the micromolar range is mediated by the basic nature of the protein's surface surrounding the active site with its catalytic FAD cofactor. The specific recognition and repair of the UV lesion occurs within the active site and its rectangular-shaped entrance. The pyrimidine dimer is flipped out of the double-stranded DNA (dsDNA) into the active site, forming numerous hydrophobic interactions at its bottom as well as hydrogen bonds between the C4-carbonyls of the 5′- and 3′-pyrimidine bases and the N6-amino group of the FAD's adenine moiety. This flavin cofactor participates in the light-mediated reactivation of UV-damaged DNA in its reduced and photoexcited FADH−* state. Within nanoseconds, injection of an electron into the CPD lesion causes the breakage of the C5-C5 and C6-C6 bonds and electron back-transfer to the FADH• intermediate (Sancar, 2003; Essen, 2006). Cocrystal structures of the class I photolyase from Anacystis nidulans (Mees et al, 2004) and the (6-4) photolyase from Drosophila melanogaster (Maul et al, 2008) with UV-damaged duplex DNA demonstrated that sequence-independent recognition of the UV lesion depended on salt bridges and hydrogen bonds mainly formed between the enzyme and the P−1, P+1, P+2 and P+3 phosphates of the DNA strand comprising the lesion. There is a distinct lack of interactions with the counter strand rationalizing why DNA photolyases catalyze very efficiently the repair of UV-B damaged ssDNA as well. With sequence identities of <16%, class II photolyases are only distantly related to other photolyase/cryptochrome subfamilies (Yasui et al, 1994) suggesting that they have arisen early in evolution (Kanai et al, 1997) and may differ in their mode of FAD binding and catalysis, although photoreduction of their FAD cofactor occurs similarly to class I via a semireduced FADH• intermediate to the catalytically active FADH− (Okafuji et al, 2010). Class II photolyases occur ubiquitously in multicellular organisms, that is, plants and animals, but are missing in placental animals like humans and mice. Despite their functional importance, overall knowledge about class II photolyases lags behind their microbial class I relatives. Here, a combined structural, spectroscopic and functional analysis of the class II photolyase from the methanogenic archaeon Methanosarcina mazei Go1, the gene product Mm0852, is reported. Its close relationship to homologous enzymes from plants (Iwamatsu et al, 2008), and animals like the mammalian infraclass of marsupials, for example Potorous tridactylus (Yasui et al, 1994), makes the described structural insights highly transferable to all other metazoan photolyases. Results and discussion An archaeal orthologue as a representative of metazoan class II photolyases A phylogenetic sequence analysis of light-dependent DNA-repair enzymes was performed to identify orthologues of class II photolyases from plants which might be better suited for structural and biophysical analysis than the available recombinant plant enzymes (Kleiner et al, 1999; Kaiser et al, 2009). The unrooted phylogenetic tree (Figure 1A) reveals a distinct cluster for class II photolyases which does not only comprise all known orthologues from higher, multicellular eukaryotes like A. thaliana, O. sativa, D. melanogaster or P. tridactylus and some viruses, but also from several eubacteria and archaea. An analysis of the INTERPRO protein subfamily of class II photolyases (IPR008148) shows currently 117 eukaryotic, 91 eubacterial and 6 archaeal members including the euryarchaeal order of Methanosarcinales. Obviously, the occurrence of class II photolyases is much more restricted in prokaryotes than that of class I, because the class I-specific INTERPRO subfamily (IPR002081) comprises 1257 eubacterial and 35 archaeal orthologues. Moreover, among the class II photolyases, the subfamily members from plants and animals cluster together with the Methanosarcinal photolyases from M. mazei, M. acetivorans and M. barkeri into a common subbranch, where they exhibit pair-wise sequence identities of 45–48% (Figure 1C and D; Supplementary Figure S4B). Accordingly, one may postulate that at least some of the overall well-conserved class II photolyases were spread by horizontal gene transfer, for example, Methanosarcinales ↔ metazoans, and may hence be a result of late evolution. For comparison, the sequence identities between class II and microbial class I photolyases of Escherichia coli and A. nidulans (EcCPDI and AnCPDI) are 16% or less (Figure 1D; Supplementary Figure S4A). Noteworthy, for class II photolyases natural fusions with transcription factors, NAD-dependent reductases or chaperones can be identified in several genomes (Figure 1B). Structural and functional analysis of methanosarcinal photolyases may be hence rewarding as they are highly representative of related photolyases from plants, animals and viruses. Figure 1.Phylogenetic analysis of the class II photolyase subfamily. (A) The class II CPD photolyase subfamily forms a distinct cluster of light-dependent DNA-repair enzymes. The scale bar of the unrooted phylogenetic tree indicates amino-acid substitutions per site. (B) Several class II photolyases (UniProt identifiers depicted at left) show various kinds of domain fusions at the N- or C-terminal subdomains (blue and green). (C) Dendrogram of 98 non-redundant members of the class II photolyase subfamily. (D) Sequence identities of prominent members of class II and class I CPD photolyases. Class II CPD photolyases: Methanosarcina mazei (MmCPDII), Methanobacterium thermoautotrophicum (MtCPDII), Arabidopsis thaliana (AtCPDII), Oryza sativa (OsCPDII), Drosophila melanogaster (DmCPDII), Potorous tridactylus (PtCPDII), Chlorobium ferrooxidans (CfCPDII) and Halothermothrix orenii (HoCPDII); Class I CPD photolyases: Escherichia coli (EcCPDI) and Anacystis nidulans (AnCPDI). Phylogenetic analysis of class II photolyases is summarized in Supplementary data. Download figure Download PowerPoint UV/Vis spectra, repair activity and DNA binding of M. mazei class II photolyase The Mm0852 gene from M. mazei, an orthologue of class II photolyases, was cloned and heterologously overexpressed in E. coli giving ∼50 mg protein per litre culture. After purification, the Mm0852 gene product was of a bright yellow colour indicating the presence of a flavin cofactor in the fully oxidized form. Its absorption spectrum exhibits peaks at 362, 377, 421, 444 and 469 nm (Figure 2A), which are characteristic for photolyase-bound FADox and differ to free FADox in solution as the latter one has peaks at 373 and 445 nm (Payne et al, 1990). The absence of any prominent absorption peak between 377 and 415 nm indicated that no antenna chromophore like MTHF is bound to the Mm0852 protein (now called MmCPDII). Figure 2.Spectroscopic characterization, repair activity and DNA binding. (A) UV/Vis spectrum of M. mazei photolyase (3.4 mg ml−1) shows fully oxidized FAD (a=362 nm; b=377 nm, c=421 nm; d=444 nm, e=469 nm). (B) UV/Vis spectra of photoreduction via illumination at 450 nm in the presence of reducing agent DTT. After addition of DTT (solid line), illumination leads to the neutral semiquinoid state of the photolyase (dashed line) with maxima at 590 nm (f) and 632 nm (g), respectively. The fully reduced state (dotted line) features a single maximum at 360 nm (h). (C) Complete repair of CPD lesions after onset of UV-A was achieved within 12 min. (D) Quantitative analysis of EMSA with M. mazei photolyase and CPD-dsDNA as well as undamaged DNA. The dashed curves correspond to non-linear fitting of the data to the simple Hill equation (Hill, 1910), whereas the solid line has been obtained by a mixed model for specific and non-specific DNA binding (see Supplementary data). (E) Scans of native EMSA-PAGE of IRDye700-labelled DNA probes incubated with M. mazei enzyme to determine dissociation constants. Observed photolyase•DNA complexes are marked with arrows. An additional distinct higher complex species is denoted with an asterisk. Download figure Download PowerPoint Like other photolyases MmCPDII is capable to photoreduce its catalytic FAD to the active FADH− form. Illumination at 450 nm of MmCPDII in presence of the reducing agent dithiothreitol (DTT) led to facile build-up of the semiquinoid, neutral state of the flavin cofactor (FADH•) by showing absorption peaks at 590 and 632 nm. During further illumination, the formation of fully reduced cofactor (FADH−) was accompanied by loss of absorption except at 360 nm (Figures 2B and 6C). Analogous intra-protein photoreductions of FAD have been described not only for various class I photolyases (Sancar, 2003) and A. thaliana cryptochrome 3 (Song et al, 2006), but also for plant class II photolyases (Okafuji et al, 2010). Catalytic activity of the recombinant MmCPDII as CPD photolyase was demonstrated by a modified repair assay using UV-damaged oligo(dT)18 (Jorns et al, 1985) and monitoring increased absorption at 265 nm due to repair of the CPD lesion upon illumination at 395 nm (Figure 2C). The MmCPDII enzyme showed unequivocal DNA-repair activity upon UV-A illumination as known from other photolyases. However, its DNA-binding characteristics differs significantly from class I photolyases. Electrophoretic mobility shift assays (EMSAs) using the experimental set-up established for A. thaliana cryptochrome 3 (Pokorny et al, 2008) were used to estimate the dissociation constants of MmCPDII to CPD-damaged and undamaged duplex DNA (Figure 2D and E). Undamaged 50mer DNA duplexes formed higher molecular mass complexes with MmCPDII at enzyme concentrations exceeding 200 nM. In contrast, CPD lesion comprising DNA duplexes showed a smaller and defined complex species already at 10 nM and above, which at higher concentrations is consumed by the large, non-specific DNA–enzyme complexes. Quantitation of the non-specific duplex DNA binding shows a dissociation constant of KD,NS=455±7 nM and a cooperativity that is consistent with the binding of four enzyme molecules per duplex DNA (n=3.9±0.2). Given the length of the oligonucleotides and the apparent lack of ladder-like complex intermediates this means that MmCPDII coaggregates along non-damaged duplex DNA and covers here about 12 base pairs or a stretch of 42 Å length. Non-specific binding by MmCPDII is much more prominent than in class I photolyases, where only high micromolar dissociation constants have been determined (EcCPDI: KD,NS=∼100 μM). Specific binding of UV-damaged duplex DNA is clearly stronger; a simple Hill analysis already gives a dissociation constant of KD,S=141±15 nM (n=1.4±0.2). Because the binding curve gives evidence between 100 and 400 nM of competing non-specific binding, a mixed model with specific 1:1 binding and cooperative non-specific binding appears to be more appropriate. This model gives specific and non-specific dissociation constants of KD,S=44±12 nM and KD,NS=305±27 nM (nD,NS=3.7±0.9), respectively. Compared with class I enzymes like EcCPDI (KD,S=∼1 nM; Sancar, 2003) specific DNA binding is by two orders of magnitude lower and the specificity ratios KD,NS/KD,S, ∼104 for EcCPDI, are reduced to a factor of 10 for MmCPDII. However, this is no shortcoming of class II photolyases, because the enzyme concentrations available in vivo can be expected to be too limiting to allow competing, cooperative non-specific binding to duplex DNA. Overall structure of M. mazei class II photolyase The crystal structure of the M. mazei class II photolyase at 1.5 Å resolution reveals the overall fold of the photolyase-cryptochrome family. Like class I (Park et al, 1995; Tamada et al, 1997; Komori et al, 2001; Fujihashi et al, 2007) and (6-4) photolyases (Maul et al, 2008; Hitomi et al, 2009) as well as cryptochromes (Brudler et al, 2003; Brautigam et al, 2004; Huang et al, 2006; Klar et al, 2007), the structure of MmCPDII is organized in an N-terminal α/β subdomain and a C-terminal all-helical subdomain (Figure 3A). The C-terminal FAD-binding subdomain (P232-Y461, α8-α19, 3104, 3105) contains the catalytic cofactor FAD in the U-shaped conformation commonly observed for photolyases and cryptochromes. The N-terminal subdomain (M3-S185) corresponds with its secondary structure elements α1-α6, β0-β5, 3101, 3102 to the Rossmann fold and lacks any kind of bound antenna chromophore. Figure 3.Overall structure of the M. mazei class II photolyase and structural comparison of members of the photolyase-cryptochrome family. (A) The N-terminal DNA-photolyase domain is shown in blue. C-terminal FAD-binding domain (green) contains the catalytic cofactor FAD (yellow) with SigmaA-weighted 2Fobs−Fcalc electron density contoured at 1σ. Domains are connected by a linker (grey) containing an α-helix (α7) with a 310-helical extension (3103). Not defined part of the linker by electron density is illustrated with a dashed line. Colouring of domain limits corresponds to Pfam database (Finn et al, 2010). (B) The M. mazei class II photolyase reveals a common overall fold to superimposed structures of the photolyase-cryptochrome family. Download figure Download PowerPoint Despite low pair-wise sequence identities to class I photolyases (Figure 1D), a comparison of MmCPDII with structures of the photolyase-cryptochrome family shows a highly preserved fold with overall r.m.s.d. of <2.5 Å for the superimposed Cα traces (Figure 3B). However, three features discriminate MmCPDII from other known members of the photolyase/cryptochrome family: the Rossmann-like fold harbours a sixth β-strand (β0) at the N-terminus; the catalytic subdomain is C-terminally truncated by ∼50 residues including the long C-terminal helix observed in class I photolyases (Figure 3B); and finally, the linker region connecting both subdomains (V186-E231) is significantly longer than in class I photolyases (EcCPDI: 25 aa, TtCPDI: 20 aa, AnCPD: 36 aa). Parts of the linker are apparently flexible as indicated by a lack of electron density for E189-V197 (Figure 3A). Such a lack of structural definition was also observed for the linker region of the class II photolyase from M. barkeri (E187-L226) whose structure was used for phasing of MmCPDII by molecular replacement (see Materials and methods and Supplementary Table SI). Overall, recombinant MmCPDII as used for crystallization and structure solution proved to be functional in photoreduction of the catalytic FAD cofactor, binding and repair of CPD lesions (Figure 2). Structure of the M. mazei photolyase•CPD–DNA complex To elucidate the recognition mode of CPD lesions within class II photolyases a stoichiometric complex between MmCPDII and a 14-meric DNA duplex with a synthetic CPD lesion in its central position has been crystallized under safe-light conditions (Mailliet et al, 2009). The MmCPDII•CPD–DNA structure at 2.2 Å resolution (Supplementary Figure S1A) reveals two complexes per asymmetric unit, where the dsDNA adopts a typical B-type conformation, but is severely kinked at the lesion site. The duplex at the 5′-side of the CPD lesion can be hence referred as 5′-arm, the other as 3′-arm. Both complexes are associated with each other by quasi-continuous arrangement of their bound DNA duplexes along the 3′-arms (Supplementary Figure S1B). Well-defined electron density for the DNA strand comprising the synthetic CPD lesion as well as for the counter strand is observed in both complexes with only a few nucleotides missing at the free termini of the 5′-arms (complex A: A1-T2; complex B: A1-C3 of the CPD strand and complex B: T14 of the counter strand, respectively). A comparison with the non-bound MmCPDII structure shows that complexation with dsDNA causes no major structural changes within the enzyme, as the r.m.s.d. between the uncomplexed photolyase and both MmCPDII•CPD–DNA complexes are rather low with 0.43 Å for complex A (396 Cα atoms) and 0.41 Å for complex B (352 Cα atoms), respectively. Similarly, only local structural changes within the enzyme upon CPD-DNA binding have previously been reported for the A. nidulans class I photolyase (Mees et al, 2004). Although both MmCPDII•CPD-DNA complexes are structurally similar, the 3′-arms of the duplex DNA remarkably exhibit deviating orientations of about 11° thus indicating plasticity in the recognition of dsDNA along the enzyme's surface (Supplementary Figure S1C). Nevertheless, as most of the protein–DNA interactions are preserved in both complexes, all following structural analyses used complex A unless otherwise stated. The CPD lesion of the kinked dsDNA is flipped out of the duplex into the active site of MmCPDII and bound there in a similar fashion as reported before for the complexes of the class I photolyase from A. nidulans (Mees et al, 2004) and the DASH cryptochrome from A. thaliana (Pokorny et al, 2008). However, the synthetic CPD lesion within the active site of MmCPDII is still intact with unbroken C5-C5 and C6-C6 bonds as indicated by unambiguous electron density in both complexes (Figure 4B) and differs in this regard from the CPD-DNA complexes reported for AnCPDI and Atcry3, where the CPD lesions have been repaired in situ by exposure to X-rays. The prevalence of an intact CPD lesion within the active site of MmCPDII is not caused by a general lack of photochemical reactivity of MmCPDII crystals. MmCPDII crystals readily undergo photoreduction at room temperature but not at cryogenic temperatures (Supplementary Figure S2E). Furthermore, MmCPDII crystals suffer from X-ray-induced reduction of the FAD chromophore (Supplementary Figure S2F), a behaviour that has been reported before for AnCPDI (Kort et al, 2004). Figure 4.Detailed view of the CPD lesion within the binding pocket of the M. mazei class II photolyase. (A) The schematic diagram illustrates that the class II photolyase is almost exclusively interacting with the CPD-comprising DNA strand. Dashed arrows indicate interactions with the protein backbone, whereas solid arrows characterize interactions with side chains. Black lines represent other mentioned interactions. Faded nucleotides are not defined by electron density. (B) Comparison of the active site of the MmCPDII•CPD–DNA complex (green) with AnCPDI (grey) bound to analogue CPD lesion (not shown). The class II enzyme features a cluster of water molecules (W1-W6) at the 3′-thymine replacing the corresponding asparagine residue (MmCPDII: G375, AnCPDI: N349). Electron density (SigmaA-weighted 2Fobs-Fcalc) is contoured at 1σ. Download figure Download PowerPoint Whereas the structures of AnCPDI and Atcry3 with in situ repaired CPD lesions show almost coplanar thymine bases with tilt angles of ∼13° (Mees et al, 2004; Pokorny et al, 2008), the cyclobutane ring of the CPD lesion within MmCPDII closely resembles with a tilt of 34° intact CPD lesions known from other structures. For example, in the crystal structure of the synthetic CPD lesion itself (Butenandt et al, 1998) the thymine bases adopt an interplanar angle of 57° (Supplementary Figure S3). Furthermore, in a set of four independent structures of the human polymerase η complexed to CPD–DNA (Biertümpfel et al, 2010), the interplanar angles of the CPD lesion vary position-dependent between 34° and 56° (Supplementary Figure S3). Apparently, the conformational plasticity of the CPD lesion is not causing a loss of the CB+ pucker of the cyclobutane ring upon binding to the active site of MmCPDII and only the class I and DASH-like photolyases are capable to select a CPD conformation that is already prone to breakage of the C5-C5 and C6-C6 bonds by X-ray-driven repair. In MmCPDII, the steric distortion induced by the active site onto the CPD lesion might be hence more relaxed due to the differences within the binding pocket of the thymine dimer including a unique water cluster (see below), although an influence of the crystallization matrix cannot be excluded. The C4 carbonyl groups of the 5′- and 3′-thymines of the CPD lesion in the MmCPDII structure form hydrogen bonds with the N6-amino group of the adenine from the catalytic FAD. This adenine is at least a crucial part of the docking site for the CPD lesion, if not even of the electron transfer pathway between the lesion and the isoalloxazine moiety. Theoretical calculations on electron transfer (Prytkova et al, 2007; Acocella et al, 2010) infer that the adenine drives the transfer to the CPD lesion from the 8-methyl group of FADH− rather indirectly by acting as an electrostatic ‘bouncer’ or as a structural organizer of the bound lesion. Accordingly, the hydrogen bonds between the thymines and the N6-amino group are maintained in the X-ray repaired states of the AnCPDI and Atcry3 CPD–DNA complexes, which correspond to structural snapshots after the first two nanoseconds of DNA repair. Despite the division into class I, class II and cryDASH-subfamilies for CPD photolyases the active sites share a high degree of structural homology as shown by a comparison between the CPD binding mode in MmCPDII with t" @default.
• W1844128776 created "2016-06-24" @default.
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• W1844128776 date "2011-09-02" @default.
• W1844128776 modified "2023-10-16" @default.
• W1844128776 title "Crystal structures of an archaeal class II DNA photolyase and its complex with UV-damaged duplex DNA" @default.
• W1844128776 cites W1544531597 @default.
• W1844128776 cites W1785791360 @default.
• W1844128776 cites W1941741329 @default.
• W1844128776 cites W1963677684 @default.
• W1844128776 cites W1965810021 @default.
• W1844128776 cites W1969672293 @default.
• W1844128776 cites W1969853783 @default.
• W1844128776 cites W1970109662 @default.
• W1844128776 cites W1970795846 @default.
• W1844128776 cites W197088108 @default.
• W1844128776 cites W1970901599 @default.
• W1844128776 cites W1980822294 @default.
• W1844128776 cites W1991593993 @default.
• W1844128776 cites W1998229535 @default.
• W1844128776 cites W1998840349 @default.
• W1844128776 cites W1999569890 @default.
• W1844128776 cites W2000840258 @default.
• W1844128776 cites W2001641653 @default.
• W1844128776 cites W2001998124 @default.
• W1844128776 cites W2002957935 @default.
• W1844128776 cites W2005307781 @default.
• W1844128776 cites W2009766773 @default.
• W1844128776 cites W2011204918 @default.
• W1844128776 cites W2016907186 @default.
• W1844128776 cites W2019306938 @default.
• W1844128776 cites W2020450410 @default.
• W1844128776 cites W2021012879 @default.
• W1844128776 cites W2029582401 @default.
• W1844128776 cites W2029677466 @default.
• W1844128776 cites W2031488163 @default.
• W1844128776 cites W2035503835 @default.
• W1844128776 cites W2038840577 @default.
• W1844128776 cites W2039950664 @default.
• W1844128776 cites W2042344660 @default.
• W1844128776 cites W2044143247 @default.
• W1844128776 cites W2044819346 @default.
• W1844128776 cites W2051248004 @default.
• W1844128776 cites W2065779555 @default.
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• W1844128776 cites W2089068314 @default.
• W1844128776 cites W2092946695 @default.
• W1844128776 cites W2101509122 @default.
• W1844128776 cites W2110671270 @default.
• W1844128776 cites W2113540334 @default.
• W1844128776 cites W2118244670 @default.
• W1844128776 cites W2124026197 @default.
• W1844128776 cites W2125502595 @default.
• W1844128776 cites W2133179696 @default.
• W1844128776 cites W2133752666 @default.
• W1844128776 cites W2156826426 @default.
• W1844128776 cites W2160986283 @default.
• W1844128776 cites W2163341755 @default.
• W1844128776 cites W2364878906 @default.
• W1844128776 cites W4210623056 @default.
• W1844128776 cites W4231612575 @default.
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