renaming BioC XML files to show PMC ID as file name and to be able to distinguish between raw and annotated XML; add folder with JSON version of annotated BioC

LICENSE.md

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annotated_BioC_JSON/PMC4774019_ann.json

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+[{"sourceid":"4774019","sourcedb":"","project":"","target":"","text":"The immunity-related GTPase Irga6 dimerizes in a parallel head-to-head fashion Background The immunity-related GTPases (IRGs) constitute a powerful cell-autonomous resistance system against several intracellular pathogens. Irga6 is a dynamin-like protein that oligomerizes at the parasitophorous vacuolar membrane (PVM) of Toxoplasma gondii leading to its vesiculation. Based on a previous biochemical analysis, it has been proposed that the GTPase domains of Irga6 dimerize in an antiparallel fashion during oligomerization. Results We determined the crystal structure of an oligomerization-impaired Irga6 mutant bound to a non-hydrolyzable GTP analog. Contrary to the previous model, the structure shows that the GTPase domains dimerize in a parallel fashion. The nucleotides in the center of the interface participate in dimerization by forming symmetric contacts with each other and with the switch I region of the opposing Irga6 molecule. The latter contact appears to activate GTP hydrolysis by stabilizing the position of the catalytic glutamate 106 in switch I close to the active site. Further dimerization contacts involve switch II, the G4 helix and the trans stabilizing loop. Conclusions The Irga6 structure features a parallel GTPase domain dimer, which appears to be a unifying feature of all dynamin and septin superfamily members. This study contributes important insights into the assembly and catalytic mechanisms of IRG proteins as prerequisite to understand their anti-microbial action. Electronic supplementary material The online version of this article (doi:10.1186/s12915-016-0236-7) contains supplementary material, which is available to authorized users. Background Immunity-related GTPases (IRGs) comprise a family of dynamin-related cell-autonomous resistance proteins targeting intracellular pathogens, such as Mycobacterium tuberculosis, Mycobacterium avium, Listeria monocytogenes, Trypanosoma cruzi, and Toxoplasma gondii. In mice, the 23 IRG members are induced by interferons, whereas the single human homologue is constitutively expressed in some tissues, especially in testis. In non-infected cells, most IRGs are largely cytosolic. However, members of a small sub-family with regulatory function associate with specific intracellular membranes, with one member favoring the endoplasmic reticulum and others the Golgi membrane and the endolysosomal system. Infection by certain intracellular pathogens initiates the redistribution of several effector members to the parasitophorous vacuole, followed by its disruption. In this way, IRGs contribute to the release of the pathogen into the cytoplasm and its subsequent destruction. Irga6, one of the effector IRG proteins, localizes to the intact parasitophorous vacuole membrane (PVM) and, after disruption of the PVM, is found associated with vesicular accumulations, presumably derived from the PVM. A myristoylation site at Gly2 is necessary for the recruitment to the PVM but not for the weak constitutive binding to the ER membrane. An internally oriented antibody epitope on helix A between positions 20 and 24 was demonstrated to be accessible in the GTP-, but not in the GDP-bound state. This indicates large-scale structural changes upon GTP binding that probably include exposure of the myristoyl group, enhancing binding to the PVM. Biochemical studies indicated that Irga6 hydrolyses GTP in a cooperative manner and forms GTP-dependent oligomers in vitro and in vivo. Crystal structures of Irga6 in various nucleotide-loaded states revealed the basic architecture of IRG proteins, including a GTPase domain and a composite helical domain. These studies additionally showed a dimerization interface in the nucleotide-free protein as well as in all nucleotide-bound states. It involves a GTPase domain surface, which is located at the opposite side of the nucleotide, and an interface in the helical domain, with a water-filled gap between the two contact surfaces. Mutagenesis of the contact surfaces suggests that this \"backside\" interface is not required for GTP-dependent oligomerization or cooperative hydrolysis, despite an earlier suggestion to the contrary. Extensive biochemical studies suggested that GTP-induced oligomerization of Irga6 requires an interface in the GTPase domain across the nucleotide-binding site. Recent structural studies indicated that a 'G interface' is typical of dynamin superfamily members, such as dynamin, MxA, the guanylate binding protein-1 (GBP-1), atlastin and the bacterial dynamin-like proteins (BDLP). For several of these proteins, formation of the G interface was shown to trigger GTP hydrolysis by inducing rearrangements of catalytic residues in cis. In dynamin, the G interface includes residues in the phosphate binding loop, the two switch regions, the 'trans stabilizing loop' and the 'G4 loop'. For Irga6, it was demonstrated that besides residues in the switch I and switch II regions, the 3'-OH group of the ribose participates in this interface. Since the signal recognition particle GTPase and its homologous receptor (called FfH and FtsY in bacteria) also employ the 3'-OH ribose group to dimerize in an anti-parallel orientation therefore activating its GTPase, an analogous dimerization model was proposed for Irga6. However, the crystal structure of Irga6 in the presence of the non-hydrolyzable GTP analogue 5'-guanylyl imidodiphosphate (GMPPNP) showed only subtle differences relative to the apo or GDP-bound protein and did not reveal a new dimer interface associated with the GTPase domain. This structure was obtained by soaking GMPPNP in nucleotide-free crystals of Irga6, an approach which may have interfered with nucleotide-induced domain rearrangements. To clarify the dimerization mode via the G interface, we determined the GMPPNP-bound crystal structure of a non-oligomerizing Irga6 variant. The structure revealed that Irga6 can dimerize via the G interface in a parallel head-to-head fashion. This dimerization mode explains previously published biochemical data, and shows in particular how the 3'-OH group of the ribose participates in the assembly. Our data suggest that a parallel dimerization mode may be a unifying feature in all dynamin and septin superfamily proteins. Results Previous results indicated that Irga6 mutations in a loosely defined surface region (the \"secondary patch\"), which is distant from the G-interface and only slightly overlapping with the backside interface (see below), individually reduced GTP-dependent oligomerization. A combination of four of these mutations (R31E, K32E, K176E, and K246E) essentially eliminated GTP-dependent assembly (Additional file 1: Figure S1) and allowed crystallization of Irga6 in the presence of GMPPNP. Crystals diffracted to 3.2 Å resolution and displayed one exceptionally long unit cell axis of 1289 Å (Additional file 1: Table S1). The structure was solved by molecular replacement and refined to Rwork/Rfree of 29.7 %/31.7 % (Additional file 1: Table S2). The asymmetric unit contained seven Irga6 molecules that were arranged in a helical pattern along the long cell axis (Additional file 1: Figure S2). Structure of the Irga6 dimer. a Schematic view of the domain architecture of mouse Irga6. The first and last amino acids of each domain are indicated. b Ribbon-type representation of the Irga6 dimer. In the left molecule, domains are colored according to the domain architecture, the right molecule is colored in grey. The nucleotide and Mg2+ ion (green) are shown in sphere representation. The GTPase domain dimer is boxed. The dotted line indicates a 2-fold axis. Secondary structure was numbered according to ref.. c\nTop view on the GTPase domain dimer. d Magnification of the contact sites. Dotted lines indicate interactions. e Superposition of different switch I conformations in the asymmetric unit; the same colors as in Additional file 1: Figure S2 are used for the switch I regions of the individual subunits. Switch I residues of subunit A (yellow) involved in ribose binding are labelled and shown in stick representation. Irga6 immunity-related GTPase 6 Like other dynamin superfamily members, the GTPase domain of Irga6 comprises a canonical GTPase domain fold, with a central β-sheet surrounded by helices on both sides (Fig. 1a-c). The helical domain is a bipartite structure composed of helices αA-C at the N-terminus and helix αF-L at the C-terminus of the GTPase domain. Overall, the seven molecules in the asymmetric unit are very similar to each other, with root mean square deviations (rmsd) ranging from 0.32 – 0.45 Å over all Cα atoms. The structures of the seven molecules also agree well with the previously determined structure of native GMPPNP-bound Irga6 (PDB: 1TQ6; rmsd of 1.00-1.13 Å over all Cα atoms). The seven Irga6 molecules in the asymmetric unit form various higher order contacts in the crystals. Within the asymmetric unit, six molecules dimerize via the symmetric backside dimer interface (buried surface area 930 Å2), and the remaining seventh molecule forms the same type of interaction with its symmetry mate of the adjacent asymmetric unit (Additional file 1: Figure S2a, b, Figure S3). This indicates that the introduced mutations in the secondary patch, from which only Lys176 is part of the backside interface, do, in fact, not prevent this interaction. Another assembly interface with a buried surface area of 450 Å2, which we call the “tertiary patch”, was formed via two interaction sites in the helical domains (Additional file 1: Figure S2c, d, S3). In this interface, helices αK from two adjacent molecules form a hydrogen bonding network involving residues 373-376. Furthermore, two adjacent helices αA form hydrophobic contacts. It was previously shown that the double mutation L372R/A373R did not prevent GTP-induced assembly, so there is currently no evidence supporting an involvement of this interface in higher-order oligomerization. Strikingly, molecule A of one asymmetric unit assembled with an equivalent molecule of the adjacent asymmetric unit via the G-interface in a symmetric parallel fashion via a 470 Å2 interface. This assembly results in a butterfly-shaped Irga6 dimer in which the helical domains protrude in parallel orientations (Fig. 1b, Additional file 1: Figure S3). In contrast, the other six molecules in the asymmetric unit do not assemble via the G interface. The G interface in molecule A can be subdivided into three distinct contact sites (Fig. 1c, d). Contact site I is formed between R159 and K161 in the trans stabilizing loops, and S132 in the switch II regions of the opposing molecules. Contact site II features polar and hydrophobic interactions formed by switch I (V104, V107) with a helix following the guanine specificity motif (G4 helix, K184 and S187) and the trans stabilizing loop (T158) of the opposing GTPase domain. In contact site III, G103 of switch I interacts via its main chain nitrogen with the exocyclic 2’-OH and 3’-OH groups of the opposing ribose in trans, whereas the two opposing exocyclic 3’-OH group of the ribose form hydrogen bonds with each other. Via the ribose contact, switch I is pulled towards the opposing nucleotide (Fig. 1e). In turn, E106 of switch I reorients towards the nucleotide and now participates in the coordination of the Mg2+ ion (Fig. 1e, Additional file 1: Figure S4). E106 was previously shown to be essential for catalysis, and the observed interactions in contact site III explain how dimerization via the ribose is directly coupled to the activation of GTP hydrolysis. The G interface is in full agreement with previously published biochemical data that indicate crucial roles of E77, G103, E106, S132, R159, K161, K162, D164, N191, and K196 for oligomerization and oligomerization-induced GTP hydrolysis. All of these residues directly participate in contacts (G103, S132, R159, and K161) or are in direct vicinity to the interface (E77, E106, K162, D164, and N191). Residues E77, K162, and D164 appear to orient the trans stabilizing loop which is involved in interface formation in contact site II. In the earlier model of an anti-parallel G interface, it was not possible to position the side chain of R159 to avoid steric conflict. In the present structure, the side-chain of R159 projects laterally along the G interface and, therefore, does not cause a steric conflict. A conserved dimerization mode via the G interface in dynamin and septin GTPases. The overall architecture of the parallel GTPase domain dimer of Irga6 is related to that of other dynamin and septin superfamily proteins. The following structures are shown in cylinder representations, in similar orientations of their GTPase domains: a the GMPPNP-bound Irga6 dimer, b the GDP-AlF4\n--bound dynamin 1 GTPase-minimal BSE construct [pdb 2X2E], c the GDP-bound atlastin 1 dimer [pdb 3Q5E], d the GDP-AlF3- bound GBP1 GTPase domain dimer [pdb 2B92], e the BDLP dimer bound to GDP [pdb 2J68] and f the GTP-bound GIMAP2 dimer [pdb 2XTN]. The GTPase domains of the left molecules are shown in orange, helical domains or extensions in blue. Nucleotide, Mg2+ (green) and AlF4\n- are shown in sphere representation, the buried interface sizes per molecule are indicated on the right. Irga6 immunity-related GTPase 6, GMPPNP 5'-guanylyl imidodiphosphate, GTP guanosine-triphosphate, BDLP bacterial dynamin like protein, GIMAP2, GTPase of immunity associated protein 2 The buried surface area per molecule (BSA) of the G interface in Irga6 is relatively small (470 Å2) compared to that of other dynamin superfamily members, such as dynamin (BSA: 1400 Å2), atlastin (BSA: 820 Å2), GBP-1 (BSA: 2060 Å2), BDLP (BSA: 2300 Å2) or the septin-related GTPase of immunity associated protein 2 (GIMAP2) (BSA: 590 Å2) (Fig. 2). However, the relative orientations of the GTPase domains in these dimers are strikingly similar, and the same elements, such as switch I, switch II, the trans activating and G4 loops are involved in the parallel dimerization mode in all of these GTPase families. Discussion IRG proteins are crucial mediators of the innate immune response in mice against a specific subset of intracellular pathogens, all of which enter the cell to form a membrane-bounded vacuole without engagement of the phagocytic machinery. As members of the dynamin superfamily, IRGs oligomerize at cellular membranes in response to GTP binding. Oligomerization and oligomerization-induced GTP hydrolysis are thought to induce membrane remodeling events ultimately leading to disruption of the PVM. Recent structural and mechanistic analyses have begun to unravel the molecular basis for the membrane-remodeling activity and mechano-chemical function of some members (reviewed in). For example, for dynamin and atlastin, it was shown that GTP binding and/or hydrolysis leads to dimerization of the GTPase domains and to the reorientation of the adjacent helical domains. The resulting domain movement was suggested to act as a “power stroke” during membrane remodeling events. However, for other dynamin superfamily members such as IRGs, the molecular basis for GTP hydrolysis and the exact role of the mechano-chemical function are still unclear. Our structural analysis of an oligomerization- and GTPase-defective Irga6 mutant indicates that Irga6 dimerizes via the G interface in a parallel orientation. Only one of the seven Irga6 molecules in the asymmetric unit formed this contact pointing to a low affinity interaction via the G interface, which is in agreement with its small size. In the crystals, dimerization via the G interface is promoted by the high protein concentrations which may mimic a situation when Irga6 oligomerizes on a membrane surface. Such a low affinity interaction mode may allow reversibility of oligomerization following GTP hydrolysis. Similar low affinity G interface interactions were reported for dynamin and MxA. The dimerization mode is strikingly different from the previously proposed anti-parallel model that was based on the crystal structure of the signal recognition particle GTPase, SRP54 and its homologous receptor. However, the G dimer interface is reminiscent of the GTPase domain dimers observed for several other dynamin superfamily members, such as dynamin, GBP1, atlastin, and BDLP. It was recently shown that septin and septin-related GTPases, such as the Tocs GTPases or GTPases of immunity related proteins (GIMAPs), also employ a GTP-dependent parallel dimerization mode. Based on phylogenetic and structural analysis, these observations suggest that dynamin and septin superfamilies are derived from a common ancestral membrane-associated GTPase that featured a GTP-dependent parallel dimerization mode. Importantly, our analysis indicates that IRGs are not outliers, but bona-fide representatives of the dynamin superfamily. Whereas the overall dimerization mode is similar in septin and dynamin GTPases, family-specific differences in the G interface and the oligomerization interfaces exist. For example, the involvement of the 2’ and 3’-OH groups of the ribose in the dimerization interface of Irga6 has not been observed for other dynamin and septin superfamily members. The surface-exposed location of the ribose in the Irga6 structure, with a wide-open nucleotide-binding pocket, facilitates its engagement in the dimerization interface. This contact, in turn, appears to activate GTP hydrolysis by inducing rearrangements in switch I and the positioning of the catalytic E106. During dimerization of GBP1, an arginine finger from the P loop reorients towards the nucleotide in cis to trigger GTP hydrolysis. In dynamin, the corresponding serine residue coordinates a sodium ion that is crucial for GTP hydrolysis. Irga6 bears Gly79 at this position, which in the dimerizing molecule A appears to approach the bridging imido group of GMPPNP via a main chain hydrogen bond. Higher resolution structures of the Irga6 dimer in the presence of a transition state analogue are required to show whether Gly79 directly participates in GTP hydrolysis or whether it may also position a catalytic cation. In dynamin, further assembly sites are provided by the helical domains which assemble in a criss-cross fashion to form a helical filament. In dynamin-related Eps15 homology domain containing proteins (EHDs), a second assembly interface is present in the GTPase domain. For Irga6, additional interfaces in the helical domain are presumably involved in oligomerization, such as the secondary patch residues whose mutation prevented oligomerization in the crystallized mutant. Further structural studies, especially electron microscopy analysis of the Irga6 oligomers, are required to clarify the assembly mode via the helical domains and to show how these interfaces cooperate with the G interface to mediate the regulated assembly on a membrane surface. Notably, we did not observe major rearrangements of the helical domain versus the GTPase domain in the Irga6 molecules that dimerized via the G interface. In a manner similar to BDLP, such large-scale conformational changes may be induced by membrane binding. Our structural analysis and the identification of the G-interface paves the way for determining the specific assembly of Irga6 into a membrane-associated scaffold as the prerequisite to understand its action as an anti-parasitic machine. Methods Protein expression and purification Selenomethionine-substituted Mus musculus Irga6R31E, K32E, K176E, K246E was expressed as a GST-fusion from the vector pGEX-4T-2 in BL21 Rosetta2(DE3) cells according to reference. Protein was purified as previously described and the protein stored in small aliquots at a concentration of 118 mg/mL in 50 mM Tris-HCl, pH 7.4, 5 mM MgCl2, 2 mM DTT. Biochemical analyses Oligomerization and GTPase assays for the Irga6R31E, K32E, K176E, K246E mutant were carried out as described in. Protein crystallization The protein was gently thawed on ice and diluted to a final concentration of 10 mg/mL with buffer containing 20 mM Tris-HCl, pH 7.5, 8 mM MgCl2, 3 mM DTT. GMPPNP was added to a final concentration of 2 mM. Crystallization was carried out in a 96 well format using the sitting drop vapor diffusion method. The reservoir contained 100 mM HEPES-NaOH pH 7.0, 9 % PEG4000, 6 % isopropanol. The sitting drop was set up using an Art Robbins Gryphon system and consisted of 200 nL protein solution and 200 nL reservoir solution. For cryo-protection, crystals were transferred into a cryo solution containing 33 % PEG4000, 3 % isopropanol, 50 mM HEPES pH 7.0, 4 mM MgCl2, 2 mM DTT, and 2 mM GMPPNP at 4 °C for at least 5 sec. Crystals were screened for diffraction at beamline BL 14.1 at BESSY II, Berlin, Germany. Data collection All data were recorded at beamline P11 at PETRA III, DESY Hamburg, Germany using a PILATUS 6 M detector. To achieve spot separation along the long cell axis, three data sets were collected with a φ increment of 0.05/0.1° at a temperature of 100 K using detector distances between 1300 and 598.5 mm (Additional file 1: Table S1). The wavelength was 0.972/0.979 Å. Calculation of an optimal data collection strategy was done with the Mosflm software. The high- and low-resolution datasets were processed and merged using the XDS program suite. Structure solution and refinement Structure solution was done by molecular replacement with Phaser employing the structure of Irga6 without nucleotide [PDB: 1TQ2] as search model. Atomic model building was done by Coot. Iterative refinement was done using Phenix at a maximum resolution of 3.2 Å. For the refinement strategy, a seven-fold non-crystallographic symmetry as well as one molecule of Irga6 [PDB: 1TQ4] as high resolution reference structure was chosen. Five percent of the measured X-ray intensities were set aside from the refinement as cross-validation. Methionine sites in the protein were confirmed by the anomalous signal of the selenium atoms. Protein superposition was done with lsqkab and the PyMol Molecular Graphics System, Version 1.3 Schrödinger, LLC. Figures were prepared using the PyMOL Molecular Graphics System, Version 1.7.4 Schrödinger, LLC. Evaluation of atom contacts and geometry of the atomic model was done by the Molprobity server. Interface sizes were calculated by the PISA server. Accession numbers The Irga6 coordinates were submitted to the Protein Data Bank (pdb) database with accession code 5fph. http://www.rcsb.org/pdb/explore/explore.do?structureId=5fph. Conclusions Our study indicates that Irg proteins dimerize via the G interface in a parallel head-to-head fashion thereby facilitating GTPase activation. These findings contribute to a molecular understanding of the anti-parasitic action of the Irg protein family and suggest that Irgs are bona-fide members of the dynamin superfamily. Additional file Abbreviations BDLP Bacterial dynamin like protein EHD2 Eps15 homology domain containing protein 2 GBP Guanylate-binding protein GDP Guanosine-diphosphate GIMAP2 GTPase of immunity associated protein 2 GMPPNP 5'-guanylyl imidodiphosphate GTP Guanosine-triphosphate IRG Immunity-related GTPase Irga6 Immunity-related GTPase 6 MxA Myxovirus resistance protein A PVM Parasitophorous vacuolar membrane Competing interests The authors declare that they have no competing interests. Authors’ contributions All authors planned the experimental design. NP cloned, characterized, and purified the Irga6 construct and found initial crystallization conditions. KS and CF optimized the crystallization condition and found suitable cryo conditions. KS and KF collected data, KF solved, and KS and KF refined the structure. KS, NP, CF, JCH, and OD wrote the manuscript. All authors read and approved the final manuscript. References Immune control of tuberculosis by IFN-gamma-inducible LRG-47 Mice deficient in LRG-47 display increased susceptibility to mycobacterial infection associated with the induction of lymphopenia Inactivation of LRG-47 and IRG-47 reveals a family of interferon gamma-inducible genes with essential, pathogen-specific roles in resistance to infection Mice deficient in LRG-47 display enhanced susceptibility to Trypanosoma cruzi infection associated with defective hemopoiesis and intracellular control of parasite growth Pathogen-specific loss of host resistance in mice lacking the IFN-gamma-inducible gene IGTP p47 GTPases regulate Toxoplasma gondii survival in activated macrophages Disruption of Toxoplasma gondii parasitophorous vacuoles by the mouse p47-resistance GTPases Balance of Irgm protein activities determines IFN-gamma-induced host defense p47 GTPases: regulators of immunity to intracellular pathogens The interferon-inducible GTPases The immunity-related GTPases in mammals: a fast-evolving cell-autonomous resistance system against intracellular pathogens Death and resurrection of the human IRGM gene The inducibly expressed GTPase localizes to the endoplasmic reticulum, independently of GTP binding Mechanisms regulating the positioning of mouse p47 resistance GTPases LRG-47 and IIGP1 on cellular membranes: retargeting to plasma membrane induced by phagocytosis Disruption of the Toxoplasma gondii parasitophorous vacuole by IFNgamma-inducible immunity-related GTPases (IRG proteins) triggers necrotic cell death Coordinated loading of IRG resistance GTPases on to the Toxoplasma gondii parasitophorous vacuole Toxoplasma's arms race with the host interferon response: a menage a trois of ROPs Vacuolar and plasma membrane stripping and autophagic elimination of Toxoplasma gondii in primed effector macrophages The gamma interferon (IFN-gamma)-inducible GTP-binding protein IGTP is necessary for toxoplasma vacuolar disruption and induces parasite egression in 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guanylate-binding proteins achieve assembly-stimulated processive cleavage of GTP to GMP Structural basis for the nucleotide-dependent dimerization of the large G protein atlastin-1/SPG3A Structures of the atlastin GTPase provide insight into homotypic fusion of endoplasmic reticulum membranes A bacterial dynamin-like protein Structure of a bacterial dynamin-like protein lipid tube provides a mechanism for assembly and membrane curving Substrate twinning activates the signal recognition particle and its receptor Oligomerization of dynamin superfamily proteins in health and disease Structural insight into filament formation by mammalian septins Crystal structure of pea Toc34, a novel GTPase of the chloroplast protein translocon Structural basis of oligomerization in septin-like GTPase of immunity-associated protein 2 (GIMAP2) Architectural and mechanistic insights into an EHD ATPase involved in membrane remodelling Atomic structures of the human immunophilin FKBP-12 complexes with FK506 and rapamycin The integration of macromolecular diffraction data XDS Phaser crystallographic software Features and development of Coot PHENIX: a comprehensive Python-based system for macromolecular structure solution Free R, value: cross-validation in crystallography A solution for the best rotation to relate two sets of vectors MolProbity: all-atom structure validation for macromolecular crystallography Inference of macromolecular assemblies from crystalline 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annotated_BioC_JSON/PMC4781976_ann.json

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+[{"sourceid":"4781976","sourcedb":"","project":"","target":"","text":"Structure of the GAT domain of the endosomal adapter protein Tom1 Cellular homeostasis requires correct delivery of cell-surface receptor proteins (cargo) to their target subcellular compartments. The adapter proteins Tom1 and Tollip are involved in sorting of ubiquitinated cargo in endosomal compartments. Recruitment of Tom1 to the endosomal compartments is mediated by its GAT domain’s association to Tollip’s Tom1-binding domain (TBD). In this data article, we report the solution NMR-derived structure of the Tom1 GAT domain. The estimated protein structure exhibits a bundle of three helical elements. We compare the Tom1 GAT structure with those structures corresponding to the Tollip TBD- and ubiquitin-bound states. Specifications table Table\t \t Subject area\tBiology\t \tMore specific subject area\tStructural biology\t \tType of data\tTable, text file, graph, figures\t \tHow data was acquired\tCircular dichroism and NMR. NMR data was recorded using a Bruker 800 MHz\t \tData format\tPDB format text file. Analyzed by CS-Rosetta, Protein Structure Validation Server (PSVS), NMRPipe, NMRDraw, and PyMol\t \tExperimental factors\tRecombinant human Tom1 GAT domain was purified to homogeneity before use\t \tExperimental features\tSolution structure of Tom1 GAT was determined from NMR chemical shift data\t \tData source location\tVirginia and Colorado, United States.\t \tData accessibility\tData is available within this article. Tom1 GAT structural data is publicly available in the RCSB Protein Data Bank (http://www.rscb.org/) under the accession number PDB: 2n9d\t \t Value of the data The Tom1 GAT domain solution structure will provide additional tools for modulating its biological function. Tom1 GAT can adopt distinct conformations upon ligand binding. A conformational response of the Tom1 GAT domain upon Tollip TBD binding can serve as an example to explain mutually exclusive ligand binding events. Data Analysis of the far-UV circular dichroism (CD) spectrum of the Tom 1 GAT domain (Fig. 1) predicts 58.7% α-helix, 3% β-strand, 15.5% turn, and 22.8% disordered regions. The Tom1 GAT structural restraints yielded ten helical structures (Fig. 2A,B) with a root mean square deviation (RMSD) of 0.9 Å for backbone and 1.3 Å for all heavy atoms (Table 1) and estimated the presence of three helices spanning residues Q216-E240 (α-helix 1), P248-Q274 (α-helix 2), and E278-T306 (α-helix 3). Unlike ubiquitin binding, data suggest that conformational changes of the Tom1 GAT α-helices 1 and 2 occur upon Tollip TBD binding (Fig. 3A,B). Experimental design, materials, and methods Protein expression and purification Human Tom1 GAT (residues 215–309) cDNA was cloned into both pGEX6P1 and pET28a vectors, and expressed as GST-tagged and His-tagged fusion proteins, respectively, using Escherichia coli [Rosetta (DE3) strain]. The 13C, 15N-labeled Tom1 GAT domain was expressed and purified as described previously. Circular dichroism Far-UV CD spectra of the His-Tom1 GAT domain were collected on a Jasco J-815 spectropolarimeter using a 1 mm path length quartz cell at room temperature. The protein (10 μM) was solubilized in 5 mM Tris–HCl (pH 7) and 100 mM KF. Spectra were obtained from five accumulated scans from 190 to 260 nm using a bandwidth of 1 nm and a response time of 1 s at a scan speed of 20 nm/min. Buffer backgrounds were employed to subtract the protein spectra. Data was processed using the Dichroweb server and the CONTIN algorithm (http://dichroweb.cryst.bbk.ac.uk/html/home.shtml). NMR structure determination NMR experiments were performed using 1 mM 13C, 15N-labeled Tom1 GAT domain in a buffer containing 20 mM d11-TrisHCl (pH 7), 50 mM KCl, 1 mM d18-DTT, and 1 mM NaN3. NMR spectra were recorded at 25 °C on a Bruker 800-MHz spectrometer (University of Virginia). The individual structure of Tom1 GAT was generated using CS-Rosetta (https://csrosetta.bmrb.wisc.edu/csrosetta). Chemical shift information (BMRB #26574) was used to obtain the structure calculation. The Rosetta calculations yielded 3000 structures of Tom1 GAT. From these, ten structures were selected based on their score and RMSDs, and converted to Protein Data Bank (PDB) format. NMR structural statistics for the ten lowest energy conformers of Tom1 GAT was generated using the Protein Structure Validation Suite. By using MolProbity, the Ramachandran analysis of the ten superimposed Tom1 GAT structures identified that 100% of the residues were in the most favored regions and there were no Ramachandran outliers in the allowed and disallowed regions. Protein structure images were obtained using PyMol (http://www.pymol.org). The structures of the ubiquitin- and Tollip TBD-bound states of the Tom1 GAT domain were obtained from data reported in Refs. and. References Tom1 modulates binding of Tollip to phosphatidylinositol 3-phosphate via a coupled folding and binding mechanism Structural basis for recognition of ubiquitinated cargo by Tom1-GAT domain Supplementary material Supplementary data associated with this article can be found in the online version at doi:10.1016/j.dib.2016.02.042. Representative far-UV CD spectrum of the His-Tom1 GAT domain. Fig. 1. (A) Stereo view displaying the best-fit backbone superposition of the refined structures for the Tom1 GAT domain. Helices are shown in orange, whereas loops are colored in green. (B) Ribbon illustration of the Tom1 GAT domain. Fig. 2. (A) Two views of the superimposed structures of the Tom1 GAT domain in the free state (gray) with that in the Tollip TBD-bound state (red). (B) Two views of the superimposed structures of the Tom1 GAT domain (gray) with that in the Ub-bound state (green). Fig. 3. NMR and refinement statistics for the Tom1 GAT domain. NMR structural statistics for lowest energy conformers of Tom1 GAT using PSVS. Table 1.\t \t \tTom1 GAT\t \tNMR distance and dihedral constraints\t\t \t Dihedral angle restraints total\t178\t \t ϕ\t89\t \t ψ\t89\t \tStructure statistics\t\t \t Dihedral angle constraints (deg)\t8.8±0.2\t \t Max. dihedral angle violation (deg)\t111±3\t \tDeviations from idealized geometry\t\t \t Bond lengths (Å)\t0.011\t \t Bond angles (deg)\t0.7\t \tAverage pairwise r.m.s. deviation (Å)a\t\t \t Protein\t\t \t Heavy\t1.3\t \t Backbone\t0.9\t \t Pairwise backbone and heavy-atom r.m.s. deviations were obtained by superimposing residues 215–309 of Tom1 GAT among 10 lowest energy refined 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annotated_BioC_JSON/PMC4822561_ann.json

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+[{"sourceid":"4822561","sourcedb":"","project":"","target":"","text":"Structure of a quinolone-stabilized cleavage complex of topoisomerase IV from Klebsiella pneumoniae and comparison with a related Streptococcus pneumoniae complex Crystal structures of the cleavage complexes of topoisomerase IV from Gram-negative (K. pneumoniae) and Gram-positive (S. pneumoniae) bacterial pathogens stabilized by the clinically important antibacterial drug levofloxacin are presented, analysed and compared. For K. pneumoniae, this is the first high-resolution cleavage complex structure to be reported.  Klebsiella pneumoniae is a Gram-negative bacterium that is responsible for a range of common infections, including pulmonary pneumonia, bloodstream infections and meningitis. Certain strains of Klebsiella have become highly resistant to antibiotics. Despite the vast amount of research carried out on this class of bacteria, the molecular structure of its topoisomerase IV, a type II topoisomerase essential for catalysing chromosomal segregation, had remained unknown. In this paper, the structure of its DNA-cleavage complex is reported at 3.35 Å resolution. The complex is comprised of ParC breakage-reunion and ParE TOPRIM domains of K. pneumoniae topoisomerase IV with DNA stabilized by levofloxacin, a broad-spectrum fluoroquinolone antimicrobial agent. This complex is compared with a similar complex from Streptococcus pneumoniae, which has recently been solved. Introduction     Klebsiella is a genus belonging to the Enterobacteriaceae family of Gram-negative bacilli, which is divided into seven species with demonstrated similarities in DNA homology: K. pneumoniae, K. ozaenae, K. rhinoscleromatis, K. oxytoca, K. planticola, K. terrigena and K. ornithinolytica. K. pneumoniae is the most medically important species of the genus owing to its high resistance to antibiotics. Significant morbidity and mortality has been associated with an emerging, highly drug-resistant strain of K. pneumoniae characterized as producing the carbapenemase enzyme (KPC-producing bacteria; Nordmann et al., 2009). The best therapeutic approach to KPC-producing organisms has yet to be defined. However, common treatments (based on in vitro susceptibility testing) are the polymyxins, tigecycline and, less frequently, aminoglycoside antibiotics (Arnold et al., 2011). Another effective strategy involves the limited use of certain antimicrobials, specifically fluoroquinolones and cephalo­sporins (Gasink et al., 2009). Several new antibiotics are under development for KPC producers. These include combinations of existing β-lactam antibiotics with new β-lactamase inhibitors able to circumvent KPC resistance. Neoglycosides are novel aminoglycosides that have activity against KPC-producing bacteria that are also being developed (Chen et al., 2012). Type II topoisomerase enzymes play important roles in prokaryotic and eukaryotic DNA replication, recombination and transcription (Drlica et al., 2008; Laponogov et al., 2013; Lee et al., 2013; Nitiss, 2009a ,b ; Schoeffler \u0026 Berger, 2008; Sissi \u0026 Palumbo, 2009; Vos et al., 2011; Wendorff et al., 2012; Wu et al., 2011, 2013). In bacteria, topoisomerase IV, a tetramer of two ParC and two ParE subunits, unlinks daughter chromosomes prior to cell division, whereas the related enzyme gyrase, a GyrA2GyrB2 tetramer, supercoils DNA and helps unwind DNA at replication forks. Both enzymes act via a double-strand DNA break involving a cleavage complex and are targets for quinolone antimicrobials that act by trapping these enzymes at the DNA-cleavage stage and preventing strand re-joining (Drlica et al., 2008). Levofloxacin is a broad-spectrum third-generation fluoro­quinolone antibiotic. It is active against Gram-positive and Gram-negative bacteria and functions by inhibiting gyrase and topoisomerase IV (Drlica \u0026 Zhao, 1997; Laponogov et al., 2010). Acquiring a deep structural and functional understanding of the mode of action of fluoroquinolones (Tomašić \u0026 Mašič, 2014) and the development of new drugs targeted against topoisomerase IV and gyrase from a wide range of Gram-positive and Gram-negative pathogenic bacteria are highly active areas of current research directed at overcoming the vexed problem of drug resistance (Bax et al., 2010; Chan et al., 2015; Drlica et al., 2014; Mutsaev et al., 2014; Pommier, 2013; Srikannathasan et al., 2015). Here, we report the first three-dimensional X-ray structure of a K. pneumoniae topoisomerase IV ParC/ParE cleavage complex with DNA stabilized by levofloxacin. The crystal structure provides structural information on topoisomerase IV from K. pneumoniae, a pathogen for which drug resistance is a serious concern. The structure of the ParC/ParE–DNA–levofloxacin binding site highlights the details of the cleavage-complex assembly that are essential for the rational design of Klebsiella topoisomerase inhibitors. Materials and methods    Cloning, expression and purification of K. pneumoniae and Streptococcus pneumoniae ParC55/ParE30    Cloning, expression and purification protocols are described in detail in the Supporting Information. Table 1 ▸ contains the sequence information for all of the components of the complexes. Fig. 1 ▸ provides information about the protein and DNA constructs used in the experimental work. Preparation of the DNA oligomer    For the K. pneumoniae cleavage complex, two DNA oligomers (5′-CGTATTACGTTGTAT-3′ and 5′-GATCATACAACGTAATACG-3′) were synthesized by solid-phase phosphoramidite chemistry and doubly HPLC purified by Metabion, Munich, Germany. The DNA sequence was designed to make a complementary DNA 34-mer that contained the ‘pre-cut’ binding-site fragment: 5′-CGTATTACGTTGTAT↓GATCATACAACGTAATACG-3′ and 3′-GCATAATGCA­ACATACTAG↓TATGTTGCATTATGC-5′ (the cuts are shown by arrows; see Fig. 1 ▸ b). For the S. pneumoniae cleavage complex, two DNA oligomers (5′-CATGAATGACTATGCACG-3′ and 5′-CGTGCATAGTCATTCATG-3′) were synthesized by solid-phase phosphoramidite chemistry and doubly HPLC purified by Metabion, Munich, Germany. The DNA sequence corresponds to the E-site 18-mer, which was found to be a better DNA length for crystallization of the S. pneumoniae topo­isomerase IV cleavage complexes in order to give stable reproducible crystals (see Fig. 1 ▸ b). DNA stock solutions were made by mixing the required oligomers (at 1 mM in 20 mM Tris pH 7.5, 200 mM NaCl, 1 mM β-mercaptothanol, 0.05% NaN3) in equal volumes. For DNA annealing, the mixtures of complementary oligomers were heated to 98°C and then slowly cooled to 4°C over a 48 h period. Crystallization and data collection    Crystallization information for both the S. pneumoniae and the K. pneumoniae topoisomerase IV cleavage complexes is summarized in Table 2 ▸. Data-collection statistics and details are provided in Table 3 ▸. Structure-solution and refinement details are provided in Table 4 ▸.  S. pneumoniae topoisomerase IV    Protein was mixed with DNA in a 1:1:1.2 molar ratio (ParC55:ParE30:18-mer E-site DNA) with an overall concentration of 4 mg ml−1. Levofloxacin and magnesium chloride were added to final concentrations of 2 and 10 mM, respectively. The mixture was pre-incubated at room temperature overnight. Initial crystallization screening was performed by sitting-drop vapour diffusion in a 96-well MRC crystallization plate (600 nl protein mixture + 400 nl reservoir solution) using a Mosquito robot (TTP Labtech; http://www.ttplabtech.com). The best crystals were obtained using capillary counter-diffusion against 50 mM sodium cacodylate pH 6.5, 2.5% Tacsimate (Hampton Research; McPherson \u0026 Cudney, 2006), 7% 2-propanol, 62.5 mM KCl, 7.5 mM MgCl2 at 304 K. The crystals were flash-cooled at 100 K in cryoprotectant buffer C [50 mM sodium cacodyl­ate pH 6.5, 2.5% Tacsimate, 62.5 mM KCl, 7.5 mM MgCl2, 1 mM β-mercapto­ethanol, 30%(v/v) MPD]. The best data set was collected on beamline I03 at Diamond Light Source at a wavelength of 0.9763 Å using an ADSC Quantum 315 detector. The data extended to 2.6 Å resolution anisotropically and were used in refinement with a maximum-likelihood target in the initial refinement cycles; they were deposited in the PDB without introducing a resolution cutoff. However, owing to the high R merge values in the outer shells, the final resolution is given as 2.9 Å and the statistics are reported according to this ‘trimmed’ resolution. The resolution cutoff was based on the rejection criteria R merge \u003c 50% and I/σ(I) \u003e 1.5 in the highest resolution shell. The data were integrated using HKL-2000 (Otwinowski \u0026 Minor, 1997). The space group was determined to be P3121, with unit-cell parameters a = b = 157.83, c = 211.15 Å. The structure was solved by molecular replacement using Phaser (McCoy et al., 2007) as implemented within the CCP4 suite (Winn et al., 2011) and our previously published topo­isomerase IV–levofloxacin structure (PDB entry 3k9f; Laponogov et al., 2010). Refinement was performed in PHENIX (Adams et al., 2002, 2010) with manual inspection and corrections performed in WinCoot (Emsley \u0026 Cowtan, 2004; Emsley et al., 2010).The structure was verified using WinCoot and PROCHECK (Laskowski et al., 1993).  K. pneumoniae topoisomerase IV    ParC55/ParE30 protein stock in incubation buffer (at 4.5 mg ml−1) was mixed with the ‘pre-cut’ 34-mer DNA stock in a 1:1.2 protein:DNA molar ratio. High-concentration stocks of levofloxacin and MgCl2 were added to give final concentrations of 2 and 10 mM, respectively. The mixture was incubated overnight at room temperature. Initial crystallization screening was performed by sitting-drop vapour diffusion in 96-well MRC crystallization plates (600 nl protein mixture + 300 nl reservoir solution) using a Mosquito robot. When the optimal crystallization conditions had been established, conventional hanging-drop vapour diffusion in 24-well Linbro plates (4 µl protein mixture + 2 µl reservoir solution) was used to increase the crystal size. Crystals formed after ∼7–10 d at room temperature. The crystallization conditions varied slightly from batch to batch in the range 0.1 M Tris pH 7.5–8.0, 0–50 mM NaCl, 4–8% PEG 4000, 12–15% glycerol. It should be mentioned that several other DNA oligomers with the same binding-site sequence were tried for crystallization (i.e. 20-mer, ‘pre-cut’ 20-mer and 34-mer DNA sequences). However, these protein–DNA–drug complexes did not produce good-quality crystals for data collection. Crystals were tested in-house for diffraction quality using an Oxford Xcalibur Nova CCD diffractometer and were then transported for high-resolution data collection at Diamond Light Source (Harwell Science and Innovation Campus, Oxfordshire, England). The data were collected on beamline I03 (wavelength 0.9762 Å) using a Pilatus 6M-F detector (0.2° oscillation per image, 100 K nitrogen stream). The best crystals diffracted to ∼3.2 Å resolution. All data sets were integrated with MOSFLM (Leslie \u0026 Powell, 2007) and merged with SCALA (Evans, 2006) as implemented in CCP4 (Winn et al., 2011). The ParC55/ParE30–DNA–levofloxacin crystals belonged to space group P21, with unit-cell parameters a = 102.07, b = 161.53, c = 138.60 Å, α = 90.00, β = 94.22, γ = 90.00°. They contained two ParC/ParE–DNA heterodimers in the asymmetric unit. Several data sets were collected, some of which contained visible diffraction to 3.2 Å resolution, but owing to potential internal twinning and space-group ambiguity (most data sets could be integrated in space groups P21 and P212121) and the fact that the structure solution could be obtained in both space groups, careful selection of the integration ranges as well as appropriate data truncation were necessary. The best region of data was integrated to 3.35 Å (see Table 3 ▸ for statistics). The resolution cutoff was based on the rejection criteria R merge \u003c 50% and I/σ(I) \u003e 1.5 in the highest resolution shell. The structure was solved by the molecular-replacement method in Phaser (McCoy et al., 2007) using the levofloxacin–DNA cleavage complex of topoisomerase IV from S. pneumoniae as a search model (PDB entry 3rae; ∼41.8% sequence identity). Refinement was performed in PHENIX (Adams et al., 2002, 2010) using secondary-structure restraints derived by superposition of the K. pneumoniae ParC/ParE model with the previously solved complex of S. pneumoniae ParC/ParE. Rigid-body, positional and TLS refinements were performed. Levofloxacin molecules and magnesium ions were placed during the final stages of refinement based on missing electron density in the σA-weighted 2F obs − F calc and F obs − F calc maps. WinCoot (Emsley \u0026 Cowtan, 2004) was used for interactive model fitting. The structure was verified using WinCoot and PROCHECK (Laskowski et al., 1993). The resulting model had good geometry, with 87.8, 9.9 and 1.3% of residues in the favoured, allowed and generously allowed regions of the Ramachandran plot, respectively, and no more than 1% of residues in disallowed regions. The data-collection and final refinement statistics are given in Tables 3 ▸ and 4 ▸. Sequence alignment was performed in ClustalW (Larkin et al., 2007, McWilliam et al., 2013). Figures were prepared using PyMOL (DeLano, 2008), CHEMDRAW (Evans, 2014) and CorelDRAW (http://www.coreldraw.com). Results and discussion    We have co-crystallized the K. pneumoniae topoisomerase IV ParC/ParE breakage-reunion domain (ParC55; residues 1–490) and ParE TOPRIM domain (ParE30; residues 390–631) with a precut 34 bp DNA duplex (the E-site), stabilized by levofloxacin. The X-ray crystal structure of the complex was determined to 3.35 Å resolution, revealing a closed ParC55 dimer flanked by two ParE30 monomers (Figs. 1 ▸, 2 ▸ and 3 ▸). The overall architecture of this complex is similar to that found for S. pneumoniae topoisomerase–DNA–drug complexes (Laponogov et al., 2009, 2010). Residues 6–30 of the N-terminal α-helix α1 of the ParC subunit again embrace the ParE subunit, ‘hugging’ the ParE subunits close to either side of the ParC dimer (Laponogov et al., 2010). Deletion of this ‘arm’ α1 results in loss of DNA-cleavage activity (Laponogov et al., 2007) and is clearly very important in complex stability (Fig. 3 ▸). This structural feature was absent in our original ParC55 structure (Laponogov et al., 2007; Sohi et al., 2008). The upper region of the topoisomerase complex consists of the E-subunit TOPRIM metal-binding domain formed of four parallel β-sheets and the surrounding α-helices. The C-subunit provides the WHD (winged-helix domain; a CAP-like structure; McKay \u0026 Steitz, 1981) and the ‘tower’ which form the U groove-shaped protein region into which the G-gate DNA binds with an induced U-shaped bend. The lower C-gate region (Fig. 3 ▸) consists of the same disposition of pairs of two long α-helices terminated by a spanning short α-helix forming a 30 Å wide DNA-accommodating cavity through which the T-gate DNA passes as found in the S. pneumoniae complex. Owing to the structural similarity, it appears that the topo­isomerases IV from K. pneumoniae and S. pneumoniae are likely to follow a similar overall topoisomerase catalytic cycle as shown in Fig. 4 ▸; we have confirmation of one intermediate from our recent structure of the full complex (the holoenzyme less the CTD β-pinwheel domain) with the ATPase domain in the open conformation (Laponogov et al., 2013). The G-gate DNA for the S. pneumoniae complex consists of an 18-base-pair E-site sequence (our designation for a DNA site which we first found from DNA-mapping studies; Leo et al., 2005; Arnoldi et al., 2013; Fig. 1 ▸). The crystallized complex was formed by turning over the topoisomerase tetramer in the presence of DNA and levofloxacin and crystallizing the product. In contrast, the K. pneumoniae complex was formed by co-crystallizing the topoisomerase tetramer complex in the presence of a 34-base-pair pre-cleaved DNA in the presence of levofloxacin. In both cases the DNA is bent into a U-form and bound snugly against the protein of the G-gate. We have been able to unambiguously read off the DNA sequences in the electron-density maps. There is 41.6% sequence identity and 54.4% sequence homology between the ParE subunit of K. pneumoniae and that of S. pneumoniae. For the ParC subunits, the figures are 40.8 identity and 55.6% homology between the two organisms. The sequence alignment is given in Supplementary Fig. S1, with the key metal-binding residues and those which give rise to quinolone resistance highlighted. The binding of levofloxacin in the K. pneumoniae complex is shown in Figs. 2 ▸, 3 ▸ and 5 ▸ and is hemi-intercalated into the DNA and stacked against the DNA bases at the cleavage site (positions −1 and +1 of the four-base-pair staggered cut in the 34-mer DNA) which is similar to that found for the S. pneumoniae complex. Fig. 5 ▸ presents side-by-side views of the K. pneumoniae and S. pneumoniae active sites which shows that levofloxacin binds in a very similar manner in these two complexes with extensive π–π stacking interaction between the bases and the drug. The methylpiperazine at C7 (using the conventional quinolone numbering; C9 in the IUPAC numbering) on the drug extends towards residues Glu474 and Glu475 for S. pneumoniae and towards Gln460 and Glu461 for K. pneumoniae, where the glutamate at 474 is substituted by a glutamine at 460 in the Klebsiella strain. Interestingly, for S. pneumoniae we observe only one possible orientation of the C7 groups in both sub­units, while for K. pneumoniae we can see two: one with the same orientation as in S. pneumoniae and other rotated 180° away. They both exist within the same crystal in the two different dimers in the asymmetric unit. The side chains surrounding them in ParE are quite disordered and are more defined in K. pneumoniae (even though this complex is at lower resolution) than in S. pneumoniae. There are no direct hydrogen bonds from the drug to these residues (although it is possible that some are formed through water, which cannot be observed at this resolution). Obviously, the drug–ParE interaction in this region is less strong compared with PD 0305970 binding to the S. pneumoniae DNA complex, where PD 0305970 forms a hydrogen bond to ParE residue Asp475 and can form one to Asp474 if the bond rotates (Laponogov et al., 2010). This may explain why drug-resistance mutations for levofloxacin are more likely to form in the ParC subunits rather than in the ParE subunits. For both complexes there is a Mg2+ ion bound to levofloxacin between the carbonyl group at position 4 of the quinolone and the carboxyl at position 6 (Figs. 2 ▸ and 5 ▸ and Supplementary Fig. 2 ▸). For S. pneumoniae topoisomerase IV, one of the O atoms of the carboxyl of Asp83 points towards the Mg2+ ion and is within hydrogen-bonding distance (5.04 Å) through an Mg2+-coordinated water. For K. pneumoniae both of the carboxyl O atoms are pointing towards the Mg2+ ion at distances of 4.86 and 4.23 Å. These residues are ordered in only one of the two dimers in the K. pneumoniae crystal (the one in which the C7 group is pointing towards the DNA away from ParE, although the conformations of these two groups on the drug are probably not correlated). The topoisomerase IV ParE27-ParC55 fusion protein from K. pneumoniae was fully active in promoting levofloxacin-mediated cleavage of DNA (Fig. 6 ▸). In the absence of the drug and ATP, the protein converted supercoiled pBR322 into a ladder of bands corresponding to relaxed DNA. The inclusion of levofloxacin produced linear DNA in a dose-dependent and ATP-independent fashion. Similar behaviour was observed for the S. pneumoniae topo­isomerase IV ParE30-ParC55 fusion protein. The CC25 (the drug concentration that converted 25% of the supercoiled DNA substrate to a linear form) was 0.5 µM for the Klebsiella enzyme and 1 µM for the pneumococcal enzyme. Interestingly, K. pneumoniae strains are much more susceptible to levofloxacin than S. pneumoniae, with typical MIC values of 0.016 and 1 mg l−1, respectively (Odenholt \u0026 Cars, 2006), reflecting differences in multiple factors (in addition to binding affinity) that influence drug responses, including membrane, peptidoglycan structure, drug-uptake and efflux mechanisms. Moreover, although topoisomerase IV is primarily the target of levofloxacin in S. pneumoniae, it is likely to be gyrase in the Gram-negative K. pneumoniae. In summary, we have determined the first structure of a quinolone–DNA cleavage complex involving a type II topo­isomerase from K. pneumoniae. 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Biol. 424, 109–124. Winn, M. D. et al. (2011). Acta Cryst. D67, 235–242. Wu, C.-C., Li, T.-K., Farh, L., Lin, L.-Y., Lin, T.-S., Yu, Y.-J., Yen, T.-J., Chiang, C.-W. \u0026 Chan, N.-L. (2011). Science, 333, 459–462. Wu, C.-C., Li, Y.-C., Wang, Y.-R., Li, T.-K. \u0026 Chan, N.-L. (2013). Nucleic Acids Res. 41, 10630–10640. Protein and DNA used in the co-crystallization experiment. (a) Coloured diagram of the protein constructs used in crystallization. (b) DNA sequences used in crystallization. The 4 bp overhang is shown in red. Cleavage points are indicated by arrows. Chemical structure of levofloxacin (a) and its conformations observed within the active sites of S. pneumoniae topoisomerase IV (b) and K. pneumoniae topoisomerase IV (c, d). Electron-density maps (2F\nobs − F\ncalc) are shown as meshes for the drug molecules contoured at 1.5σ and are limited to a distance of 2.3 Å from the drug atoms. Overall orthogonal views of the cleavage complex of topoisomerase IV from K. pneumoniae in surface (left) and cartoon (right) representations. The ParC subunit is in blue, ParE is in yellow and DNA is in cyan. The bound quinolone molecules (levofloxacin) are in red and are shown using van der Waals representation. Schematic representation of the catalytic cycle of type II topoisomerases. The ParC N-terminal domain (ParC55) is in grey, the ParC C-terminal β-­pinwheel domain is in silver, the ParE N-terminal ATPase domain is in red, the ParE C-terminal domain (ParE30) is in yellow, the G-gate DNA is in green and the T-segment DNA is in purple. Bound ATP is indicated by pink circles in the ATPase domains (reproduced with permission from Fig. 1 of Lapanogov et al., 2013). Detailed views of the active sites of topoisomerase IV from S. pneumoniae and K. pneumoniae with quinolone molecules bound. The magnesium ions and their coordinating amino acids are shown in purple. The drug molecules and residues known to lead to drug resistance upon mutation are in red. The active-site tyrosine and arginine are in orange. The DNA is shown in silver/cyan. The ParC and ParE backbones are shown in blue and yellow, respectively. Comparison of DNA cleavage by topoisomerase IV core ParE-ParC fusion proteins from K. pneumoniae (KP) and S. pneumoniae (SP) promoted by levofloxacin. Supercoiled plasmid pBR322 (400 ng) was incubated with topoisomerase IV proteins (400 ng) in the absence or presence of levofloxacin at the indicated concentrations. After 60 min incubation, samples were treated with SDS and proteinase K to remove proteins covalent bound to DNA, and the DNA products were examined by gel electrophoresis in 1% agarose. Lane A, supercoiled pBR322 DNA; N, L and S, nicked, linear and supercoiled pBR322, respectively. Macromolecule-production information \nK. pneumoniae topoisomerase IV. Source organism\tK. pneumoniae (strain ATCC 35657)\t \tExpression vector\tpET-29a\t \tExpression host\tE. coli BL21(λDE3) pLysS\t \tComplete amino-acid sequence of the construct produced\t \t Topoisomerase IV (ParE CTD 390–631 and ParC NTD 1–490 fused) \tMKKLTSGPALPGKLADCTAQDLNRTELFLVEGDSAGGSAKQARDREYQAIMPLKGKILNTWEVSSDEVLASQEVHDISVAIGIDPDSDDLSQLRYGKICILADADSDGLHIATLLCALFVRHFRTLVKEGHVYVALPPLYRIDLGKEVYYALTEEEKTGVLEQLKRKKGKPNVQRFKGLGEMNPMQLRETTLDPNTRRLVQLVISDEDEQQTTAIMDMLLAKKRSEDRRNWLQEKGDMADLEVEFMSDMAERLALHEFTENAYLNYSMYVIMDRALPFIGDGLKPVQRRIVYAMSELGLNASAKFKKSARTVGDVLGKYHPHGDSACYEAMVLMAQPFSYRYPLGDGQGNWGAPDDPKSFAAMRYTESRLSKYAELLLSELGQGTVDWVPNFDGTLQEPKMLPARLPNILLNGTTGIAVGMATDIPPHNLREVAKAAITLIEQPKTTLDELLDIVQGPDFPTEAEIITSRAEIRKIYQNGRGSVRMRAVWSKEDGAVVITALPHQVSGAKVLEQIAAQMRNKKLPMVDDLRDESDHENPTRLVIVPRSNRVDMEQVMNHLFATTDLEKSYRINLNMIGLDGRPAVKNLLEILSEWLVFRRDTVRRRLNHRLEKVLKRLHILEGLLVAFLNIDEVIEIIRTEDEPKPALMSRFGISETQAEAILELKLRHLAKLEEMKIRGEQSELEKERDQLQAILASERKMNNLLKKELQADADAFGDDRRSPLHEREEAKAMSHHHHHH\t \t Symmetrized E-site (pre-cut) DNA1\t5′-CGTATTACGTTGTAT-3′\t \t Symmetrized E-site (pre-cut) DNA2\t5′-GATCATACAACGTAATACG-3′\t \t \nS. pneumoniae topoisomerase IV. Source organism\tS. pneumoniae (isolate 7785 St George’s Hospital; Pan \u0026 Fisher, 1996)\t \tExpression vector\tpET-19b (N-terminal His10), pET-29a (C-terminal His6)\t \tExpression host\tE. coli BL21(λDE3) pLysS\t \tComplete amino-acid sequence of the construct produced\t \t ParC55\tMSNIQNMSLEDIMGERFGRYSKYIIQDRALPDIRDGLKPVQRRILYSMNKDSNTFDKSYRKSAKSVGNIMGNFHPHGDSSIYDAMVRMSQNWKNREILVEMHGNNGSMDGDPPAAMRYTEARLSEIAGYLLQDIEKKTVPFAWNFDDTEKEPTVLPAAFPNLLVNGSTGISAGYATDIPPHNLAEVIDAAVYMIDHPTAKIDKLMEFLPGPDFPTGAIIQGRDEIKKAYETGKGRVVVRSKTEIEKLKGGKEQIVITEIPYEINKANLVKKIDDVRVNNKVAGIAEVRDESDRDGLRIAIELKKDANTELVLNYLFKYTDLQINYNFNMVAIDNFTPRQVGIVPILSSYIAHRREVILARSRFDKEKAEKRLHIVEGLIRVISILDEVIALIRASENKADAKENLKVSYDFTEEQAEAIVTLQLYRLTNTDVVVLQEEEAELREKIAMLAAIIGDERTMYNLMKKELREVKKKFATPRLSSLEDTAKALEHHHHHH\t \t ParE30\tMGHHHHHHHHHHSSGHIDDDDKHMKNKKDKGLLSGKLTPAQSKNPAKNELYLVEGDSAGGSAKQGRDRKFQAILPLRGKVINTAKAKMADILKNEEINTMIYTIGAGVGADFSIEDANYDKIIIMTDADTDGAHIQTLLLTFFYRYMRPLVEAGHVYIALPPLYKMSKGKGKKEEVAYAWTDGELEELRKQFGKGATLQRYKGLGEMNADQLWETTMNPETRTLIRVTIEDLARAERRVNVLMGDKVEPRRKWIEDNVKFTLEEATVF \t \t E-site DNA1 \t5′-CATGAATGACTATGCACG-3′\t \t E-site DNA2 \t5′-CGTGCATAGTCATTCATG-3′\t \t Crystallization  \tK. pneumoniae topoisomerase IV\tS. pneumoniae topoisomerase IV\t \tMethod\tVapour diffusion\tCapillary counter-diffusion\t \tPlate type\t24-well Limbro\tN/A\t \tTemperature (K)\t298\t304\t \tProtein concentration (mg ml−1)\t4.5 \t4\t \tBuffer composition of protein solution\t20 mM Tris pH 7.5, 100 mM NaCl, 1 mM β-mercaptoethanol, 0.05% NaN3\t \tComposition of reservoir solution\t0.1 M Tris pH 7.5–8.0, 0–50 mM NaCl, 4–8% PEG 4000, 12–15% glycerol\t50 mM sodium cacodylate pH 6.5, 2.5% Tacsimate, 7% 2-propanol, 62.5 mM KCl, 7.5 mM MgCl2\t \tVolume and ratio of drop\t4 + 2 µl\tN/A\t \tVolume of reservoir (ml)\t0.5\tN/A\t \t Data collection and processing Values in parentheses are for the outer shell.  \tK. pneumoniae topoisomerase IV\tS. pneumoniae topoisomerase IV\t \tDiffraction source\tBeamline I03, Diamond Light Source\t \tWavelength (Å)\t0.97620\t0.97630\t \tTemperature (K)\t100.0\t100.0\t \tDetector\tPilatus 6M-F\tADSC Quantum 315\t \tCrystal-to-detector distance (mm)\t502.22\t377.629\t \tRotation range per image (°)\t0.2\t0.25\t \tTotal rotation range (°)\t180\t75\t \tExposure time per image (s)\t0.2\t1.0\t \tSpace group\tP21\tP3121\t \ta, b, c (Å)\t102.07, 161.53, 138.60\t157.83, 157.83, 211.15\t \tα, β, γ (°)\t90, 94.22, 90\t \t \tMosaicity (°)\t0.237\t0.466\t \tResolution range (Å)\t86.12–3.35 (3.53–3.35)\t50–2.90 (3.00–2.90)\t \tTotal No. of reflections\t160764\t311576\t \tNo. of unique reflections\t63406\t67471\t \tCompleteness (%)\t98.5 (98.4)\t99.4 (99.9)\t \tMultiplicity\t2.53 (2.59)\t4.6 (4.7)\t \t〈I/σ(I)〉\t3.48 (1.95)\t16.14 (3.48)\t \tRr.i.m.†\t0.116 (0.434)\t0.08 (0.515)\t \tOverall B factor from Wilson plot (Å2)\t53.29\t73.37\t \t Estimated R\nr.i.m. = R\nmerge[N/(N − 1)]1/2, where N is the data multiplicity. Structure solution and refinement Values in parentheses are for the outer shell.  \tK. pneumoniae topoisomerase IV\tS. pneumoniae topoisomerase IV\t \tResolution range (Å)\t85.01–3.35 (3.40–3.35)\t41.83–2.90 (2.93–2.90)\t \tCompleteness (%)\t98.3\t99.5\t \tσ Cutoff\tF \u003e 1.350σ(F)\tF \u003e 1.34σ(F)\t \tNo. of reflections, working set\t60158 (2615)\t67471 (1992)\t \tNo. of reflections, test set\t3208 (142)\t6838 (218)\t \tFinal Rcryst\t0.224 (0.2990)\t0.186 (0.2806)\t \tFinal Rfree\t0.259 (0.3537)\t0.226 (0.3562)\t \tNo. of non-H atoms\t \t Protein\t18741\t10338\t \t Nucleic acid\t1608\t730\t \t Ligand\t104\t52\t \t Ion\t8\t6\t \t Water\t—\t54\t \t Total\t20461\t11180\t \tR.m.s. deviations\t \t Bonds (Å)\t0.002\t0.008\t \t Angles (°)\t0.611\t1.221\t \tAverage B factors (Å2)\t \t Protein\t58.05\t76.7\t \t Nucleic acid\t64.85\t90.7\t \t Ligand\t60.14\t95.7\t \t Ion\t42.62\t84.5\t \t Water\t—\t64.2\t \tRamachandran plot\t \t Most favoured (%)\t93\t94\t \t Allowed (%)\t6\t6\t 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annotated_BioC_JSON/PMC4855620_ann.json

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+[{"sourceid":"4855620","sourcedb":"","project":"","target":"","text":"Structure‐activity relationship of the peptide binding‐motif mediating the BRCA2:RAD51 protein–protein interaction  RAD51 is a recombinase involved in the homologous recombination of double‐strand breaks in DNA. RAD51 forms oligomers by binding to another molecule of RAD51 via an ‘FxxA’ motif, and the same recognition sequence is similarly utilised to bind BRCA2. We have tabulated the effects of mutation of this sequence, across a variety of experimental methods and from relevant mutations observed in the clinic. We use mutants of a tetrapeptide sequence to probe the binding interaction, using both isothermal titration calorimetry and X‐ray crystallography. Where possible, comparison between our tetrapeptide mutational study and the previously reported mutations is made, discrepancies are discussed and the importance of secondary structure in interpreting alanine scanning and mutational data of this nature is considered. Abbreviations  BRCA2, breast cancer type‐2 susceptibility protein  HR, homologous recombination  ITC, isothermal titration calorimetry  PPI, protein–protein interaction  SAR, structure activity relationship Eukaryotic RAD51, archeal RadA and prokaryotic RecA are a family of ATP‐dependent recombinases involved in homologous recombination (HR) of double‐strand breaks in DNA 1. RAD51 interacts with BRCA2, and is thought to localise RAD51 to sites of DNA damage 2, 3. Both BRCA2 and RAD51 together are vital for helping to repair and maintain a high fidelity in DNA replication. BRCA2 especially has garnered much attention in a clinical context, as many mutations have been identified that drive an increased risk of cancer in individuals 4, 5. Although the inactivation of the BRCA2:RAD51 DNA repair pathway can cause genomic instability and eventual tumour development, an inability to repair breaks in DNA may also engender a sensitivity to ionising radiation 6, 7. For this reason it is hypothesised that in tumour cells with an intact BRCA2:RAD51 repair pathway, small molecules which prevent the interaction between RAD51 and BRCA2 may confer radiosensitivity by disabling the HR pathway 8. The interaction between the two proteins is mediated by eight BRC repeats, which are characterised by a conserved ‘FxxA’ motif 9. RAD51 and RadA proteins also contain an ‘FxxA’ sequence (FTTA for human RAD51) through which it can bind to other RAD51 and RadA molecules, and oligomerise to form higher order filament structures on DNA. The common FxxA motifs of both the BRC repeats and RAD51 oligomerisation sequence are recognised by the same FxxA‐binding site of RAD51. In general, the dominant contribution of certain residues to the overall binding energy of a protein–protein interaction are known as ‘hot‐spot’ residues. Interestingly, small molecule inhibitors of PPIs are often found to occupy the same pockets which are otherwise occupied by hot‐spot residues in the native complex. It is therefore of great interest to identify hot‐spots in an effort to guide drug discovery efforts against a PPI. Further, a correlation between residues that are strongly conserved and hot‐spot residues has been reported 10. Purely based on the amino acid consensus sequence reported by Pellegrini et al., 11 phenylalanine and alanine would both be expected to be hot‐spots and to a lesser extent, threonine. However, whilst the identification of highly conserved residues may be a good starting point for identifying hot‐spots, experimental validation by mutation of these sequences is vital. The importance of residues in the FxxA motif has been probed by a variety of techniques, collated in Table 1. Briefly, mutating phenylalanine to glutamic acid inactivated the BRC4 peptide and prevented RAD51 oligomerisation 11, 12. A phenylalanine‐truncated BRC4 is also found to be inactive 13, however, introducing a tryptophan for phenylalanine was found to have no significant effect on BRC4 affinity 12. A glutamine replacing the histidine in BRC4 maintains BRC4 activity 13. The ability of BRC3 to interact with RAD51 nucleoprotein filaments is disrupted when threonine is mutated to an alanine 3. Similarly, mutating alanine to glutamic acid in the RAD51 oligomerisation sequence 11 or to serine in BRC4 13 leads to loss of interaction in both cases. The BRC5 repeat in humans has serine in the place of alanine, and is thought to be a nonbinding repeat 12. Mutations identified in the clinic, in the FxxA region of the BRC repeats of BRCA2 are collated in Table 1 14. It is difficult to state the clinical relevance of these mutations as they are annotated as ‘unvalidated’, that is, it is not known whether they contribute to the disease phenotype or are neutral variants. For completeness, we present them here with this caveat, and to make the comment that these clinical mutations are consistent with abrogating the interaction with RAD51. Summary of FxxA‐relevant mutations previously reported and degree of characterisation. The mutation, relevant peptide context, resulting FxxA motif sequence and experimental technique for each entry is given. For clarity, mutated residues are shown in bold Mutation contexta\tMutation\tFxxA motif\tTechnique used\tEffect\t \tRAD51 (FTTA)\tF86E\tETTA\tImmunoprecipitation 11\tNo binding\t \tBRC4 (FHTA)\tF1524E\tEHTA\tCompetitive ELISA 12\tPeptide inactive\t \tBRC4 (FHTA)\tF1524W\tWHTA\tCompetitive ELISA 12\tComparable activity to WT\t \tBRC4 (FHTA)\tF1524V\tVHTA\tBRCA2 mutations database 14\t–\t \tBRC4 (FHTA)\tΔF1524\t‐HTA\tDissociation of RAD51‐DNA complex 13\tPeptide inactive\t \tBRC4 (FHTA)\tH1525Q\tFQTA\tDissociation of RAD51‐DNA complex 13\tComparable activity\t \tBRC7 (FSTA)\tS1979R\tFRTA\tBRCA2 mutations database 14\t–\t \tBRC3 (FQTA)\tT1430A\tFQAA\tRAD51:DNA bandshift assay 3\tPeptide inactive\t \tBRC3 (FQTA)\tT1430A\tFQAA\tElectron microscopic visualisation of nucleoprotein filaments 3\tPeptide inactive\t \tBRC1 (FRTA)\tT1011R\tFRRA\tBRCA2 mutations database 14\t–\t \tBRC2 (FYSA)\tS1221P\tFYPA\tBRCA2 mutations database 14\t–\t \tBRC2 (FYSA)\tS1221Y\tFYYA\tBRCA2 mutations database 14\t–\t \tRAD51 (FTTA)\tA89E\tFTTE\tImmunoprecipitation 11\tNo binding\t \tBRC4 (FHTA)\tA1527S\tFHTS\tDissociation of RAD51‐DNA complex 13\tPeptide inactive\t \t The wild‐type FxxA sequence is indicated in parenthesis. In this work, we report the most detailed study of systematic mutations of peptides to probe the FxxA‐binding motif to date. We have chosen to focus on tetrapeptides, which allows us to examine the effect of mutation on the fundamental unit of binding, FxxA, rather than in the context of either the BRC repeat or self‐oligomerisation sequence. Affinities of peptides were measured directly using Isothermal Titration Calorimetry (ITC) and the structures of many of the peptides bound to humanised RadA were determined by X‐ray crystallography. The use of ITC is generally perceived as a gold‐standard in protein–ligand characterisation, rather than a competitive assay which may be prone to aggregation artefacts. Wild‐type human RAD51, however, is a heterogeneous mixture of oligomers and when monomerised by mutation, is highly unstable. In this context, we have previously reported the use of stable monomeric forms of RAD51, humanised from Pyrococcus furiosus homologue RadA, for ITC experiments and X‐ray crystallography 8, 15. Materials and methods Peptide synthesis Peptides were synthesised using solid‐phase FMOC chemistry by Alta Biosciences (Birmingham, UK) or the Protein and Nucleic Acid Service at the Department of Biochemistry (University of Cambridge). All peptides prepared and used in the study were N‐acetylated and C‐amide terminated. Protein preparation Protein expression and purification was performed as described previously 15. In brief, monomeric HumRadA2 was expressed in E. coli using T7‐based expression vector at 37 °C for 3 h. Soluble cell lysate was heat treated to precipitate most of the cellular proteins and the soluble fraction containing HumRadA2 was purified using a combination of cation exchange chromatography at pH 6.0 and size‐exclusion chromatography in 10 mm MES, 100 mm NaCl pH 6.0 buffer. Protein concentration was determined using the calculated extinction coefficient at 280 nm, and stored at −80 °C in small aliquots after flash freezing. Isothermal titration calorimetry Isothermal titration calorimetry experiments were performed at 25 °C on a MicroCal iTC200. HumRadA2 (600 μm in 20 mm MES pH 6.0 with 100 mm NaCl and 0.5 mm EDTA) was diluted with Tris buffer (200 mm, pH 7.5 with 100 mm NaCl) to 64–83 μm. Peptides were dissolved in MilliQ water (50 mm) and an aliquot taken and diluted with 200 mm Tris, pH 7.5, 100 mm NaCl to give a ligand solution of 2.5–5 mm. The peptide solution was titrated into the protein solution; 16 injections (2.4 μL) of 4.8 s duration were made at 80‐s intervals. The initial injection of ligand (0.4 μL) was discarded during data analysis. Control experiments of peptides to buffer showed insignificant heats. The data were processed and thermodynamic parameters obtained by fitting the data to a single‐site‐binding model using Origin software and fixing the stoichiometry as 1.0 for weak‐binding ligands 16. All data from ITC measurements are shown in the Figs S1 and S2. X‐ray crystallography Monomerised RadA proteins were crystallised in the same conditions as described previously 15. Peptides were soaked into the crystals at 2–5 mm concentration overnight in the presence of 10% glycerol as a cryoprotectant. Crystals were cryo‐cooled in liquid nitrogen and data collected at synchrotron light sources and processed using XDS: details of this are found in crystallographic table (Table S1 in Supporting Information). Structures were solved by molecular replacement using unliganded, monomeric RadA coordinates (PDB: 4b3b, after removal of FHTA peptide) as a search model and refined with an automated procedure using Refmac5 17. After inspection of the resulting electron density, the bound peptides were modelled into the density and structures were further refined using Refmac5 18 and phenix.refine 19, and manually rebuilt using Coot 20. Coordinates and structure factors have been deposited in the PDB under accession codes as listed in Table 2 and in the crystallographic data table in the Supporting Information. With the exception of FATA peptide complex, which was crystallised with wild‐type RadA, the structures are determined using HumRadA1 mutant. Summary of peptide‐binding data determined by ITC against HumRadA2. Mutated residues are highlighted in bold. All peptides were N‐acetylated and C‐amide terminated Table entry\tPeptide\tKD/μm\tΔH/cal·mol−1\tTΔS/cal·mol−1\tPDB code\t \tFirst position variation\t \t1\tFHTA\t280 ± 20\t−2388 ± 94\t2453\t4b3b15\t \t2\tWHTA\t93 ± 3\t−2768 ± 34\t2727\t5fow\t \tSecond position variation\t \t3\tFATA\t280 ± 29\t−1820 ± 109\t3010\t5fpka\t \t4\tFNTA\t613 ± 44\t−4036 ± 177\t346\t–\t \t5\tFPTA\tNo detectable binding\t\t\t–\t \tThird position variation\t \t6\tFHPA\t113 ± 3\t−2155 ± 26\t3218\t5fou\t \t7\tFHAA\t675 ± 60\t−7948 ± 466\t−3636\t5fox\t \tFourth position variation\t \t8\tFHTG\t1590 ± 300\t−5518 ± 924\t−1702\t5fov\t \t9\tFHTU\t680 ± 51\t−14 600 ± 771\t−10 281\t5fot\t \tCombination\t \t10\tWHPA\t330 ± 25\t−6801 ± 318\t−2044\t–\t \tPeptide truncations\t \t11\tFH\t\t \tNo binding detected\t \tNo binding detected\t \tNo binding detected\t \t\t\t\t–\t \t12\tFHT\t–\t \t13\tHTA\t–\t \t Structure solved with wild‐type RadA. Sequence analysis Sequences of mammalian RAD51 proteins and archeal RadA orthologues were obtained from Ensembl (www.ensembl.org) and Uniprot (www.uniprot.org) databases. Sequences were aligned using ClustalX2, and aligned sequences for the FxxA motifs were used in WebLogo (weblogo.berkely.edu/logo.cgi) server 21 to derive the consensus diagrams shown in Figs 1 and 4. All the sequences used in these analyses are shown in Figs S4, S5 and S6. Conservation of FxxA motif (A) BRC4 peptide (green cartoon) bound to truncated human RAD51 (grey surface) (PDB: 1n0w, 11). The blue dashed box highlights the FxxA interaction pocket. (B) Two interacting protein molecules of RAD51 from Saccharomyces cerevisiae are shown. One RAD51 (green cartoon) interacts with another molecule of RAD51 (grey and pink surface) via the FxxA pocket indicated by the dashed blue box. The N‐terminal domain of one RAD51 protomer is highlighted in pink for clarity and the green arrow indicates the location of this protomer's FxxA oligomerisation sequence (PDB: 1szp, 29). (C) Conservation of FxxA motif across the human BRC repeats and (D) across 21 eukaryotic RAD51s and 24 RadAs, with the size of the letters proportional to the degree of conservation. Sequence figures generated using Weblogo 3.0 21, sequence details are found in the Supporting Information. Results We have mutated and truncated the tetrapeptide epitope FHTA, and examined the effects both structurally and on the binding affinity with humanised RadA. As a comparative reference, we are using the FHTA sequence derived from the most tightly binding BRC repeat, BRC4 22. The peptides used are N‐acetylated and C‐amide terminated in order to provide the most relevant peptide in the context of a longer peptide chain. A summary of the peptide sequence, PDB codes and K D data measured by ITC with the corresponding ΔH and TΔS values are collated in Table 2. Phe1524 of BRC4 binds in a small surface pocket of human RAD51, defined by the hydrophobic side chains of residues Met158, Ile160, Ala192, Leu203 and Met210. The residue is highly conserved across BRC repeats and oligomerisation sequences. Consistent with this, the truncated HTA tripeptide could not be detected to bind to humanised, monomeric RadA, HumRadA2 (Table 2, entry 13). As previously discussed, there is some evidence that substituting a tryptophan for the phenylalanine at this position was tolerated in the context of BRC4 12. Therefore, the WHTA peptide was tested and found to not only be tolerated, but to increase the binding affinity of the peptide approximately threefold. The second position of the tetrapeptide was found to be largely invariant to changes in the side chains that were investigated. The residue makes no interactions with the RAD51 protein, but may make an internal hydrogen bond with Thr1520 in the context of BRC4, Fig. 3A. Replacing the histidine with an asparagine, chosen to potentially mimic the hydrogen bond donor–acceptor nature of histidine, resulted in a moderate, twofold decrease in potency (Table 2, entry 4). Mutating to an alanine, recapitulated the potency of FHTA, implying that the interactions made by histidine do not contribute overall to binding affinity (Table 2, entry 3). FPTA was also tested, but was found to have no affinity for the protein (Table 2, entry 5). Modelling suggests that a proline in the second position would be expected to clash sterically with the surface of the protein, and provides a rationale for the lack of binding observed. Threonine was mutated to an alanine, resulting in only a moderately weaker K D (twofold, Table 2, entry 7). In the context of a tetrapeptide at least, this result implies a lack of importance of a threonine at this position. Interestingly, it was found that a proline at this position improved the affinity almost threefold, to 113 μm (Table 2, entry 6). This beneficial mutation was incorporated with another previously identified variant to produce the peptide WHPA. Disappointingly, the combined effect of the mutations was not additive and the potency was weakened to 690 μm. While the importance of the phenylalanine may be possible to predict from examination of the crystal structure, the alanine appears to be of much less importance in this regard. It is, however, a highly conserved residue and clearly of interest for systematic mutation. Removing the alanine residue entirely produced the truncated tripeptide FHT, which did not bind (Table 2, entry 12). The unnatural amino acid, α‐amino butyric acid (U), was introduced at the fourth position, positioning an ethyl group into the alanine pocket (Table 2, entry 9). Perhaps surprisingly, it was accommodated and the affinity dropped only by twofold as compared to FHTA. The effect of simply removing the β‐carbon of alanine, by mutation to glycine (FHTG), produced an approximately sixfold drop in binding affinity (Table 2, entry 8). This is in line with the observation that alanine is not 100% conserved and some archeal RadA proteins contain a glycine in the place of alanine 23. Structural characterisation of peptide complexes Structures of the key tetrapeptides were solved by soaking into crystals of a humanised form of RAD51, HumRadA1, which we have previously reported as a convenient surrogate system for RAD51 crystallography 15. The corresponding PDB codes are indicated in Table 2 and crystallographic data are found in the Supporting Information. All structures are of high resolution (1.2–1.7 Å) and the electron density for the peptide was clearly visible after the first refinement using unliganded RadA coordinates (Fig. S1). Some of the SAR observed in the binding analysis can be interpreted in terms of these X‐ray crystal structures. For example, an overlay of the bound poses of the ligands FHTA and FHPA (Fig. 2B) reveals a high similarity in the binding modes, indicating that the conformational rigidity conferred by the proline is compatible with the FHTA‐binding mode, and a reduction in an entropic penalty of binding may be the source of the improvement in affinity. WHTA peptide shows a relative dislocation when compared to FHTA (Fig 2A), with the entire ligand backbone of WHTA shifted to accommodate the change in the position of the main chain carbon of the first residue, as the larger indole side chain fills the Phe pocket. This shift is translated all the way to the alanine side chain. It is possible that this mutation is beneficial in the tetrapeptide context and neutral in the full‐length BRC4 context because the smaller peptide is less constrained and allowed to explore more conformations. An attempt to combine both the tryptophan and proline mutations, however, led to no improvement for WHPA peptide compared to FHTA. One possible explanation is that the ‘shifted’ binding mode observed in WHTA was not compatible with the conformational restriction that the proline of WHPA introduced. Comparison of different peptide complexes (A) Overlay with FHTA (grey) and WHTA (purple) showing a small relative displacement of the peptide backbone. (B) Superposition of FHTA (grey) and FHPA (yellow), showing conservation of backbone orientation (C) Overlay of FHTU (green), FHTA (grey) and FHTG (cyan). The thermodynamic data of peptide binding are also shown in Table 2. Although we have both thermodynamic data and high‐quality X‐ray structural information for some of the mutant peptides, we do not attempt to interpret differences in thermodynamic profiles between ligands, that is, to analyse ΔΔH and ΔΔS. Although ΔH and ΔS are tabulated, the K Ds measured are relatively weak and necessarily performed under low c‐value conditions. In this experimental regime, nonsigmoidal curves are generated and therefore errors in ΔH are expected to be much higher than the errors from model fitting given in Table 2 16. As ΔS is derived from ΔG by subtracting ΔH, errors in ΔH will be correlated with errors in ΔS, giving rise to a ‘phantom’ enthalpy–entropy compensation. Such effects have been discussed by Klebe 24 and Chodera and Mobley 25 and will frustrate attempts to interpret the measured ΔΔH and ΔΔS. Understanding mutations, residue conservation and epitope secondary structure The conserved phenylalanine and alanine residues of the FHTA sequence were both found to be essential for binding by ITC. Conversely the second position histidine residue, corresponding to the unconserved His1525 in the BRC4 sequence, could be mutated without significant effect on the peptide affinity. The more general correlation between hot‐spot residues in protein–protein interactions and the high conservation of such residues has been previously reported 10, 26. Interestingly, however, the highly conserved threonine residue could be mutated without affecting the peptide affinity. This unexpected result, in the light of its very high conservation in the BRC and oligomerisation sequences, begs the question of what the role of Thr1526 is and highlights a potential pitfall and need for caution in the experimental design of alanine mutation studies. As the FHTA peptide is potentially a surrogate peptide for both the BRC repeat peptides and the RAD51 self‐oligomerisation peptide, it is useful to examine the role of Thr1526 (BRC4) and the analogous Thr87 (RAD51) in both binding contexts in more detail. Structural information for these two interactions is limited. Only one structure of BRC4 is published in complex with human RAD51 (PDB: 1n0w). Figure 3A shows the binding pose of BRC4 when bound to RAD51 and the intrapeptide hydrogen bonds that are made by BRC4. While Phe1524 and Ala1527 are buried in hydrophobic pockets on the surface, His1525 is close enough to form a hydrogen bond with the carbonyl of Thr1520, but the rotamer of His1525, supported by clearly positioned water molecules, is not compatible with hydrogen bonding. Also, Thr1520 is constrained by crystal contacts in this structure. Lack of conservation of this residue supports the idea that this interaction is not crucial for RAD51:BRC repeat binding. (A) Highlight of intra‐BRC4 interactions when bound to RAD51 (omitted for clarity) (PDB: 1n0w), with key residues shown in colour. (B) Intrapeptide interactions from oligomerisation epitope of S. cerevisiae \nRAD51 when bound to next RAD51 in the filament (PDB: 1szp). Colouring as in (A). Residue numbering relates to the S. cerevisiae \nRAD51 protein, the corresponding human residues are in parentheses. Either a threonine or serine is most commonly found in the third position of the FxxA motif. Thr1526 makes no direct interactions with the RAD51 protein, but instead forms a hydrogen bond network with the highly conserved S1528 and K1530 (Fig. 1C). The high degree of conservation of these three residues suggests an important possible role in facilitating a turn and stabilising the conformation of the peptide as it continues its way to a second interaction site on the side of RAD51. With respect to understanding the RAD51:RAD51 interaction, no human crystal structure has been published, however, several oligomeric structures of archaeal RadA as well that of Saccharomyces cerevisiae RAD51 have been reported 27, 28, 29. Figure 3B shows a highlight of the FxxA portion of oligomerisation peptide from the S. cerevisiae RAD51 structure, with residues in parentheses corresponding to the human RAD51 protein. The conserved threonine residue at the third position forms a hydrogen bond with the peptide backbone amide, which forms the base of an α‐helix. In both structural contexts, the role of the third position threonine in FxxA seems to be in stabilising secondary structure; a β‐turn in the case of BRC binding and an α‐helix in the case of RAD51 oligomerisation. In the tetrapeptide context these secondary interactions are not present and mutation of threonine to alanine would be expected to have little effect on affinity. In line with this, although we observe a slight twofold weakening of peptide affinity, the effect is far from being as drastic or inactivating as reported in longer peptide backgrounds 3. It would be interesting to investigate the importance of this residue in the context of the BRC4 peptide, and the oligomerisation peptide. Rather than indifference to alanine mutation, a significant effect, via lack of secondary structure stabilisation, would be predicted, as indeed has been reported for BRC3 3. Conclusions The key observations from this work are shown in Fig 4. Two residues in the FxxA motif, phenylalanine and alanine, are highly conserved (Fig 4a). Phenylalanine mutated to tryptophan, in the context of the tetrapeptide improved potency, contrary to the reported result of comparable activity in the context of BRC4 12. Proline at the third position similarly improved potency. Activity was lost by mutating the terminal alanine to glycine, but recovered somewhat with the novel α‐amino butyric acid (U). Threonine was found to be relatively unimportant in the tetrapeptides but has been previously reported to be crucial in the context of BRC3. The reason for this disconnection is suggested to be that threonine plays a role in stabilising the β‐turn in the BRC repeats, which is absent in the tetrapeptides studied. This may lead to a more general caution, that hot‐spot data should be interpreted by considering the bound interaction with the protein, as well as the potential role in stabilising the bound peptide secondary structure. In either case, the requirement for structural data in correctly interpreting alanine‐scanning experiments is reinforced. Summary of key observations (A) FxxA motif sequence conservation of Rad51 oligomerisation sequences and BRC repeats. (B) Highlight of SAR identified for the tetrapeptide. The differences in ΔG for different peptide variants relative to FHTA are shown in the bar chart with colouring matching with the structural overlay below. (C) Overlay of tetrapeptide structures, with wild‐type FHTA peptide across the figure for reference and truncated segments of mutated residues shown in each panel. Purple carbon is WHTA, light blue is FATA, yellow is FHPA, cyan is FHTG and grey carbon is FHTA. Note the C‐terminal amide changes position in FHTG without the anchoring methyl group. Supporting information References RAD51 protein involved in repair and recombination in S. cerevisiae is a RecA‐like protein Cancer susceptibility and the functions of BRCA1 and BRCA2 Role of BRCA2 in control of the RAD51 recombination and DNA repair protein The emerging landscape of breast cancer susceptibility Variation of breast cancer risk among BRCA1/2 carriers Clinical relevance of normal and tumour cell radiosensitivity in BRCA1/BRCA2 mutation carriers: a review Embryonic lethality and radiation hypersensitivity mediated by Rad51 in mice lacking Brca2 Small‐molecule inhibitors that target protein‐protein interactions in the RAD51 family of recombinases Sequence fingerprints in BRCA2 and RAD51: implications for DNA repair and cancer Hot regions in protein–protein interactions: the organization and contribution of structurally conserved hot spot residues Insights into DNA recombination from the structure of a RAD51‐BRCA2 complex Two modules in the BRC repeats of BRCA2 mediate structural and functional interactions with the RAD51 recombinase Design of potent inhibitors of human RAD51 recombinase based on BRC motifs of BRCA2 protein: modeling and experimental validation of a chimera peptide Description and analysis of genetic variants in French hereditary breast and ovarian cancer families recorded in the UMD‐BRCA1/BRCA2 databases Using a fragment‐based approach to target protein‐protein interactions On the value of c: can low affinity systems be studied by isothermal titration calorimetry? REFMAC5 for the refinement of macromolecular crystal structures Refinement of macromolecular structures by the maximum‐likelihood method PHENIX: a comprehensive Python‐based system for macromolecular structure solution Coot: model‐building tools for molecular graphics WebLogo: a sequence logo generator Two classes of BRC repeats in BRCA2 promote RAD51 nucleoprotein filament function by distinct mechanisms Crystal structure of archaeal recombinase RADA: a snapshot of its extended conformation Applying thermodynamic profiling in lead finding and optimization Entropy‐enthalpy compensation: role and ramifications in biomolecular ligand recognition and design Evolutionary predictions of binding surfaces and interactions Crystal structure of a Rad51 filament Crystal structure of Methanococcus voltae RadA in complex with ADP: hydrolysis‐induced conformational change Full‐length archaeal RAD51 structure and mutants: mechanisms for RAD51 assembly and control by 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annotated_BioC_JSON/PMC4871749_ann.json

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+[{"sourceid":"4871749","sourcedb":"","project":"","target":"","text":"The Taf14 YEATS domain is a reader of histone crotonylation The discovery of new histone modifications is unfolding at startling rates, however, the identification of effectors capable of interpreting these modifications has lagged behind. Here we report the YEATS domain as an effective reader of histone lysine crotonylation – an epigenetic signature associated with active transcription. We show that the Taf14 YEATS domain engages crotonyllysine via a unique π-π-π-stacking mechanism and that other YEATS domains have crotonyllysine binding activity. Crotonylation of lysine residues (crotonyllysine, Kcr) has emerged as one of the fundamental histone post-translational modifications (PTMs) found in mammalian chromatin. This epigenetic PTM is widespread and enriched at active gene promoters and potentially enhancers. The crotonyllysine mark on histone H3K18 is produced by p300, a histone acetyltransferase also responsible for acetylation of histones. Owing to some differences in their genomic distribution, the crotonyllysine and acetyllysine (Kac) modifications have been linked to distinct functional outcomes. p300-catalyzed histone crotonylation, which is likely metabolically regulated, stimulates transcription to a greater degree than p300-catalyzed acetylation. The discovery of individual biological roles for the crotonyllysine and acetyllysine marks suggests that these PTMs can be read by distinct readers. While a number of acetyllysine readers have been identified and characterized, a specific reader of the crotonyllysine mark remains unknown (reviewed in). A recent survey of bromodomains (BDs) demonstrates that only one BD associates very weakly with a crotonylated peptide, however it binds more tightly to acetylated peptides, inferring that bromodomains do not possess physiologically relevant crotonyllysine binding activity. The family of acetyllysine readers has been expanded with the discovery that the YEATS (Yaf9, ENL, AF9, Taf14, Sas5) domains of human AF9 and yeast Taf14 are capable of recognizing the histone mark H3K9ac. The acetyllysine binding function of the AF9 YEATS domain is essential for the recruitment of the histone methyltransferase DOT1L to H3K9ac-containing chromatin and for DOT1L-mediated H3K79 methylation and transcription. Similarly, activation of a subset of genes and DNA damage repair in yeast require the acetyllysine binding activity of the Taf14 YEATS domain. Consistent with its role in gene regulation, Taf14 was identified as a core component of the transcription factor complexes TFIID and TFIIF. However, Taf14 is also found in a number of chromatin-remodeling complexes (i.e., INO80, SWI/SNF and RSC) and the histone acetyltransferase complex NuA3, indicating a multifaceted role of Taf14 in transcriptional regulation and chromatin biology. In this study, we identified the Taf14 YEATS domain as a reader of crotonyllysine that binds to histone H3 crotonylated at lysine 9 (H3K9cr) via a distinctive binding mechanism. We found that H3K9cr is present in yeast and is dynamically regulated. To elucidate the molecular basis for recognition of the H3K9cr mark, we obtained a crystal structure of the Taf14 YEATS domain in complex with H3K9cr5-13 (residues 5–13 of H3) peptide (Fig. 1, Supplementary Results, Supplementary Fig. 1 and Supplementary Table 1). The Taf14 YEATS domain adopts an immunoglobin-like β sandwich fold containing eight anti-parallel β strands linked by short loops that form a binding site for H3K9cr (Fig. 1b). The H3K9cr peptide lays in an extended conformation in an orientation orthogonal to the β strands and is stabilized through an extensive network of direct and water-mediated hydrogen bonds and a salt bridge (Fig. 1c). The most striking feature of the crotonyllysine recognition mechanism is the unique coordination of crotonylated lysine residue. The fully extended side chain of K9cr transverses the narrow tunnel, crossing the β sandwich at right angle in a corkscrew-like manner (Fig. 1b and Supplementary Figure 1b). The planar crotonyl group is inserted between Trp81 and Phe62 of the protein, the aromatic rings of which are positioned strictly parallel to each other and at equal distance from the crotonyl group, yielding a novel aromatic-amide/aliphatic-aromatic π-π-π-stacking system that, to our knowledge, has not been reported previously for any protein-protein interaction (Fig. 1d and Supplementary Fig. 1c). The side chain of Trp81 appears to adopt two conformations, one of which provides maximum π-stacking with the alkene functional group while the other rotamer affords maximum π-stacking with the amide π electrons (Supplementary Fig. 1c). The dual conformation of Trp81 is likely due to the conjugated nature of the C=C and C=O π-orbitals within the crotonyl functional group. In addition to π-π-π stacking, the crotonyl group is stabilized by a set of hydrogen bonds and electrostatic interactions. The π bond conjugation of the crotonyl group gives rise to a dipole moment of the alkene moiety, resulting in a partial positive charge on the β-carbon (Cβ) and a partial negative charge on the α-carbon (Cα). This provides the capability for the alkene moiety to form electrostatic contacts, as Cα and Cβ lay within electrostatic interaction distances of the carbonyl oxygen of Gln79 and of the hydroxyl group of Thr61, respectively. The hydroxyl group of Thr61 also participates in a hydrogen bond with the amide nitrogen of the K9cr side chain (Fig. 1d). The fixed position of the Thr61 hydroxyl group, which facilitates interactions with both the amide and Cα of K9cr, is achieved through a hydrogen bond with imidazole ring of His59. Extra stabilization of K9cr is attained by a hydrogen bond formed between its carbonyl oxygen and the backbone nitrogen of Trp81, as well as a water-mediated hydrogen bond with the backbone carbonyl group of Gly82 (Fig 1d). This distinctive mechanism was corroborated through mapping the Taf14 YEATS-H3K9cr binding interface in solution using NMR chemical shift perturbation analysis (Supplementary Fig. 2a, b). Binding of the Taf14 YEATS domain to H3K9cr is robust. The dissociation constant (Kd) for the Taf14 YEATS-H3K9cr5-13 complex was found to be 9.5 μM, as measured by fluorescence spectroscopy (Supplementary Fig. 2c). This value is in the range of binding affinities exhibited by the majority of histone readers, thus attesting to the physiological relevance of the H3K9cr recognition by Taf14. To determine whether H3K9cr is present in yeast, we generated whole cell extracts from logarithmically growing yeast cells and subjected them to Western blot analysis using antibodies directed towards H3K9cr, H3K9ac and H3 (Fig. 2a, b, Supplementary Fig. 3 and Supplementary Table 2). Both H3K9cr and H3K9ac were detected in yeast histones; to our knowledge, this is the first report of H3K9cr occurring in yeast. We next asked if H3K9cr is regulated by the actions of histone acetyltransferases (HATs) and histone deacetylases (HDACs). Towards this end, we probed extracts derived from yeast cells in which major yeast HATs (HAT1, Gcn5, and Rtt109) or HDACs (Rpd3, Hos1, and Hos2) were deleted. As shown in Figure 2a, b and Supplementary Fig. 3e, H3K9cr levels were abolished or reduced considerably in the HAT deletion strains, whereas they were dramatically increased in the HDAC deletion strains. Furthermore, fluctuations in the H3K9cr levels were more substantial than fluctuations in the corresponding H3K9ac levels. Together, these results reveal that H3K9cr is a dynamic mark of chromatin in yeast and suggest an important role for this modification in transcription as it is regulated by HATs and HDACs. We have previously shown that among acetylated histone marks, the Taf14 YEATS domain prefers acetylated H3K9 (also see Supplementary Fig. 3b), however it binds to H3K9cr tighter. The selectivity of Taf14 towards crotonyllysine was substantiated by 1H,15N HSQC experiments, in which either H3K9cr5-13 or H3K9ac5-13 peptide was titrated into the 15N-labeled Taf14 YEATS domain (Fig. 2c and Supplementary Fig. 4a, b). Binding of H3K9cr induced resonance changes in slow exchange regime on the NMR time scale, indicative of strong interaction. In contrast, binding of H3K9ac resulted in an intermediate exchange, which is characteristic of a weaker association. Furthermore, crosspeaks of Gly80 and Trp81 of the YEATS domain were uniquely perturbed by H3K9cr and H3K9ac, indicating a different chemical environment in the respective crotonyllysine and acetyllysine binding pockets (Supplementary Fig. 4a). These differences support our model that Trp81 adopts two conformations upon complex formation with the H3K9cr mark as compared to H3K9ac (Supplementary Figs. 1c, d and 4c). One of the conformations, characterized by the π stacking involving two aromatic residues and the alkene group, is observed only in the YEATS-H3K9cr complex. To establish whether the Taf14 YEATS domain is able to recognize other recently identified acyllysine marks, we performed solution pull-down assays using H3 peptides acetylated, propionylated, butyrylated, and crotonylated at lysine 9 (residues 1–20 of H3). As shown in Figure 2d and Supplementary Fig. 5a, the Taf14 YEATS domain binds more strongly to H3K9cr1-20, as compared to other acylated histone peptides. The preference for H3K9cr over H3K9ac, H3K9pr and H3K9bu was supported by 1H,15N HSQC titration experiments. Addition of H3K9ac1-20, H3K9pr1-20, and H3K9bu1-20 peptides caused chemical shift perturbations in the Taf14 YEATS domain in intermediate exchange regime, implying that these interactions are weaker compared to the interaction with the H3K9cr1-20 peptide (Supplementary Fig. 5b). We concluded that H3K9cr is the preferred target of this domain. From comparative structural analysis of the YEATS complexes, Gly80 emerged as candidate residue potentially responsible for the preference for crotonyllysine. In attempt to generate a mutant capable of accommodating a short acetyl moiety but discriminating against a longer, planar crotonyl moiety, we mutated Gly80 to more bulky residues, however all mutants of Gly80 lost their binding activities towards either acylated peptide, suggesting that Gly80 is absolutely required for the interaction. In contrast, mutation of Val24, a residue located on another side of Trp81, had no effect on binding (Fig. 2d and Supplementary Fig. 5a, c). To determine if the binding to crotonyllysine is conserved, we tested human YEATS domains by pull-down experiments using singly and multiply acetylated, propionylated, butyrylated, and crotonylated histone peptides (Supplementary Fig. 6). We found that all YEATS domains tested are capable of binding to crotonyllysine peptides, though they display variable preferences for the acyl moieties. While YEATS2 and ENL showed selectivity for the crotonylated peptides, GAS41 and AF9 bound acylated peptides almost equally well. Unlike the YEATS domain, a known acetyllysine reader, bromodomain, does not recognize crotonyllysine. We assayed a large set of BDs in pull-down experiments and found that this module is highly specific for acetyllysine and propionyllysine containing peptides (Supplementary Fig. 7). However, bromodomains did not interact (or associated very weakly) with longer acyl modifications, including crotonyllysine, as in the case of BDs of TAF1 and BRD2, supporting recent reports. These results demonstrate that the YEATS domain is currently the sole reader of crotonyllysine. In conclusion, we have identified the YEATS domain of Taf14 as the first reader of histone crotonylation. The unique and previously unobserved aromatic-amide/aliphatic-aromatic π-π-π-stacking mechanism facilitates the specific recognition of the crotonyl moiety. We further demonstrate that H3K9cr exists in yeast and is dynamically regulated by HATs and HDACs. As we previously showed the importance of acyllysine binding by the Taf14 YEATS domain for the DNA damage response and gene transcription, it will be essential in the future to define the physiological role of crotonyllysine recognition and to differentiate the activities of Taf14 that are due to binding to crotonyllysine and acetyllysine modifications. Furthermore, the functional significance of crotonyllysine recognition by other YEATS proteins will be of great importance to elucidate and compare. ONLINE METHODS Protein expression and purification The Taf14 YEATS constructs (residues 1–132 or 1–137) were expressed in E. coli BL21 (DE3) RIL in either Luria Broth or M19 minimal media supplemented with 15NH4Cl and purified as N-terminal GST fusion proteins. Cells were harvested by centrifugation and resuspended in 50 mM HEPES (pH 7.5) supplemented with 150 mM NaCl and 1 mM TCEP. Cells are lysed by freeze-thaw followed by sonication. Proteins were purified on glutathione Sepharose 4B beads and the GST tag was cleaved with PreScission protease. X-ray data collection and structure determination Taf14 YEATS (residues 1–137) was concentrated to 9 mg/mL in 25 mM MES (pH 6.5) and incubated with 2 molar equivalence of the H3K9cr5-13 at RT for 30 mins prior to crystallization. Crystals were obtain via sitting drop diffusion method at 18°C by mixing 800 nL of protein/peptide solution with 800 nL of well solution composed of 44% PEG600 (v/v) and 0.2 M citric acid (pH 6.0). X-ray diffraction data was collected at a wavelength of 1.54 Å at 100 K from a single crystal on the UC Denver Biophysical Core home source composed of a Rigaku Micromax 007 high frequency microfocus X-ray generator with a Pilatus 200K 2D area detector. HKL3000 was used for indexing, scaling, and data reduction. Solution was solved via molecular replacement with Phaser using the Taf14 YEATS domain (PDB 5D7E) as search model with waters, ligands, and peptide removed. Phenix was used for refinement of structure and waters were manually placed by inception of difference maps in Coot. Ramachandran plot indicates good stereochemistry of the three-dimensional structure with 100% of all residues falling within the favored (98%) and allowed (2%) regions. The crystallographic statistics are shown in Supplementary Table 1. NMR spectroscopy NMR spectroscopy was carried out on a Varian INOVA 600 MHz spectrometer outfitted with a cryogenic probe. Chemical shift perturbation (CSP) analysis was performed using uniformly 15N-labeled Taf14 (1–132). 1H,15N heteronuclear single quantum coherence (HSQC) spectra of the Taf14 YEATS domain were collected in the presence of increasing concentrations of either H3K9cr5-13, H3K9ac5-13, H3K9cr1-20, H3K9ac1-20 H3K9pr1-20, H3K9bu1-20 or free Kcr in PBS buffer pH 6.8, 8% D2O. Fluorescence binding assays Tryptophan fluorescence measurements were performed on a Fluorolog spectrofluorometer at room temperature as described. The samples containing 2 μM of Taf14 YEATS in PBS (pH 7.4) and increasing concentrations of H3K9cr5-13 were excited at 295 nm. Emission spectra were recorded from 310 to 340 nm with a 1 nm step size and a 0.5 sec integration time. The Kd value was determined using a nonlinear least-squares analysis and the equation:   where [L] is the concentration of the peptide, [P] is the concentration of the protein, ΔI is the observed change of signal intensity, and ΔImax is the difference in signal intensity of the free and bound states. The Kd values were averaged over 3 separate experiments, with error calculated as the standard deviation (SD). Peptide pull-downs YEATS domains in pGEX vectors were expressed in SoluBL21 cells (Amsbio) by induction with 1 mM IPTG at 16–18°C overnight with shaking. Cells were lysed by freeze-thaw and sonication then purified over glutathione agarose (Pierce) in a buffer containing 50 mM Tris pH 8.0, 500 mM NaCl, 20% glycerol (v/v) and 1 mM dithiothreitol (DTT). Peptide pull-downs were performed essentially as described except that the assay buffer contained 50 mM Tris pH 8.0, 500 mM NaCl, and 0.1% NP-40, and 500 pmols of biotinylated histone peptides were loaded onto streptavidin coated magnetic beads before incubation with 40 pmols of protein. Bound proteins were detected with rabbit GST antibody (Sigma, G7781). Point mutants were generated by site-directed mutagenesis and purified/assayed as described above. The YEATS domains of Taf14, AF9, ENL, and GAS41 were previously described. Western blotting Yeast cultures were grown in YPD media at 30°C to mid-log phase and extracts were prepared as previously described. Proteins from cell lysates were separated by SDS-PAGE and transferred to a PVDF membrane. Anti-H3K9ac (Millipore, 07-352) and anti-H3K9cr (PTM Biolabs, PTM-516) were diluted to 1:2000 and 1:1000, respectively, in 1x Superblock (ThermoScientific). An HRP-conjugated anti-rabbit (GE Healthcare) was used for detection. Bands were quantified using the ImageJ program. Dot blotting Increasing concentrations of biotinylated histone peptides (0.06–1.5 μg) were spotted onto a PVDF membrane then probed with the anti-H3K9ac (Millipore, 07-352) or H3K9cr (PTM Biolabs, PTM-516) at 1:2000 in a 5% non-fat milk solution and detected with an HRP-conjugated anti-rabbit by enhanced chemiluminesence (ECL). Bromodomains pull-downs cDNAs of GST-fused bromodomains were obtained either from EpiCypher Inc. or as a kind gift from Katrin Chua (Stanford University). GST fusions were expressed as described above except that the preparation buffer contained 50 mM Tris (pH 7.5), 150 mM NaCl, 10% glycerol (v/v), and 1 mM DTT. Pull-down assays were preformed as described above except that the assay buffer contained 50 mM Tris (pH 8.0), 300 mM NaCl, and 0.1% NP-40. Supplementary Material Accession codes. Coordinates and structure factors have been deposited in the Protein Data Bank under accession codes 5IOK. Author contributions F.H.A., S.A.S., E.K.S., J.B.B., A.G., I.K.T and K.K. performed experiments and together with X.S., B.D.S and T.G.K. analyzed the data. F.H.A., S.A.S., B.D.S. and T.G.K. wrote the manuscript with input from all authors. Competing Financial Interest The authors declare no competing financial interests. Additional information Any supplementary information is available in the online version of this paper. Identification of 67 histone marks and histone lysine crotonylation as a new type of histone modification Intracellular Crotonyl-CoA Stimulates Transcription through p300-Catalyzed Histone Crotonylation Protein lysine acylation and cysteine succination by intermediates of energy metabolism Identification of ‘erasers’ for lysine crotonylated histone marks using a chemical proteomics approach Perceiving the epigenetic landscape through histone readers Interpreting the language of histone and DNA modifications A Subset of Human Bromodomains Recognizes Butyryllysine and Crotonyllysine Histone Peptide Modifications Histone recognition and large-scale structural analysis of the human bromodomain family YEATS domain proteins: a diverse family with many links to chromatin modification and transcription AF9 YEATS domain links histone acetylation to DOT1L-mediated H3K79 methylation Association of Taf14 with acetylated histone H3 directs gene transcription and the DNA damage response Anc1 interacts with the catalytic subunits of the general transcription factors TFIID and TFIIF, the chromatin remodeling complexes RSC and INO80, and the histone acetyltransferase complex NuA3 Preparation and analysis of the INO80 complex TFG/TAF30/ANC1, a component of the yeast SWI/SNF complex that is similar to the leukemogenic proteins ENL and AF-9 The something about silencing protein, Sas3, is the catalytic subunit of NuA3, a yTAF(II)30-containing HAT complex that interacts with the Spt16 subunit of the yeast CP (Cdc68/Pob3)-FACT complex The essential role of acetyllysine binding by the YEATS domain in transcriptional regulation Phaser crystallographic software PHENIX: a comprehensive Python-based system for macromolecular structure solution Features and development of Coot Molecular basis for chromatin binding and regulation of MLL5 Association of UHRF1 with methylated H3K9 directs the maintenance of DNA methylation Association of Taf14 with acetylated histone H3 directs gene transcription and the DNA damage response The Saccharomyces cerevisiae histone H2A variant Htz1 is acetylated by NuA4 A phosphatase complex that dephosphorylates gammaH2AX regulates DNA damage checkpoint recovery The structural mechanism for the recognition of H3K9cr (a) Chemical structure of crotonyllysine. (b) The crystal structure of the Taf14 YEATS domain (wheat) in complex with the H3K9cr5-13 peptide (green). (c) H3K9cr is stabilized via an extensive network of intermolecular electrostatic and polar interactions with the Taf14 YEATS domain. (d) The π-π-π stacking mechanism involving the alkene moiety of crotonyllysine. H3K9cr is a selective target of the Taf14 YEATS domain (a, b) Western blot analysis comparing the levels of H3K9cr and H3K9ac in wild type (WT), HAT deletion, or HDAC deletion yeast strains. Total H3 was used as a loading control. (c) Superimposed 1H,15N HSQC spectra of Taf14 YEATS recorded as H3K9cr5-13 and H3K9ac5-13 peptides were titrated in. Spectra are color coded according to the protein:peptide molar ratio. (d) Western blot analyses of peptide pull-down assays using wild-type and mutated Taf14 YEATS domains and indicated 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+[{"sourceid":"4885502","sourcedb":"","project":"","target":"","text":"Using Cryo-EM to Map Small Ligands on Dynamic Metabolic Enzymes: Studies with Glutamate Dehydrogenase Cryo-electron microscopy (cryo-EM) methods are now being used to determine structures at near-atomic resolution and have great promise in molecular pharmacology, especially in the context of mapping the binding of small-molecule ligands to protein complexes that display conformational flexibility. We illustrate this here using glutamate dehydrogenase (GDH), a 336-kDa metabolic enzyme that catalyzes the oxidative deamination of glutamate. Dysregulation of GDH leads to a variety of metabolic and neurologic disorders. Here, we report near-atomic resolution cryo-EM structures, at resolutions ranging from 3.2 Å to 3.6 Å for GDH complexes, including complexes for which crystal structures are not available. We show that the binding of the coenzyme NADH alone or in concert with GTP results in a binary mixture in which the enzyme is in either an “open” or “closed” state. Whereas the structure of NADH in the active site is similar between the open and closed states, it is unexpectedly different at the regulatory site. Our studies thus demonstrate that even in instances when there is considerable structural information available from X-ray crystallography, cryo-EM methods can provide useful complementary insights into regulatory mechanisms for dynamic protein complexes. Introduction Recent advances in cryo-electron microscopy (cryo-EM) allow determination of structures of small protein complexes and membrane proteins at near-atomic resolution, marking a critical shift in the structural biology field. One specific area of broad general interest in drug discovery is the localization of bound ligands and cofactors under conditions in which efforts at crystallization have not been successful because of structural heterogeneity. Recent cryo-EM analyses have already demonstrated that it is now possible to use single-particle cryo-EM methods to localize small bound ligands or inhibitors on target proteins. Whether ligand binding can be visualized at high resolution is an important question, even in the more general case when multiple conformations are present simultaneously. Here, we address this question using mammalian glutamate dehydrogenase as an example. Glutamate dehydrogenase (GDH) is a highly conserved enzyme expressed in most organisms. GDH plays a central role in glutamate metabolism by catalyzing the reversible oxidative deamination of glutamate to generate α-ketoglutarate and ammonia, with the concomitant transfer of a pair of electrons to either NAD+ or NADP+. Regulation of GDH is tightly controlled through multiple allosteric mechanisms. Extensive biochemical and crystallographic studies have characterized the enzymatic activity of GDH and its modulation by a chemically diverse group of compounds such as nucleotides, amino acids, steroid hormones, antipsychotic drugs, and natural products. X-ray crystallographic studies have shown that the functional unit of GDH is a homohexamer composed of a trimer of dimers, with a 3-fold axis and an equatorial plane that define its D3 symmetry (Fig. 1A). Each 56-kDa protomer consists of three domains. The first is located near the dimer interface and forms the core of the hexamer. The second, a nucleotide-binding domain (NBD) with a Rossmann fold, defines one face of the catalytic cleft bounded by the core domain. During the catalytic cycle, the NBD executes a large movement, hinged around a “pivot” helix, that closes the catalytic cleft, and drives a large conformational change in the hexamer from open to closed states (Fig. 1B). The third domain, dubbed the “antenna,” is an evolutionary acquisition in protista and animals. Antennae of adjacent protomers in each trimer intercalate to form a bundle, perpendicular to the pivot helices, that protrudes along the distal extremes of the 3-fold axis. When a protomer undergoes a conformational change, the rotation of its pivot helix is transferred through the antenna to the adjacent subunit. The influence of the antenna, present only in protozoan and metazoan enzymes, has been proposed to explain its cooperative behavior, which is absent in bacterial homologs. Deletion of this domain leads to loss of cooperativity. Structure and quaternary conformational changes in GDH. (A) Views of open (PDB ID 1NR7) and closed (PDB 3MW9) states of the GDH hexamer, shown in ribbon representation perpendicular to the 2-fold symmetry axis (side view, top) and 3-fold symmetry axis (top view, bottom). Only three protomers are shown in the top view for purposes of visual clarity. The dashed lines and arrows, respectively, highlight the slight extension in length, and twist in shape that occurs with transition from open to the closed state. The open state shown is for unliganded GDH, whereas the closed state has NADH, GTP, and glutamate bound. (B) Superposition of structures for closed and open conformations, along with a series of possible intermediate conformations along the trajectory that serve to illustrate the extent of change in structure across different regions of the protein. The transition between “closed” and “open” states of GDH is modulated by two allosteric sites in each protomer (Fig. 1A), which are differentially bound by GTP (an inhibitor) and ADP (an activator). These allosteric modulators tightly control GDH function in vivo. In the first site, which sits next to the pivot helix at the base of the antenna (the “GTP binding site”), GTP binding is known to act as an inhibitor, preventing release of the reaction product from the catalytic site by stabilizing the closed conformation of the catalytic cleft. In the second “regulatory site”, which is situated near the pivot helix between adjacent protomers, ADP acts as an activator of enzymatic activity, presumably by hastening the opening of the catalytic cleft that leads to the release of the reaction product. Interestingly, it has also been shown that the coenzyme NADH can bind to the regulatory site (also bound by the activator ADP), exerting a converse, inhibitory effect on GDH product release, although the role this may play in vivo is not entirely clear. Although there are numerous crystal structures available for GDH in complex with cofactors and nucleotides, they are limited to the combinations that have been amenable to crystallization. Nearly all X-ray structures of mammalian GDH are in the closed conformation, and the few structures that are in the open conformation are at lower resolution (Table 1). Of those structures in the closed conformation, most include NAD[P]H, GTP, and glutamate (or, alternately, NAD+, GTP, and α-ketoglutarate). However, the effects of coenzyme and GTP, bound alone or in concert in the absence of glutamate, have not been analyzed by crystallographic methods. Here, we report single-particle cryo-electron microscopy (cryo-EM) studies that show that under these conditions enzyme complexes coexist in both closed and open conformations. We show that the structures in both states can be resolved at near-atomic resolution, suggesting a molecular mechanism for synergistic inhibition of GDH by NADH and GTP (see Table 2 for detailed information on all cryo-EM-derived structures that we report in this work). X-ray structures of mammalian GDH reported in both the open and closed conformations GDH\tLigands\tPDB ID\tConformation\tResolution\t \tWT\tNADH + GLU + GTP\t3MW9\tClosed\t2.4\t \tWT\tGlu, GTP, NADPH, and Bithionol\t3ETD\tClosed\t2.5\t \tWT\tGlu, NADPH, GTP + GW5074\t3ETG\tClosed\t2.5\t \tWT\tapo\t1L1F\tOpen\t2.7\t \tWT\tNADPH, glutamate, and GTP\t1HWZ\tClosed\t2.8\t \tWT\tNADPH + GLU + GTP + Zinc\t3MVQ\tClosed\t2.94\t \tWT\tNADPH, Glu, GTP, Hexachlorophene\t3ETE\tClosed\t3\t \tWT\tNAD, PO4, and 2-oxoglutarate\t1HWY\tClosed\t3.2\t \tWT\tNADPH + GLU + Eu\t3MVO\tClosed\t3.23\t \tR463A mutant\tapo\t1NR1\tOpen\t3.3\t \tWT\tapo\t1NR7\tOpen\t3.3\t \tWT\tADP\t1NQT\tOpen\t3.5\t \tWT\tNADPH and Epicatechin-3-gallate (Ecg)\t3QMU\tOpen\t3.62\t \t Cryo-EM structures of mammalian GDH determined for this study GDH\tLigands\tEMDB ID\tPDB ID\tConformation\tResolution\tParticles\t \tWT\tapo\tEMD-6630\t3JCZ\tOpen\t3.26\t22462\t \tWT\tGTP\tEMD-6631\t3JD0\tOpen\t3.47\t39439\t \tWT\tNADH\tEMD-6635\t3JD2\tOpen\t3.27\t34716\t \tWT\tNADH\tEMD-6634\t3JD1\tClosed\t3.27\t34926\t \tWT\tNADH + GTP\tEMD-6632\t3JD3\tOpen\t3.55\t14793\t \tWT\tNADH + GTP\tEMD-6633\t3JD4\tClosed\t3.40\t20429\t \t Materials and Methods Specimen Preparation. Bovine glutamate dehydrogenase (Enzyme Commission 1.4.1.3; Sigma-Aldrich/MilliporeSigma, St. Louis, MO) was dialyzed overnight against fractionation buffer (100 mM potassium phosphate, pH 6.8) prior to fractionation by size-exclusion chromatography using a Superdex 200 10/30 column connected to an ÄKTA FPLC apparatus (GE Healthcare Bio-Sciences, Piscataway, NJ ). The concentration of GDH was adjusted to ∼2 mg/ml by rapid mixing with potassium phosphate buffer containing the concentration of ligand as necessary and with n-octyl glucopyranoside at a final concentration of 0.1%. The final concentration of each ligand was 20 mM. Small volumes of sample, typically 3 µl, were deposited on 200 mesh Quantifoil R2/2 grids (Quantifoil Micro Tools, Großlöbichaum, Germany), blotted, and plunge-frozen in liquid ethane using an FEI Vitrobot Mark IV (FEI Company, Hillsboro, OR). Frozen grids were mounted into autoloader cartridges and transferred to the microscope. Cryo-Electron Microscopy. Specimens were imaged on an FEI Titan Krios microscope (FEI Company) aligned for parallel illumination and operated at 300 kV. The instrument was furnished with a Gatan K2 Summit camera placed at the end of a GIF Quantum energy filter (Gatan Inc., Pleasanton, CA), operated in zero-energy-loss mode with a slit width of 20 eV. Images were collected manually at a dose rate of ∼5 e– pixel−1 s−1, i.e., in the linear range of the detector. The physical pixel size at the plane of the specimen was 1.275 Å, corresponding to a super-resolution pixel size of 0.6375 Å. The total exposure time was 15.2 s, and intermediate frames were recorded every 0.4 s, giving an accumulated dose of ∼45 e–/Å2 and a total of 38 frames per image. The majority of images were collected at under-focus values between 1 μm and 3 μm. Data Processing. Drift and beam-induced motion were compensated by whole-frame alignment of movies, and CTF was estimated as described in. Integrated frames were manually examined and selected on the basis of the quality of the CTF estimation, astigmatism, drift, and particle distribution. Molecular images were automatically identified from selected integrated micrographs by detecting the local maxima of correlation of each image with a Gaussian disk of 150 Å in radius. Individual particle projections were extracted from integrated super-resolution images using a binning factor of 4 and a box size of 96 × 96 pixels and assigned into 20–100 groups by iterative reference free two-dimensional classification as implemented in EMAN2. Following 8 iterations of classification, a subset of classes depicting intact particles were used to build symmetric (D3) density maps using the program e2initialmodel.py from the EMAN2 suite. One or more maps were selected as reference for further processing on the basis of consistency between projections and the original classes. These “initial models” were refined to ∼15–20 Å using e2refine_easy.py. Unbinned particles were then re-extracted from the original super-resolution images using a binning of 2 and a box size of 384 × 384, and subject to classification in three dimensions using the maximum likelihood method implemented in RELION (; MRC Laboratory, Cambridge, UK) (regularization parameter of T=4). Unless noted otherwise, D3 symmetry was imposed for three-dimensional classification runs, the number of classes was initially determined on the basis of the number of particles included in the analysis and later adjusted on the basis of the number of conformations detected in the sample (see Supplemental Table 1 for details). Iteration over classification in three dimensions was continued until convergence as judged by resolution and distribution of particles among the classes. Particles belonging to “good” three-dimensional classes were pooled into one or more classes, depending on the conformational landscape of the complex, and refined using the “gold standard” method in RELION. The refined maps were corrected for the MTF of the camera and for B-factor in the framework of RELION. Figures were generated using UCSF Chimera and Maxon Cinema4D (Maxon Computer Inc., Newbury Park, CA), and two-dimensionally composited in Adobe Photoshop and Illustrator. Building of Atomic Models. The deposited models for unliganded GDH (1NR7), the binary complex with ADP (1NQT), and the quaternary complex with NADH, GTP, and Glu (3MW9) were used to derive models from the six structures reported here. Conflicts in sequence between the deposited models were solved by conforming to the primary sequence as reported in 3MW9. The models were placed in the corresponding map (open or closed) by rigid body fitting as implemented in Chimera. In the closed state, in which all relative orientations of the ligands are known, only the ligands known to be present in each structure were retained, all other nonstandard residues were deleted. For the open state, ligands were initially placed on the basis of their orientation relative to the corresponding binding site. The models were refined against a map derived from one-half of the dataset using Rosetta as described (https://faculty.washington.edu/dimaio/files/density_tutorial.pdf), followed by real space refinement in PHENIX (; PHENIX, Berkeley, CA) and evaluated by calculating the Fourier shell correlation between the model and the map derived from the second half of the dataset. For each complex, ten best scoring instances on the basis of the Fourier shell correlation were selected among one hundred runs, visually examined, and the one deemed best interactively corrected in Coot. Each corrected model was subject to a final instance of real space refinement using PHENIX. Results and Discussion To explore the conformational landscape of apo-GDH, we first determined its structure in the absence of any added ligands (Supplemental Fig. 1, Fig. 2, A–C). The density map, refined to an average resolution of ∼3.0 Å (Supplemental Fig. 2), is in the open conformation and closely matches the model of unliganded GDH derived by X-ray crystallography at 3.3 Å resolution (PDB ID 1NR7). The variation in local resolution from the core to the periphery, as reported by ResMap (Supplemental Fig. 3D), is consistent with the B-factor gradient observed in the crystal structure (Supplemental Fig. 3A). Extensive classification without imposing symmetry yielded only open structures and failed to detect any closed catalytic cleft in the unliganded enzyme, suggesting that all six protomers are in the open conformation. Consistent with this conclusion, the loops connecting the β-strands of the Rossmann fold are well-defined (Fig. 2B), implying that there is little movement at the NBD, as the transition between closed and open states is associated with NBD movement (Fig. 1B). Cryo-EM structures of GDH in unliganded and NADH-bound states. (A) Refined cryo-EM map of unliganded GDH at ∼3 Å resolution. (B, C) Illustration of density map in the regions that contain the Rossmann nucleotide binding fold (B), pivot and antenna helices (C) in the unliganded GDH map. (D) Cryo-EM-derived density maps for two coexisting conformations that are present when GDH is bound to the cofactor NADH. Each protomer is shown in a different color and densities for NADH bound in both regulatory (red) and catalytic (purple) sites on one protomer are indicated. The overall quaternary structures of the two conformations are essentially the same as that of the open and closed states observed by X-ray crystallography. When GDH is bound to NADH, GTP, and glutamate, the enzyme adopts a closed conformation; this “abortive complex” has been determined to 2.4-Å resolution by X-ray crystallography (PDB 3MW9). However, crystal structures of GDH bound only to NADH or to GTP have not yet been reported. To test the effect of NADH binding on GDH conformation in solution, we determined the structure of this binary complex using cryo-EM methods combined with three-dimensional classification. Two dominant conformational states, in an all open or all closed conformation were detected, segregated (Fig. 2D), and further refined to near-atomic resolution (∼3.3 Å; Supplemental Fig. 2). Densities for 12 molecules of bound NADH were identified in maps of both open and closed states (Supplemental Fig. 4). The NADH-bound closed conformation matches the structure of the quaternary complex observed by X-ray crystallography, with the exception that density corresponding to GTP and glutamate was absent in the cryo-EM-derived map. Comparison of the NADH-bound closed conformation to the NADH-bound open conformation shows that, as expected, the catalytic cleft is closed and the NBDs are displaced toward the equatorial plane, accompanied by a rotation of the pivot helix by ∼7°, concomitant with a large conformational change in the antennae domains (Figs. 1 and 2D). A comparison between NADH-bound open and closed conformations also involves a displacement of helix 5 (residues 171–186), as well as a tilt of the core β-sheets relative to the equatorial plane of the enzyme (residues 57–97, 122–130) and α-helix 2 (residues 36–54), and a bending of the N-terminal helix. Thus, closure of the catalytic cleft is accompanied by a quaternary structural change that can be described as a global bending of the structure about an axis that runs parallel to the pivot helix, accompanied by an expansion of the core (Figs. 1A and 2D). Detailed analysis of the GDH/NADH structures shows that both the adenosine and nicotinamide moieties of NADH bind to the catalytic site within the NBD in nearly the same orientation in both the open and the closed states, and display closely comparable interactions with the Rossmann fold (Fig. 3, A and B). At the regulatory site, where either ADP can bind as an activator or NADH can bind as an inhibitor, the binding of the adenine moiety of NADH is nearly identical between the two conformers. However, there is a significant difference in the orientation of the nicotinamide and phosphate moieties in the two conformational states (Fig. 3, C and D). In the closed state, the nicotinamide group is oriented toward the center of the hexamer, inserted into a narrow cavity between two adjacent subunits in the trimer. There are extensive interactions between NADH and the residues lining this cavity, which may explain the well-defined density of this portion of NADH in the closed state. In contrast, in the open conformation, the cavity present in the closed state becomes too narrow for the nicotinamide group; instead, the group is oriented in the opposite direction, parallel to the pivot helix with the amido group extending toward the C-terminal end of the helix. Detailed view of NADH conformation in catalytic and regulatory sites. (A, B) NADH density (purple) and interactions in the catalytic sites of closed (A) and open (B) states. (C, D) NADH density (red) and interactions in the regulatory sites of closed (C) and open (D) states. Although there is a difference in orientation of the nicotinamide moiety between the closed and open states in the regulatory site, in both structures the adenine portion of NADH has a similar binding pocket and is located in almost exactly the same position as ADP, a potent activator of GDH function (Supplemental Fig. 5). In the open state, the binding of ADP or NADH is further stabilized by His209, a residue that undergoes a large movement during the transition from open to closed conformation (Fig. 3, C and D). In the open conformation, the distance between His209 and the α-phosphate of NADH is ∼4.4 Å, which is comparable with the corresponding distance in the ADP-bound conformation. In the closed conformation, however, this key histidine residue is \u003e10.5 Å away from the nearest phosphate group on NADH, altering a critical stabilization point within the regulatory site. This suggests that although the conformation of NADH in the open state regulatory site more closely mimics the binding of ADP, the conformation of NADH in the closed state regulatory site is significantly different; these differences may contribute to the opposite effects of NADH and ADP on GDH enzymatic activity. In the absence of NADH, GTP binds weakly to GDH with a dissociation constant of ∼20 μM. Cryo-EM analysis of GDH incubated with GTP resulted in a structure at an overall resolution of 3.5 Å, showing that it is in an open conformation (Supplemental Fig. 6), with all NBDs in the open state. The density for GTP is not very well defined, suggesting considerable wobble in the binding site. Subtraction of the GTP-bound map with that of the apo state shows that GTP binding can nevertheless be visualized specifically in the GTP binding site (Supplemental Fig. 6). Importantly, the binding of GTP alone does not appear to drive the transition from the open to the closed state of GDH. To further dissect the roles of NADH and GTP in the transition from the open to closed conformations, we next determined structures of GDH in complex with both NADH and GTP, but without glutamate. When NADH and GTP are both present, classification reveals the presence of both closed and open GDH conformations, similar to the condition when only NADH is present (Fig. 4, A and B). Reconstruction without classification, however, yields a structure clearly in the closed conformation, suggesting that, in coordination with NADH, GTP may further stabilize the closed conformation. The location of GTP in the open and closed states of the GDH/NADH/GTP complex is similar to that in the crystal structure observed in the presence of NADH, GTP, and glutamate. Likewise, the position of NADH in the open and closed states closely resembles the position of NADH in the GDH/NADH open and closed structures. One key difference between the open and closed states of these structures is the position of the His209 residue: As mentioned above, His209 swings away from the adenine moiety of NADH in the closed state. When GTP is present in the GTP binding site, His209 instead interacts with GTP, probably stabilizing the closed conformation (Fig. 4, C and D). Thus, GTP binding to GDH appears synergistic with NADH and displaces the conformational landscape toward the closed state. Cryo-EM structure of GDH bound to both NADH and GTP. (A, B) Observation of co-existing open (A) and closed (B) conformations in the GDH-NADH-GTP ternary complex. Densities for GTP (yellow) as well as NADH bound to both catalytic (purple) and regulatory (red) sites in each protomer are shown. (C, D) Detailed inspection of the interactions near the regulatory site show that the orientation of His209 switches between the two states, which may allow interactions with bound GTP in the closed (D), but not open (C) conformation. Our structural studies thus establish that whether or not GTP is bound, NADH binding is detectable at catalytic and regulatory sites, in both the open and closed conformational states. Whereas the orientation in which NADH binds at the catalytic site is similar for both conformations, the orientation of the nicotinamide portion of NADH in the regulatory site is different between the open and closed conformations (Figs. 3 and 4). In the closed state, the nicotinamide moiety is inserted into a well-defined cavity at the interface between two adjacent protomers in the trimer. As mentioned above, this cavity is much narrower in the open state, suggesting that this cavity may be unavailable to the NADH nicotinamide moiety when the enzyme is in the open conformation. These structural features provide a potential explanation of the weaker density for the nicotinamide moiety of NADH in the open state, and may account for the higher reported affinity of NADH for the closed state. The role of the nicotinamide moiety in acting as a wedge that prevents the transition to the open conformation also suggests a structural explanation of the mechanism by which NADH binding inhibits the activity of the enzyme by stabilizing the closed conformation state. The rapid emergence of cryo-EM as a tool for near-atomic resolution structure determination provides new opportunities for complementing atomic resolution information from X-ray crystallography, as illustrated here with GDH. Perhaps the most important contribution of these methods is the prospect that when there are discrete subpopulations present, the structure of each state can be determined at near-atomic resolution. What we demonstrate here with GDH is that by employing three-dimensional image classification approaches, we not only can isolate distinct, coexisting conformations, but we can also localize small molecule ligands in each of these conformations. These kinds of approaches will probably become increasingly important in molecular pharmacology, especially in the context of better understanding drug-target interactions in dynamic protein complexes. Supplementary Material This research was supported by funds from the  National Cancer Institute Center for Cancer Research, the IATAP program at NIH, and the NIH-FEI Living Laboratory for Structural Biology (S.S., J.L.S.M.). This work was supported by the Intramural Research Program of the National Institutes of Health National Cancer Institute. dx.doi.org/10.1124/mol.116.103382. This article has supplemental material available at molpharm.aspetjournals.org. Abbreviations cryo-EM cryo-electron microscopy GDH glutamate dehydrogenase NBD nucleotide binding domain Authorship Contributions Participated in research design: Borgnia, Banerjee, Merk, Subramaniam, Milne. Conducted experiments: Borgnia, Banerjee, Merk, Rao, Pierson. Performed data analysis: Borgnia, Banerjee, Merk, Matthies, Bartesaghi, Earl, Falconieri, Subramaniam, Milne. Wrote or contributed to the writing of the manuscript: Borgnia, Banerjee, Earl, Falconieri, Subramaniam, Milne. References PHENIX: a comprehensive Python-based system for macromolecular structure solution Evolution of glutamate dehydrogenase regulation of insulin homeostasis is an example of molecular exaptation Regulation of bovine glutamate dehydrogenase. The effects of pH and ADP Structural studies on ADP activation of mammalian glutamate dehydrogenase and the evolution of regulation Structure of β-galactosidase at 3.2-Å resolution obtained by cryo-electron microscopy Inhibition of ox brain glutamate dehydrogenase by perphenazine Studies of glutamate dehydrogenase. The interaction of ADP, GTP, and NADPH in complexes with glutamate dehydrogenase Features and development of Coot Glutamate Dehydrogenase. V. 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annotated_BioC_JSON/PMC4888278_ann.json

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+[{"sourceid":"4888278","sourcedb":"","project":"","target":"","text":"Structural determinant for inducing RORgamma specific inverse agonism triggered by a synthetic benzoxazinone ligand Background The nuclear hormone receptor RORγ regulates transcriptional genes involved in the production of the pro-inflammatory interleukin IL-17 which has been linked to autoimmune diseases such as rheumatoid arthritis, multiple sclerosis and inflammatory bowel disease. This transcriptional activity of RORγ is modulated through a protein-protein interaction involving the activation function 2 (AF2) helix on the ligand binding domain of RORγ and a conserved LXXLL helix motif on coactivator proteins. Our goal was to develop a RORγ specific inverse agonist that would help down regulate pro-inflammatory gene transcription by disrupting the protein protein interaction with coactivator proteins as a therapeutic agent. Results We identified a novel series of synthetic benzoxazinone ligands having an agonist (BIO592) and inverse agonist (BIO399) mode of action in a FRET based assay. We show that the AF2 helix of RORγ is proteolytically sensitive when inverse agonist BIO399 binds. Using x-ray crystallography we show how small modifications on the benzoxazinone agonist BIO592 trigger inverse agonism of RORγ. Using an in vivo reporter assay, we show that the inverse agonist BIO399 displayed specificity for RORγ over ROR sub-family members α and β. Conclusion The synthetic benzoxazinone ligands identified in our FRET assay have an agonist (BIO592) or inverse agonist (BIO399) effect by stabilizing or destabilizing the agonist conformation of RORγ. The proteolytic sensitivity of the AF2 helix of RORγ demonstrates that it destabilizes upon BIO399 inverse agonist binding perturbing the coactivator protein binding site. Our structural investigation of the BIO592 agonist and BIO399 inverse agonist structures identified residue Met358 on RORγ as the trigger for RORγ specific inverse agonism. Electronic supplementary material The online version of this article (doi:10.1186/s12900-016-0059-3) contains supplementary material, which is available to authorized users. Background Retinoid-related orphan receptor gamma (RORγ) is a transcription factor belonging to a sub-family of nuclear receptors that includes two closely related members RORα and RORβ. Even though a high degree of sequence similarity exists between the RORs, their functional roles in regulation for physiological processes involved in development and immunity are distinct. During development, RORγ regulates the transcriptional genes involved in the functioning of multiple pro-inflammatory lymphocyte lineages including T helper cells (TH17cells) which are necessary for IL-17 production. IL-17 is a pro-inflammatory interleukin linked to autoimmune diseases such as rheumatoid arthritis, multiple sclerosis and inflammatory bowel disease; making its transcriptional regulation through RORγ an attractive therapeutic target. RORγ consists of an N-terminal DNA binding domain (DBD) connected to a C-terminal ligand binding domain (LBD) via a flexible hinge region. The DBD is composed of two zinc fingers that allow it to interact with specifically encoded regions on the DNA called the nuclear receptor response elements. The LBD consists of a coactivator protein binding pocket and a hydrophobic ligand binding site (LBS) which are responsible for regulating transcription. The coactivator binding pocket of RORγ recognizes a conserved helix motif LXXLL (where X can be any amino acid) on transcriptional coactivator complexes and recruits it to activate transcription. Like other nuclear hormone receptors, RORγ’s helix12 which makes up the C-termini of the LBD is an essential part of the coactivator binding pocket and is commonly referred to as the activation function helix 2 (AF2). In RORγ, the conformation of the AF2 helix required to form the coactivator binding pocket is mediated by a salt bridge between His479 and Tyr502 in addition to π- π interactions between Tyr502 and Phe506. The conformation of the AF2 helix can be modulated through targeted ligands which bind the LBS and increase the binding of the coactivator protein (agonists) or disrupt binding (inverse agonists) thereby enhancing or inhibiting transcription. Since RORγ has been demonstrated to play an important role in pro-inflammatory gene expression patterns implicated in several major autoimmune diseases, our aim was to develop RORγ inverse agonists that would help down regulate pro-inflammatory gene transcription. FRET results for agonist BIO592 (a) and Inverse Agonist BIO399 (b) Here we present the identification of two synthetic benzoxazinone RORγ ligands, a weak agonist BIO592 (Fig. 1a) and an inverse agonist BIO399 (Fig. 1b) which were identified using a Fluorescence Resonance Energy transfer (FRET) based assay that monitored coactivator peptide recruitment. Using partial proteolysis in combination with mass spectrometry analysis we demonstrate that the AF2 helix of RORγ destabilizes upon BIO399 (inverse agonist) binding. Finally, comparing binding modes of our benzoxazinone RORγ crystal structures to other ROR structures, we hypothesize a new mode of action for achieving inverse agonism and selectivity. Methods Cloning, protein expression and purification of RORγ518 GST-RORγ518 was constructed by sub-cloning residues 259 to 518 of a human RORγ cDNA into a pGEX-6P vector with a cleavable N-terminal GST fusion tag. BL21 (DE3) Escherichia coli cells were transformed with the plasmid encoding the GST-PreScission-hRORgamma 259–518 protein (GST-RORγ518) and were grown at 37 °C in LB media supplemented with ampicillin to an OD of 1. The temperature was reduced to 18 °C and protein expression was induced by adding 1 mM IPTG and was shaking for an additional 16 h. The cells were harvested and resuspended in lysis buffer (25 mM TRIS pH 8.0, 250 mM NaCl, 10 % Glycerol, 5 mM DTT and Roche EDTA-free protease inhibitor cocktail) and were lysed using a microfluidizer. The lysate was clarified by centrifugation at 20,000 × g for 1 h at 4 °C and GST-RORγ518 was captured by batch binding to Glutathione Sepharose resin overnight at 4 °C. The resin was washed with buffer A (25 mM TRIS pH 8.0, 250 mM NaCl, 10 % glycerol, 5 mM DTT) and loaded onto a XK column and washed until no non-specific unbound protein was detected. GST- RORγ518 was eluted from the column using buffer A supplemented with 10 mM Glutathione pH 8.0 and analyzed by SDS-PAGE. The eluate was then treated with PreScission Protease (10units/mg of protein) and further purified on a Superdex 75 column equilibrated in buffer B (25 mM TRIS pH 8.0, 250 mM NaCl, 5 % glycerol and 2 mM DTT). RORγ518 eluted as a monomer and was approximately 95 % pure as observed by SDS-PAGE. Additional constructs including c-terminal truncations, surface entropy reduction and cysteine scrubbed mutations were also expressed and purified in the same manner as RORγ518 if an expression level of \u003e1 mg/L was achieved. RORγ FRET based assay and GAL4 reporter assay FRET-based (Fluorescence Resonance Energy Transfer) assay and the GAL4 Reporter assay were performed as described previously. BIO592 and BIO399 were synthesized (Additional file 1) and belonged to a proprietary library where they were identified as RORγ activity modulators using the FRET-based assay. Partial proteolysis of RORγ518 RORγ518 at 8 mg/ml or in complex with 1 mM BIO399 or 1 mM BIO592 and 0.5 mM coactivator peptide EBI96 EFPYLLSLLGEVSPQ (New England Peptide) were treated with Actinase E (Hampton Research) added at a ratio of 1.25ugs of protease/1 mg of RORγ518 for 6 h at 4 °C. The reactions were quenched using 1X Protease inhibitor cocktail (Roche) + 1 mM EDTA and subjected to mass spectrometry analysis. Mass spectrometry of partially proteolyzed RORγ518 Proteolyzed RORγ518 samples were reduced with 50 mM dithiothreitol in 50 mM Tris pH 8.0, 150 mM NaCl containing 4 M urea and 5 mM EDTA. The sample was then analyzed on a LC-MS system comprised of a UPLC (ACQUITY, Waters Corp.), a TUV dual-wavelength UV detector (Waters Corp.), and a ZQ mass spectrometer (Waters Corp.). A Vydac C4 cartridge was used for desalting. Molecular masses for the Actinase E treated RORγ518 samples were obtained by deconvoluting the raw mass spectra using MaxLynx 4.1 software (Waters Corp.). Crystallization of RORγ518 with agonist BIO592 and inverse agonist BIO399 RORγ518 was concentrated to 8 mg/ml and EBI96 was added to a final concentration of 0.5 mM and agonist BIO592 to 1 mM and incubated on ice for 1 h. The coactivator peptide EBI96 which was identified by phage display was chosen for crystallization because of its strong interaction with RORγ in a mammalian two-hybrid analysis system that assessed the transactivation of RORγ. Diffraction quality crystals were grown through vapor diffusion in a buffer containing 0.1 M HEPES pH 8.0, 25 % PEG3350 and 0.2 M NaCl at 18 °C. Crystals were cryoprotected in the mother liquor containing 20 % glycerol as cryoprotectant prior to being frozen in liquid nitrogen for data collection. Actinase E proteolyzed RORγ518 BIO399 concentrated to 8 mg/ml was crystallized using vapor diffusion in a buffer containing 0.1 M BisTRIS pH 5.5, 0.2 M ammonium acetate and 15 % PEG3350 at 18 °C. Crystals were cryoprotected for data collection by transferring them to a mother liquor containing 15 % PEG400 prior to being frozen in liquid nitrogen. Data collection and structure determination for RORγ518 BIO592 and BIO399 complexes X-ray diffraction data for all the crystals were measured at beam line ID31 at the Argonne Photon Source. The data were processed with Mosflm in case of the RORγ518-BIO592-EBI96 ternary complex and with HKL2000 in the case of the Actinase E treated aeRORγ518/BIO399 complex. For both datasets, PDB ID: 3LOL was used as the search model, and the molecular replacement solutions were determined using MOLREP. The refinement was carried out using Refmac5 and model building was carried out in Coot. The data processing and refinement statistics are provided in Additional file 2. RORγ518-BIO592-EBI96 ternary complex: The data for the ternary complex were measured to 2.63 Å. It crystallized in a P21 space group with four molecules of the ternary complex in the asymmetric unit. The final model was refined to a Rcryst of 19.9 % and Rfree of 25.5 %. aeRORγ518/BIO399 complex: Diffraction data for the aeRORγ518-BIO399 complex were measured to 2.35 Å. It crystallized in C2 space group with two molecules in the asymmetric unit. The final model was refined to a Rcryst of 21.1 % and Rfree of 26.3 %. Results and discussion Identification of BIO592 and BIO399 as ligands that modulate RORγ coactivator peptide recruitment Using a FRET based assay we discovered agonist BIO592 (Fig. 1a) which increased the coactivator peptide TRAP220 recruitment to RORγ (EC50 0f 58nM and Emax of 130 %) and a potent inverse agonist BIO399 (Fig. 1b) which inhibited coactivator recruitment (IC50: 4.7nM). Interestingly, the structural difference between the agonist BIO592 and inverse agonist BIO399 was minor; with the 2,3-dihydrobenzo[1,4]oxazepin-4-one ring system of BIO399 being 3 atoms larger than the benzo[1,4]oxazine-3-one ring system of BIO592. In order to understand how small changes in the core ring system leads to inverse agonism, we wanted to structurally determine the binding mode of both BIO592 and BIO399 in the LBS of RORγ using x-ray crystallography. Structure of the RORγ518-BIO592-EBI96 ternary complex is in a transcriptionally active conformation \na The ternary structure of RORγ518 BIO592 and EBI96. b RORγ AF2 helix in the agonist conformation. c EBI96 coactivator peptide bound in the coactivator pocket of RORγ RORγ518 bound to agonist BIO592 was crystallized with a truncated form of the coactivator peptide EBI96 to a resolution of 2.6 Å (Fig. 2a). The structure of the ternary complex had features similar to other ROR agonist coactivator structures in a transcriptionally active canonical three layer helix fold with the AF2 helix in the agonist conformation. The agonist conformation is stabilized by a hydrogen bond between His479 and Tyr502, in addition to π-π interactions between His479, Tyr502 and Phe506 (Fig. 2b). The hydrogen bond between His479 and Tyr502 has been reported to be critical for RORγ agonist activity. Disrupting this interaction through mutagenesis reduced transcriptional activity of RORγ. This reduced transcriptional activity has been attributed to the inability of the AF2 helix to complete the formation of the coactivator binding pocket necessary for coactivator proteins to bind. Electron density for the coactivator peptide EBI96 was observed for residues EFPYLLSLLG which formed a α-helix stabilized through hydrophobic interactions with the coactivator binding pocket on RORγ (Fig. 2c). This interaction is further stabilized through a conserved charged clamp wherein the backbone amide of Tyr7 and carbonyl of Leu11 of EBI96 form hydrogen bonds with Glu504 (helix12) and Lys336 (helix3) of RORγ. Formation of this charged clamp is essential for RORγ’s function for playing a role in transcriptional activation and this has been corroborated through mutagenic studies in this region. BIO592 binds in a collapsed conformation stabilizing the agonist conformation of RORγ \na Collapsed binding mode of agonist BIO592 in the hydrophobic LBS of RORγ. b Benzoxazinone ring system of agonist BIO592 packing against His479 of RORγ stabilizing agonist conformation of the AF2 helix BIO592 bound in a collapsed conformational state in the LBS of RORγ with the xylene ring positioned at the bottom of the pocket making hydrophobic interactions with Val376, Phe378, Phe388 and Phe401, with the ethyl-benzoxazinone ring making several hydrophobic interactions with Trp317, Leu324, Met358, Leu391, Ile 400 and His479 (Fig. 3a, Additional file 3). The sulfonyl group faces the entrance of the pocket, while the CF3 makes a hydrophobic contact with Ala327. Hydrophobic interaction between the ethyl group of the benzoxazinone and His479 reinforce the His479 sidechain position for making the hydrogen bond with Tyr502 thereby stabilizing the agonist conformation (Fig. 3b). RORγ AF2 helix is sensitive to proteolysis in the presence of Inverse Agonist BIO399 Next, we attempted co-crystallization with the inverse agonist BIO399. However, extensive crystallization efforts with BIO399 and RORγ518 or other AF2 intact constructs did not produce crystals. We hypothesized that the RORγ518 coactivator peptide interaction in the FRET assay was disrupted upon BIO399 binding and that a conformational rearrangement of the AF2 helix could have occurred, hindering crystallization. Specific proteolytic positions on RORγ518 when treated with Actinase E alone (Green) or in the presence of BIO399 (Red) and shared proteolytic sites (Yellow) The unfolding of the AF2 helix has been observed for other nuclear hormone receptors when bound to an inverse agonist or antagonist. We used partial proteolysis in combination with mass spectrometry to determine if BIO399 was causing the AF2 helix to unfold. Results of the Actinase E proteolysis experiments on RORγ518, the ternary complex of RORγ518 with agonist BIO592 and coactivator EBI96, or in the presence of inverse agonist BIO399 supported our hypothesis. Analysis of the fragmentation pattern showed minimal proteolytic removal of the AF2 helix by Actinase E on RORγ518 alone (ending at 504 to 506) and the ternary complex remained primarily intact (ending at 515/518) (Additional file 4). However, in the presence of inverse agonist BIO399, the proteolytic pattern showed significantly less protection, albeit the products were more heterogeneous (majority ending at 494/495), indicating the destabilization of the AF2 helix compared to either the APO or ternary agonist complex (Fig. 4, Additional file 5). Several rounds of cocrystallization attempts with RORγ518 or other RORγ AF2 helix containing constructs complexed with BIO399 had not produced crystals. We attributed the inability to form crystals to the unfolding of the AF2 helix induced by BIO399. We reasoned that if we could remove the unfolded AF2 helix using proteolysis we could produce a binary complex more amenable to crystallization. AF2 truncated RORγ BIO399 complex is more amenable to crystallization \na The binary structure of AF2-truncated RORγ and BIO399. b The superposition of inverse agonist BIO399 (Cyan) and agonist BIO592 (Green). c Movement of Met358 and His479 in the BIO399 (Cyan) and BIO592 (Green) structures The Actinase E treated RORγ518 BIO399 ternary complex (aeRORγ493/4) co-crystallized readily in several PEG based conditions. The structure of aeRORγ493/4 BIO399 complex was solved to 2.3 Å and adopted a similar core fold to the BIO592 agonist crystal structure (Fig. 5a, Additional file 3). The aeRORγ493/4 BIO399 structure diverged at the c-terminal end of Helix 11 from the RORγ518 BIO592 EBI96 structure, where helix 11 unwinds into a random coil after residue L475. Inverse agonist BIO399 uses Met358 as a trigger for inverse agonism BIO399 binds to the ligand binding site of RORγ adopting a collapsed conformation as seen with BIO592 where the two compounds superimpose with an RMSD of 0.72 Å (Fig. 5b). The majority of the side chains within 4 Å of BIO399 and BIO592 adopt similar rotomer conformations with the exceptions of Met358 and His479 (Fig. 5c). The difference density map showed clear positive density for Met358 in an alternate rotomer conformation compared to the one observed in the molecular replacement model or the other agonist containing models (Additional file 6). We tried to refine Met358 in the same conformation as the molecular replacement model or the other agonist containing models, but the results clearly indicated that this was not possible, thus confirming the new rotamer conformation for the Met358 sidechain in the inverse agonist bound structure. The change in rotomer conformation of Met358 between the agonist and inverse agonist structures is attributed to the gem-dimethyl group on the larger 7 membered benzoxazinone ring system of BIO399. The comparison of the two structures shows that the agonist conformation observed in the BIO592 structure would be perturbed by BIO399 pushing Met358 into Phe506 of the AF2 helix indicating that Met358 is a trigger for inducing inverse agonism in RORγ (Fig. 5c). BIO399 and Inverse agonist T0901317 bind in a collapsed conformation distinct from other RORγ Inverse Agonists Cocrystal structures \na Overlay of RORγ structures bound to BIO596 (Green), BIO399 (Cyan) and T0901317 (Pink). b Overlay of M358 in RORγ structure BIO596 (Green), BIO399 (Cyan), Digoxin (Yellow), Compound 2 (Grey), Compound 48 (Salmon) and Compound 4j (Orange) The co-crystal structure of RORγ with T0901317 (PDB code: 4NB6), an inverse agonist of RORγ (IC50 of 54nM in an SRC1 displacement FRET assay and an IC50 of 59nM in our FRET assay (Additional file 7)) shows that it adopts a collapsed conformation similar to the structure of BIO399 described here. The two compounds superimpose with an RMSD of 0.81 Å (Fig. 6a). The CF3 group on the hexafluoropropanol group of T0901317 was reported to fit the electron density in two conformations one of which pushes Met358 into the vicinity of Phe506 in the RORγ BIO592 agonist structure. We hypothesize that since the Met358 sidechain conformation in the T0901317 RORγ structure is not in the BIO399 conformation, this difference could account for the 10-fold reduction in the inverse agonism for T0901317 compared to BIO399 in the FRET assay. Co-crystal structures of RORγ have been generated with several potent inverse agonists adopting a linear conformation distinct from the collapsed conformations seen for BIO399 and T090131718. The inverse agonist activity for these compounds has been attributed to orientating Trp317 to clash with Tyr502 or a direct inverse agonist hydrogen bonding event with His479, both of which would perturb the agonist conformation of RORγ. BIO399 neither orients the sidechain of Trp317 toward Tyr502 nor forms a hydrogen bond with His479 suggesting its mode of action is distinct from linear inverse agonists (Additional file 8). In the linear inverse agonist crystal structures the side chain of Met358 resides in a similar position as the rotomer observed in RORγ agonist structures with BIO592 described here or as observed in the hydroxycholesterol derivatives and therefore would not trigger inverse agonism with these ligands (Fig. 6b). BIO399 shows selectivity for RORγ over RORα and RORβ in a GAL4 Cellular Reporter Assay GAL4 cell assay selectivity profile for BIO399 toward RORα and RORβ in GAL4 ROR\tγ\tα\tβ\t \tIC50 (uM)\t0.043 (+/− 0.01uM; N = 6)\t\u003e10 (N = 2)\t\u003e1.2 (N = 2)\t \tSelectivity (X)\t-\t\u003e235\t\u003e28.2\t \t \na Overlay of RORα (yellow), β (pink) and γ (cyan) showing side chain differences at Met358 inverse agonism trigger position and (b) around the benzoxazinone ring system of BIO399 In order to assess the in vivo selectivity profile of BIO399 a cellular reporter assay was implemented where the ligand binding domains of ROR α, β and γ were fused to the DNA binding domain of the transcriptional factor GAL4. The ROR-GAL4 fusion proteins were expressed in cells with the luciferase reporter gene under the control of a GAL4 promoter. BIO399 inhibited the luciferase activity when added to the cells expressing the RORγ-GAL4 fusion with an in vivo IC50 of 42.5nM while showing \u003e235 and 28 fold selectivity over cells expressing GAL4 fused to the LBD of ROR α or β, respectively (Table 1). The LBS of RORs share a high degree of similarity. However, the inverse agonism trigger of BIO399, residue Met358, is a leucine in both RORα and β. This selectivity profile for BIO399 is attributed to the shorter leucine side chain in RORα and β which would not reach the phenylalanine on the AF2 helix further underscoring the role of Met358 as a trigger for RORγ specific inverse agonism (Fig. 7a). Furthermore, RORα contains two phenylalanine residues in its LBS whereas RORβ and γ have a leucine in the same position (Fig. 6b). We hypothesize that the two phenylalanine residues in the LBS of RORα occlude the dihydrobenzoxazepinone ring system of BIO399 from binding it and responsible for the increase in selectivity for RORα over β. Conclusions We have identified a novel series of synthetic benzoxazinone ligands which modulate the transcriptional activity of RORγ in a FRET based assay. Using partial proteolysis we show a conformational change which destabilizes the AF2 helix of RORγ when the inverse agonist BIO399 binds. The two RORγ co-crystal structures reported here show how a small change to the core ring system can modulate the mode of action from agonist (BIO592) to inverse agonism (BIO399). Finally, we are reporting a newly identified trigger for achieving RORγ specific inverse agonism in an in vivo setting through Met358 which perturbs the agonist conformation of the AF2 helix and prevents coactivator protein binding. Abbreviations AF2, activation function 2; BisTRIS, 2-[Bis(2-hydroxyethyl)amino]-a-(hydroxymethyl)propane-1,3-diol; DND, DNA binding domain; DTT, 1,4-Dithiothreitol; EDTA, 2-({2-[Bis(carboxymethyl)amino]ethyl}(carboxymethyl)amino)acetic acid; FRET, fluorescence resonance energy transfer; GST, Glutathione-S-Transferase; HEPES, 2-[4(2-hydroxyethyl)-1-piperazineethanesulfonic acid; IC50, half maximal inhibitory concentration; IL-17, Interleukin-17; IPTG, isopropyl β-D-1-thiogalactopyranoside; LBD, Ligand Binding Domain; LBS, ligand binding site; LC-MS, liquid chromatography/mass spectrometry; PDB, Protein Data Bank; ROR, retinoid orphan receptor; SRC-1, steroid receptor coactivator-1; TH17 Cells, T helper cells; TRIS, 2-amino-2-hydroxymethyl-propane-1,3,diol. Additional files Competing interests The authors declare that they have no competing interests. Consent to publish Not applicable. Ethics Not applicable. References Jetten AM. 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