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+MIT License
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+Copyright (c) 2023, Melanie Vollmar
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+Permission is hereby granted, free of charge, to any person obtaining a copy of this software and associated documentation files (the “Software”), to deal in the Software without restriction, including without limitation the rights to use, copy, modify, merge, publish, distribute, sublicense, and/or sell copies of the Software, and to permit persons to whom the Software is furnished to do so, subject to the following conditions:
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+[{"sourceid":"4746701","sourcedb":"","project":"","target":"","text":"Crystal structure of SEL1L: Insight into the roles of SLR motifs in ERAD pathway Terminally misfolded proteins are selectively recognized and cleared by the endoplasmic reticulum-associated degradation (ERAD) pathway. SEL1L, a component of the ERAD machinery, plays an important role in selecting and transporting ERAD substrates for degradation. We have determined the crystal structure of the mouse SEL1L central domain comprising five Sel1-Like Repeats (SLR motifs 5 to 9; hereafter called SEL1Lcent). Strikingly, SEL1Lcent forms a homodimer with two-fold symmetry in a head-to-tail manner. Particularly, the SLR motif 9 plays an important role in dimer formation by adopting a domain-swapped structure and providing an extensive dimeric interface. We identified that the full-length SEL1L forms a self-oligomer through the SEL1Lcent domain in mammalian cells. Furthermore, we discovered that the SLR-C, comprising SLR motifs 10 and 11, of SEL1L directly interacts with the N-terminus luminal loops of HRD1. Therefore, we propose that certain SLR motifs of SEL1L play a unique role in membrane bound ERAD machinery. Protein quality control in the endoplasmic reticulum (ER) is essential for maintenance of cellular homeostasis in eukaryotes and is implicated in many severe diseases. Terminally misfolded proteins in the lumen or membrane of the ER are retrotranslocated into the cytosol, polyubiquitinated, and degraded by the proteasome. The process is called ER-associated protein degradation (ERAD) and is conserved in all eukaryotes. Accumulating studies have identified key components for ERAD, including HRD1, SEL1L (Hrd3p), Derlin-1, -2, -3 (Der1p), HERP-1, -2 (Usa1p), OS9 (Yos9), XTP-B, and Grp94, all of which are involved in the recognition and translocation of the ERAD substrates in yeast and metazoans. The components are differentially localized from the lumen and membrane of the ER to the cytosol, and have different functions in the ERAD process. Yeast ERAD components, which have been extensively characterized through genetic and biochemical studies, are comparable with mammalian ERAD components, sharing similar molecular functions and structural composition. The HRD1 E3 ubiquitin ligase, which is embedded in the ER membrane, is involved in translocating ERAD substrates across the ER membrane and catalyzing substrate ubiquitination via its cytosolic RING finger domain. SEL1L, the mammalian homolog of Hrd3p, associates with HRD1, mediates HRD1 interactions with the ER luminal lectin OS9, and recognizes substrates to be degraded. In particular, SEL1L is crucial for translocation of Class I major histocompatibility complex (MHC) heavy chains (HCs). Recent research based on the inducible Sel1l knockout mouse model highlights the physiological functions of SEL1L. SEL1L is required for ER homeostasis, which is essential for protein translation, pancreatic function, and cellular and organismal survival. However, despite the functional importance of SEL1L, the molecular structure of SEL1L has not been solved. Previous biochemical studies reveal that SEL1L is a type I transmembrane protein and has a large luminal domain comprising sets of repeated Sel1-like (SLR) motifs. The SLR motif is a structural motif that closely resembles the tetratricopeptide-repeat (TPR) motif, which is a protein-protein interaction module. Although there is evidence that the luminal domain of SEL1L is involved in substrate recognition or in forming complexes with chaperones, it is not known how the unique structure of the repeated SLR motifs contributes to the molecular function of the HRD1-SEL1L E3 ligase complex and affects ERAD at the molecular level. Furthermore, while repeated SLR motifs are commonly found in tandem arrays, the SLR motifs in SEL1L are, according to the primary structure prediction of SEL1L, interspersed among other sequences in the luminal domain and form three SLR domain clusters. Therefore, the way in which these unique structural features of SEL1L are related to its critical function in ERAD remains to be elucidated. To clearly understand the biochemical role of the SLR domains of SEL1L in ERAD, we determined the crystal structure of the central SLR domain of SEL1L. We found that the central domain of SEL1L, comprising SLR motifs 5 through 9 (SEL1Lcent), forms a tight dimer with two-fold symmetry due to domain swapping of the SLR motif 9. We also found that SLR-C, consisting of SLR motifs 10 and 11, directly interacts with the N-terminus luminal loop of HRD1. Based on these observations, we propose a model wherein the SLR domains of SEL1L contribute to the formation of stable oligomers of the ERAD translocation machinery, which is indispensable for ERAD. Results Structure Determination of SEL1Lcent The Mus musculus SEL1L protein contains 790 amino acids and has 17% sequence identity to its yeast homolog, Hrd3p. Mouse SEL1L contains a fibronectin type II domain at the N-terminus, followed by 11 SLR motifs and a single transmembrane domain at the C-terminus (Fig. 1A). The 11 SLR motifs are located in the ER lumen and account for more than two thirds of the mass of full-length SEL1L. The SLR motifs can be grouped into three regions due to the presence of linker sequences among the groups of SLR motifs: SLR-N (SLR motifs 1 to 4), SLR-M (SLR motifs 5 to 9), and SLR-C (SLR motifs 10 to 11) (Fig. 1A). Sequence alignment of the SLR motifs revealed that there is a short linker sequence (residues 336–345) between SLR-N and SLR-M and a long linker sequence (residues 528–635) between SLR-M and SLR-C (Fig. 1A). We first tried to prepare the full-length mouse SEL1L protein, excluding the transmembrane domain at the C-terminus (residues 735–755), by expression in bacteria. However, the full-length SEL1L protein aggregated in solution and produced no soluble protein. To identify a soluble form of SEL1L, we generated serial truncation constructs of SEL1L based on the predicted SLR motifs and highly conserved regions across several different species. Both SLR-N (residues 194–343) and SLR-C (residues 639–719) alone could be solubilized with an MBP tag at the N-terminus, but appeared to be polydisperse when analyzed by size-exclusion chromatography. However, the central region of SEL1L, comprising residues 337–554, was soluble and homogenous in size, as determined by size-exclusion chromatography. To define compact domain boundaries for the central region of SEL1L, we digested the protein with trypsin and analyzed the proteolysis products by SDS-PAGE and N-terminal sequencing. The results of this preliminary biochemical analysis suggested that SEL1L residues 348–533 (SEL1Lcent) would be amenable to structural analysis (Fig. 1A). Crystals of SEL1Lcent grew in space group P21 with four copies of SEL1Lcent (a total of 82 kDa) in the asymmetric unit. The structure was determined by the single-wavelength anomalous diffraction (SAD) method using selenium as the anomalous scatterer (Table 1 and Methods). The assignment of residues during model building was aided by the selenium atom positions, and the structure was refined with native data to 2.6 Å resolution with Rwork/Rfree values of 20.7/27.7%. Statistics for data collection and refinement are presented in Table 1. Overall Structure of SEL1Lcent The mouse SEL1Lcent crystallized as a homodimer, and there were two homodimers in the crystal asymmetric unit (Fig. 1B,C, Supplementary Fig. 1). The two SEL1Lcent molecules dimerize in a head-to-tail manner through a two-fold symmetry interface resulting in a cosmos-like shaped structure (Fig. 1B). The resulting structure resembles the yin-yang symbol with overall dimensions of 60 × 60 × 25 Å, where a SEL1Lcent monomer corresponds to half the symbol. The dimer formation buries a surface area of 1670 Å2 for each monomer, and no significant differences between the protomers were displayed (final root mean square deviation (RMSD) of 0.6 Å for all Cα atoms). Each protomer is composed of ten α-helices, which form the five SLRs, resulting in an elongated curved structure, confirming the primary structure prediction (Fig. 1D). The α-helices subdivide the structure into five pairs (A and B) as shown in a number of TPRs and SLRs. Helices A and B are 14 and 13 residues long, respectively, and the two helices are connected by a short turn and loop (Fig. 1D). In addition, a longer loop, consisting of approximately eight amino acids, is inserted between helix B of one SLR and helix A of the next SLR. This arrangement is a unique feature for SLRs among the major classes of repeats containing an α-solenoid. Starting from its N-terminus, the α-solenoid of SEL1L extends across a semi-circle in a right-handed superhelix fashion along the rotation axis of the yin-yang circle. However, the last helix, 9B, at the C-terminus adopts a different conformation, lying parallel to the long axis of helix 9A instead of forming an antiparallel SLR. This unique conformation of helix 9B most likely contributes to formation of the dimer structure of SEL1Lcent, as detailed below. With the exception of the last SLR, the four α-helix pairs possess similar conformations, with RMSD values of 0.7 Å for all Cα atoms. Although the sequence similarity for the pairwise alignments varies between 25% and 35%, all the residues present in the SLR motifs are conserved among the five pairs. The SLR domain of SLR-M ends at residue 524, and C-terminal amino acids 525–533 of the protein are not visible in the electron density map, suggesting that this region is highly flexible. Since no information regarding dimer formation by SEL1L through its SLR motifs is available, we tested whether the SEL1Lcent dimer shown in our crystal structure is a biological unit. First, we cross-linked SEL1Lcent or a longer construct of SEL1L (SEL1Llong, residues 337–554) using various concentrations of glutaraldehyde (GA) or dimethyl suberimidate (DMS) and analyzed the products by SDS-PAGE. We detected bands at the mass of a dimer for both SEL1Lcent and SEL1Llong when cross-linked with low concentrations of GA (0.005%) or DMS (0.3 mM) (Supplementary Fig. 2A,B). Next, we conducted analytical ultracentrifugation of SEL1Lcent. Consistent with the cross-linking data, analytical ultracentrifugation revealed that the molecular weight of SEL1Lcent corresponds to a dimer (Supplementary Fig. 2C). Taken together, these data indicate that some kind of dimer is formed in solution. Dimer Interface of SEL1Lcent In contrast to a previously described SLR motif containing proteins that exist as monomers in solution, SEL1Lcent forms an intimate two-fold homotypic dimer interface (Figs 1B and 2A). The concave surface of each SEL1L domain comprising helix 5A to 9A encircles its dimer counterpart in an interlocking clasp-like arrangement. However, no interactions were seen between the two-fold-related protomers through the concave inner surfaces themselves. Rather, the unique structure of SLR motif 9, consisting of two parallel helices (9A and 9B), is located in the space generated by the concave surface and provides an extensive dimerization interface between the two-fold-related molecules (Fig. 2A). Helix 9B from one protomer inserts into the empty space surrounded by the concave region in the other monomer, forming a domain-swapped conformation. Three major contact interfaces are involved in the interactions, and all interfaces are symmetrically related between the dimer subunits (Fig. 2A). Structure-based sequence alignment of 135 SEL1L phylogenetic sequences using a ConSurf server revealed that the surface residues in the dimer interfaces were highly conserved among the SEL1L orthologs (Fig. 1E). First, helix 9B of each SEL1Lcent subunit interacts with residues lining the inner groove from the SLR α-helices (5B, 6B, 7B, and 8B) from its counterpart. In this interface, Leu 516 and Tyr 519 on helix 9B are located in the center, making hydrophobic interactions with Trp 478 on helix 8B, Val 444 on helix 7B, Phe 411 on helix 6B, and Leu 380 on helix 5B from the SEL1Lcent counterpart (Fig. 2A, Interface 1 detail). In addition to hydrophobic interactions, the side chain hydroxyl group of Tyr 519 and the main-chain oxygen of Ile 515 form H-bonds to the side chain of the conserved Gln 377 and His 381 on helix 5B of the two-fold-related protomer. The side chain of Gln 523 forms an H-bond to the side chain of Asp 480 on the two-fold-related protomer (Fig. 2A, Interface 1 detail). Second, the residues from helix 9A interact with the residues from helix 5A of its counterpart in a head-to-tail orientation. In this interface, the interacting residues on helix 9A, including Leu 503, Tyr 499, and the aliphatic side chain of Lys 500, form an extensive network of van der Waals contacts with the hydrophobic residues of the counterpart helix 5A, including Tyr 360, Leu 356, Tyr 359, and Leu 363. In addition to hydrophobic interactions, the side chains of Asn 507 and Ser 510 on helix 9A make H-bonds with highly conserved Arg 384 in the loop between helix 5B and 6A from the two-fold-related protomer (Fig. 2A, Interface 2 detail). Third, the helix 9B from each protomer is involved in the dimer interaction by forming a two-fold antiparallel symmetry. In particular, the side chains of hydrophobic residues, including Phe 518, Leu 521, and Met 524, are directed toward each other, where they make both inter- and intramolecular contacts (Fig. 2A, Interface 3 detail). To further investigate the interactions observed in our crystal structure, we generated a C-terminal deletion mutant (SEL1L348–497) lacking SLR motif 9 (helix 9A and 9B) from SEL1Lcent for comparative analysis. The deletion mutant and the wild-type SEL1Lcent showed no difference in spectra by CD spectroscopy, indicating that the deletion of the SLR motif 9 did not affect the secondary structure of SEL1Lcent (Supplementary Fig. 3). However, the mutant behaved as a monomer in size-exclusion chromatography and analytical ultracentrifugation experiments (Fig. 2B, Supplementary Fig. 2C). Additionally, to further validate the key residues involved in dimer formation, we generated a triple point mutant (Interface 1, I515A, L516A, and Y519A) of the hydrophobic residues that are involved in dimerization. The triple point mutant eluted at the monomer position upon size-exclusion chromatography, while the negative control point mutant (Q460A) eluted at the same position as the wild-type. Notably, a single-residue mutation (L521A in interface 3) abolished the dimerization of SEL1Lcent (Fig. 2B). Leu 521 is located in the dimerization center of the antiparallel 9B helices in the SEL1Lcent dimer. Taken together, these structural and biochemical data demonstrate that SEL1Lcent exists as a dimer in solution and that SLR motif 9 in SEL1Lcent plays an important role in generating a two-fold dimerization interface. The Two Glycine Residues (G512 and G513) Create a Hinge for Domain Swapping of SLR Motif 9 SLRs of mouse SEL1L were predicted using the TPRpred server. Based on the prediction, full-length SEL1L contains a total of 11 SLR motifs, and our construct corresponds to SLR motifs 5 through 9. Although amino acid sequences from helix 9A and 9B correctly aligned with the regular SLR repeats and corresponded to SLR motif 9 (Fig. 3A), the structural arrangement of the two helices deviated from the common structure for the SLR motif. According to our crystal structure, the central axis of helix 9B is almost parallel to that of helix 9A (Fig. 3B). However, this unusual conformation of SLR motif 9 seems to be essential for dimer formation, as described earlier. For this structural geometry, two adjacent residues, Gly 512 and Gly 513, in SEL1L confer flexibility at this position by adopting main-chain dihedral angles that are disallowed for non-glycine residues. The phi and psi dihedrals are 100° and 20° for Gly 512, and 110° and −20° for Gly 513, respectively (Fig. 3C). Gly 513 is conserved among other SLR motifs in the SEL1Lcent, but Gly 512 is present only in the SLR motif 9 of SEL1Lcent (Fig. 3A). Thus, the Gly-Gly residues generate an unusual sharp bend at the C-terminal SLR motif 9. The involvement of a glycine residue in forming a hinge for domain swapping has been reported previously. The significance of Gly 513 is further highlighted by its absolute conservation among different species, including the budding yeast homolog Hrd3p. To further investigate the importance of Gly 512 and Gly 513 in the unusual SLR motif geometry, we generated a point mutation (Gly to Ala), which restricts the flexibility. Although the Gly 512 and Gly 513 residues are closely surrounded by helix 9B from the counter protomer, there is enough space for the side chain of alanine, suggesting that no steric hindrance would be caused by the mutation (Fig. 3C). This means that the effect of the mutation is mainly to generate a more restricted geometry at the hinge region. G512A or G513A alone showed no differences from wild-type in terms of the size-exclusion chromatography elution profile (Fig. 3D), suggesting that the restriction for single glycine flexibility would not be enough to break the swapped structure of helix 9B. However, the double mutant (G512A/G513A) eluted over a broad range and much earlier than the wild-type, suggesting that mutation of the residues involved in the hinge linking helix 9A and 9B significantly affected the geometry of helix 9B in generating domain swapping, and eventually altered the overall oligomeric state of SEL1Lcent into a polydisperse pattern (Fig. 3D, Supplementary Fig. 6). When the residues were mutated to lysine (G512K/G513K), the mutant not only restricted the geometry of residues at the hinge but also generated steric hindrance during interaction with the counter protomer of SEL1Lcent, thereby inhibiting self-association of SEL1Lcent completely. The G512K/G513K double mutant eluted at the monomer position in size-exclusion chromatography (Fig. 3D). A previous study shows that induction of steric hindrance by mutation destabilizes the dimerization interface of a different protein, ClC transporter. Collectively, these data suggest that the Gly 512 and Gly 513 at the connection between helix 9A and 9B play a crucial role in forming the domain-swapped conformation that enables dimer formation. SEL1L Forms Self-oligomers through SEL1Lcent domain in vivo Next, we examined if SEL1L also forms self-oligomers in vivo using HEK293T cells. We generated full-length SEL1L-HA and SEL1L-FLAG fusion constructs and co-transfected the constructs into HEK293T cells. A co-immunoprecipitation assay using an anti-FLAG antibody followed by Western blot analysis using an anti-HA antibody showed that full-length SEL1L forms self-oligomers in vivo (Fig. 4A). To further examine whether the SEL1Lcent domain is sufficient to physically interact with full-length SEL1L, we generated SEL1Lcent and SLR motif 9 deletion (SEL1L348–497) construct, which were fused to the C-terminus of SEL1L signal peptides. Co-immunoprecipitation analysis showed that the SEL1Lcent was sufficient to physically interact with the full-length SEL1L, while SEL1L348–497 failed to do so (Fig. 4A). Interestingly, however, the expression level of SEL1L348–497 was consistently lower than that of SEL1Lcent (Fig. 4A,B). Semi-quantitative RT-PCR revealed no significant difference in transcriptional levels of the two constructs (data not shown). We speculated that SEL1L348–497 could be secreted while the SEL1Lcent is retained in the ER by association with the endogenous ERAD complex. Indeed, immunoprecipitation followed by western blot analysis using the culture medium detected secreted SEL1L348–497 fragment, but not SEL1Lcent (Fig. 4B). We next examined if the reason why SEL1L348–497 failed to bind to the full-length SEL1L may be because of the lower level of SEL1L348–497 in the ER lumen compared to SEL1Lcent fragment. In order to retain two SEL1L fragments in the ER lumen, we added KDEL ER retention sequence to the C-terminus of both fragments. Indeed, the addition of KDEL peptide increased the level of SEL1L348–497 in the ER lumen (Fig. 4D,E) and the immunostaining analysis showed both constructs were well localized to the ER (Fig. 4C). We further analyzed whether SEL1Lcent may competitively inhibit the self-oligomerization of SEL1L in vivo. To this end, we co-transfected the differentially tagged full-length SEL1L (SEL1L-HA and SEL1L-FLAG) and increasing doses of SEL1Lcent-KDEL, SEL1L348–497-KDEL or SEL1Lcent (L521A)-KDEL, respectively. Co-immunoprecipitation assay revealed that wild-type SEL1Lcent-KDEL, indeed, competitively disrupted the self-association of the full-length SEL1L (Fig. 4E). In contrast, SEL1L348–497-KDEL and the single-residue mutation L521A in SEL1Lcent did not competitively inhibit the self-association of full-length SEL1L (Fig. 4E,F). These data suggest that the SEL1L forms self-oligomers and the oligomerization is mediated by the SEL1Lcent domain in vivo. Structural Comparison of SEL1L SLRs with TPRs or SLRs of Other Proteins Previous studies reveal that TPRs and SLRs have similar consensus sequences, suggesting that their three-dimensional structures are also similar. The superposition of isolated TPRs from Cdc23 (S. pombe, cell division cycle 23 homolog, PDB code 3ZN3) and SLRs from HcpC (Helicobacter Cysteine-rich Protein C, PDB code 1OUV) yields RMSDs below 1 Å, confirming that the isolated repeats are indeed similar. This is relevant to SLR motifs in SEL1L, as isolated SLR motifs from SEL1Lcent showed good structural alignment with isolated TPRs (RMSD 1.6 Å for all Cα chains) from Cdc23N-term and SLRs (RMSD 0.6 Å for all Cα chains) from HcpC (Fig. 5A). However, superimposing the structure of SLR motifs 5 to 9 from SEL1Lcent onto the overall Cdc23N-term or full-length HcpC structures revealed that SLR motifs 5 to 9 in SEL1Lcent have a different superhelical structure than either Cdc23 or HcpC (RMSD values of \u003e2.5 Å for Cα atoms) (Fig. 5B). The differences may result from the differing numbers of residues in the loops and differences in antiparallel helix packing. Moreover, there are conserved disulfide bonds in the SLR motifs of HcpC and HcpB, but no such bonds are observed in SEL1Lcent. These factors contribute to the differences in the overall conformation of the SLR motifs in SEL1L and other SLR or TPR motif-containing proteins. Another major difference in the structure of SLR motifs between SEL1L and HcpC is the oligomeric state of proteins. The TPR motif is involved in the dimerization of proteins such as Cdc23, Cdc16, and Cdc27. In particular, the N-terminal domain of Cdc23 (Cdc23N-term) has a TPR-motif organization similar to that of the SLR motif in SEL1Lcent. The seven TPR motifs of Cdc23N-term are assembled into a superhelical structure, generating a hollow surface and encircling its dimer counterpart in an interlocking clasp-like arrangement (Fig. 5C). The TPR motif 1 (TPR1) of each Cdc23N-term subunit is located in the hollow surface of the counter subunit and interacts with residues lining the inner groove TPR α-helices, generating two-fold symmetry homotype interactions. However, in this structure, a conformational change in the TPR motif itself is not observed. Self-association of HcpC has not been reported, and there is no domain-swapped structure in the SLR motifs of HcpC, in contrast to that observed in SEL1Lcent. Although SEL1L contains a number of SLR motifs comparable to HcpC, the SLR motifs in SEL1L are interrupted by other sequences, making three SLR motif clusters (Fig. 1A). The interrupted SLR motifs may be required for dimerization of SEL1Lcent, as five SLR motifs are more than enough to form the semicircle of the yin-yang symbol (Fig. 1B). Helix 5A from SLR motif 5 meets helix 9A from SLR motif 9 of the counterpart SEL1L. If the SLR motifs 5 to 9 were not isolated from other SLR motifs, steric hindrance could interfere with dimerization of SEL1L. This is one of the biggest differences from TPRs in Cdc23 and from the SLRs in HcpC, where the motifs exist in tandem. TPR and SLR motifs are generally involved in protein-protein interaction modules, and the sequences between the SLR motifs of SEL1L might actually facilitate the self-association of this protein. SLR-C of SEL1L Binds HRD1 N-terminus Luminal Loop Based on the structural data presented herein, a possible arrangement of membrane-associated ERAD components in mammals, highlighting the molecular functions of SLR domains in SEL1L, is shown in Fig. 6C. We suggest that the middle SLR domains are involved in the dimerization of SEL1L based on the crystal structure and biochemical data. SLR-C, which contains SLR motifs 10 to 11, might be involved in the interaction with HRD1. Indirect evidence from a previous yeast study shows that the circumscribed region of C-terminal Hrd3p, specifically residues 664–695, forms contacts with the Hrd1 luminal loops. The Hrd3p residues 664–695 correspond to mouse SEL1L residues 696–727, which include the entire helix 11B (residue 697–709) of SLR motif 11 and a well-conserved adjacent region (Supplementary Fig. 4). This observation is supported by the following: (1) the meticulous range of SLR motif 10 to 11 is newly established from a structure-guided SLR motif alignment, based on the present structure study, and (2) the relatively high sequence conservation between mammalian SEL1L and yeast Hrd3p around SLR motifs 10 to 11, which contain contact regions with HRD1 (Hrd1p) (Supplementary Figs. 4 and 5). To address this hypothesis, we prepared constructs encoding mouse HRD1 luminal fragments fused to GST as shown in Fig. 6A, and tested their ability to bind certain SLR motifs in SEL1L. The fusion proteins were immobilized on glutathione-Sepharose beads and probed for binding to SLR-N, SLR-M, SLR-C, and monomer form of SLR-M (SLR-ML521A). Figure 6B shows that the SLR-C, consisting of SLR motifs 10 and 11, exclusively interacts with N-terminal luminal loop (residues 21–42) of HRD1. The molecular functions of SLR-N are unclear. One possibility is that SLR-N contributes to substrate recognition of proteins to be degraded because there are a couple of putative glycosylation sites within the SLR-N domain (Fig. 1A). SEL1Lcent contains a putative N-glycosylation site, Asn 427, which is highly conserved among different species and structurally exposed to the surface of the SEL1L dimer according to the crystal structure (Fig. 6C). Discussion Many reports demonstrate that membrane-bound ERAD machinery proteins in yeast, such as Hrd1p, Der1p, and Usa1p, are involved in oligomerization of ERAD components. The Hrd1p complex forms dimers upon sucrose gradient sedimentation and size-exclusion chromatography. Previous data show that HA-epitope-tagged Hrd3p or Hrd1p efficiently co-precipitate with unmodified Hrd3p and Hrd1p, respectively, suggesting that both Hrd1p and Hrd3p homodimers are involved in self-association of the Hrd complex. Considering that the functional and structural composition of ERAD components are conserved in both yeast and mammals, we propose that the mammalian ERAD components also form self-associating oligomers. This hypothesis is supported by cross-linking data suggesting that human HRD1 forms a homodimer. Consistent with the previous data, our crystal structure and biochemical data demonstrate that mouse SEL1Lcent exists as a homodimer in the ER lumen via domain swapping of SLR motif 9. We need to further test whether there are contacts involved in dimer formation in SEL1L in addition to those in the SLR-M region. In yeast, Usa1p acts as a scaffold for Hrd1p and Der1p, in which the N-terminus of Usa1p interacts with the C-terminal 34 amino acids of Hrd1p in the cytosol to induce oligomerization of Hrd1p, which is essential for its activity. However, metazoans lack a clear Usa1p homolog. Although mammalian HERP has sequences and domains that are conserved in Usa1p, the molecular function of HERP is not clearly related to that of Usa1p. Rather, recent research shows that a transiently expressed HRD1-SEL1L complex alone associates with the ERAD lectins OS9 or XTP-B and is sufficient to facilitate the retrotranslocation and degradation of the model ERAD substrate α-antitrypsin null Hong-Kong (NHK) and its variant, NHK-QQQ, which lacks the N-glycosylation sites. Assuming that the correct oligomerization of ERAD components may be critical for their function, we hypothesize that homodimer formation of SEL1L in the ER lumen may stabilize oligomerization of the HRD complex, given that SEL1L forms a stoichiometric complex with HRD1. This is further supported by our data showing that the SLR-C of SEL1L directly interacts with the luminal fragment of HRD1 in the ER lumen. Although the organization of membrane-bound HRD complex components may be very similar between metazoans and yeast, the molecular details of interactions between the components may not necessarily be conserved. In yeast, it is unclear whether self-association of Hrd3p is due to SLR motifs because the sequence of Hrd3p does not align precisely with the SLR motifs in SEL1L. Furthermore, we are uncertain whether self-association of Hrd3p contributes to formation of the active form of the Hrd1p complex. Recently, a truncated version of Yos9 was shown to form a dimer in the ER lumen and to contribute to the dimeric state of the Hrd1p complex. This interaction seems to be weak because direct Yos9-Yos9 interactions were not detected in immunoprecipitation experiments from yeast cell extracts containing different epitope-tagged variants of Yos9. However, the dimerization of Yos9 could provide a higher stability for the Hrd1p complex oligomer. Likewise, the dimerization of SEL1L might provide stability for the mammalian HRD oligomer complex. Further cell biological studies are required to clarify whether SEL1L (Hrd3p) dimerization could be cooperative with the oligomerization of the HRD complex. Considering that it is very important for the function of the HRD complex that the components assemble as oligomers, we believe that the self-association of SEL1L strongly contributes to generating active forms of the HRD complex, even in the absence of Usa1p, in metazoans. These findings should provide a foundation for molecular-level studies to understand the membrane-associated HRD complex assembly in ERAD. Methods Protein Production The expression and purification of SEL1L was performed as described previously. Crystallization and SAD Structure Determination of SEL1Lcent Crystals were grown using the hanging-drop vapor diffusion method at 4 °C. For crystallization of the M. musculus SEL1Lcent, 1 μl of protein solution (in 25 mM Tris-HCl, 150 mM NaCl, and 5 mM DTT, pH 7.5) was equilibrated with 1 μl of well solution (30% isopropanol, 100 mM NaCl, 100 mM Tris, 5 mM DTT, and 20 mM phenol, pH 8.5). The crystals, which appeared after 4 days, contain two SEL1Lcent dimers in the asymmetric unit (space group P21, a = 29.13, b = 110.52, c = 109.81 Å, α = 90.00, β = 90.61, γ = 90.00, 44% solvent). For X-ray diffraction experiments, crystals were transferred to well solution plus paraffin-oil, then flash frozen in liquid nitrogen. SAD data were collected with a Se-Met crystal at beamline 7A of the Pohang Accelerator Laboratory (PAL) and processed using HKL2000 software. Native data (2.6 Å resolution) were collected from a single frozen crystal at the same beamline of PAL and were integrated and scaled as described above. The SAD data analysis was performed using Phenix software using data between 50 and 2.9 Å resolution. Phenix identified 31 of the 32 selenium sites and refined these to give a mean f.o.m. = 0.472. Electron density modification, including non-crystallographic symmetry (NCS) averaging, using the RESOLVE software yielded an initial electron density map of excellent quality. Model building and refinement were carried out with the Coot and Phenix programs, respectively. The final model was refined to an R factor of 20.7% (Rfree = 27.7%) for native data between 30 and 2.6 Å resolution (Table 1). The final model consisted of 5402 protein atoms and 47 water molecules. There were no outliers in a Ramachandran plot of the final model. The model contained four copies of SEL1Lcent (residues 348–533) in the asymmetric unit. Of these, the following residues were not modeled due to weak electron densities: SEL1Lcent residues 348–351, 420, 421, and 525–533 in the first copy; residues 348–351 and 525–533 in the second and third copies; and residues 348–352 and 525–533 in the fourth copy. The X-ray data and refinement statistics are summarized in Table 1. Cell Culture and Plasmids Construction HEK293T cells were cultured in DMEM (Gibco) supplemented with 10% FBS. The mouse Sel1L gene was cloned into pCS108 and the 3 × HA or 3 × FLAG tag was fused to the C-terminus of SEL1L. The signal peptide from Xenopus Sel1L was cloned into pCS108 and the mouse SEL1Lcent domain, SEL1L (348-497) fragments, and SEL1Lcent (L521A) were fused to the C-terminus of the signal peptide. Then, a 3 × HA or a 3 × FLAG tag was fused to the C-terminus of the constructs. For the ER retention signal, the KDEL sequences were added to the C-terminus of the fragments. The plasmids were transfected using Lipofectamine 2000 (Life Technologies) according to the manufacturer’s manual. Western Blot Analysis and Immunostaining For western blot analysis, HEK293T cells were transfected with the indicated construct and harvested after washing in PBS. The cells were homogenized in lysis buffer (50 mM Tris, pH 7.4, 150 mM NaCl, 0.1% Triton X-100, 5% glycerol), supplemented with protease and phosphatase inhibitor cocktails. Homogenates were cleared by centrifugation at 13,200 rpm for 15 minutes at 4 °C. The lysates were subsequently used for either co-immunoprecipitation experiment or western blot analysis. For the western blot analysis, the samples were run onto 6–12% polyacrylamide gel. Blots were blocked in 5% TBS + 0.05% Tween 20 and incubated with anti-DDDD-K (Abcam) or anti-HA (Roche) antibodies. Proteins were visualized using HRP-conjugated secondary antibodies (1:4000) and SuperSignal West Pico Chemiluminescent Substrate or SuperSignal West Dura Extended Duration Substrate (Thermo) and exposed to ChemiDoc MP (Bio-Rad). For immunostaining, the cells were fixed in 4% formaldehyde and incubated with the indicated antibodies. The coverslips were incubated in blocking solution (10% FBS + 2% DMSO in TBS + 0.1% Triton X-100) at room temperature for 30 minutes to block non-specific binding. Fluorescent labeling was performed using Alexa Fluor 555 or 488-conjugated secondary antibodies and nuclei were stained with DAPI. The samples were mounted and confocal images were obtained using a Zeiss LSM700. GST Pull-down Assay For pull-down experiments, 400 μg of HRD1 luminal fragment GST-fusion proteins were incubated with 5 μl of a 50% (v/v) slurry of glutathione sepharose 4B beads (GE Healthcare) for 50 min at 4 °C. Beads were washed twice with buffer A (150 mM NaCl, 25 mM sodium phosphate pH 7.5, 5 mM DTT), and then mixed with 100 μg of MBP-SEL1L protein (SLR-N, SLR-M, SLR-C, and SLR-ML521A) in buffer A, in a total assay volume of 500 μl. The assay mix was incubated at 4 °C for 15 minutes, and beads were washed twice with 500 μl buffer A. Proteins were eluted with SDS sample buffer, and analyzed by SDS-PAGE. Additional Information Accession Numbers: The coordinates and structure factors have been deposited in the Protein Data Bank with the accession code of 5B26. How to cite this article: Jeong, H. et al. Crystal structure of SEL1L: Insight into the roles of SLR motifs in ERAD pathway. Sci. Rep. 6, 20261; doi: 10.1038/srep20261 (2016). Supplementary Material Protein misfolding, functional amyloid, and human disease Cleaning up: ER-associated degradation to the rescue Defining human ERAD networks through an integrative mapping strategy Road to ruin: targeting proteins for degradation in the endoplasmic reticulum Distinct ubiquitin-ligase complexes define convergent pathways for the degradation of ER proteins OS-9 and GRP94 deliver mutant α1-antitrypsin to the Hrd1–SEL1L ubiquitin ligase complex for ERAD A luminal surveillance complex that selects misfolded glycoproteins for ER-associated degradation The Hrd1p ligase complex forms a linchpin between ER-lumenal substrate selection and Cdc48p recruitment The ubiquitylation machinery of the endoplasmic reticulum Endoplasmic reticulum degradation requires lumen to cytosol signaling transmembrane control of Hrd1p by Hrd3p Stringent requirement for HRD1, SEL1L, and OS-9/XTP3-B for disposal of ERAD-LS substrates A complex of Yos9p and the HRD ligase integrates endoplasmic reticulum quality control into the degradation machinery Multiprotein complexes that link dislocation, ubiquitination, and extraction of misfolded proteins from the endoplasmic reticulum membrane SEL1L nucleates a protein complex required for dislocation of misfolded glycoproteins SEL1L, the homologue of yeast Hrd3p, is involved in protein dislocation from the mammalian ER The ER-associated degradation adaptor protein Sel1L regulates LPL secretion and lipid metabolism Sel1L is indispensable for mammalian endoplasmic reticulum-associated degradation, endoplasmic reticulum homeostasis, and survival Sel1-like repeat proteins in signal transduction The structure of the tetratricopeptide repeats of protein phosphatase 5: implications for TPR-mediated protein-protein interactions The crystal structure of Helicobacter cysteine-rich protein C at 2.0 Å resolution: similar peptide-binding sites in TPR and SEL1-like repeat proteins The Crystal Structure of Helicobacter pyloriCysteine-rich Protein B Reveals a Novel Fold for a Penicillin-binding Protein ConSurf 2005: the projection of evolutionary conservation scores of residues on protein structures TPRpred: a tool for prediction of TPR-, PPR-and SEL1-like repeats from protein sequences Control of domain swapping in bovine odorant-binding protein Design, function and structure of a monomeric ClC transporter The four canonical TPR subunits of human APC/C form related homo-dimeric structures and stack in parallel to form a TPR suprahelix The E3 ubiquitin ligases Hrd1 and gp78 bind to and promote cholera toxin retro-translocation Herp regulates Hrd1-mediated ubiquitylation in a ubiquitin-like domain-dependent manner Der1 promotes movement of misfolded proteins through the endoplasmic reticulum membrane Usa1 functions as a scaffold of the HRD-ubiquitin ligase Role of HERP and a HERP-related Protein in HRD1-dependent Protein Degradation at the Endoplasmic Reticulum Retrotranslocation of a misfolded luminal ER protein by the ubiquitin-ligase Hrd1p SEL1L protein critically determines the stability of the HRD1-SEL1L endoplasmic reticulum-associated degradation (ERAD) complex to optimize the degradation kinetics of ERAD substrates Structural and biochemical basis of Yos9 protein dimerization and possible contribution to self-association of 3-hydroxy-3-methylglutaryl-coenzyme A reductase degradation ubiquitin-ligase complex Crystallization and preliminary X-ray diffraction analysis of the Sel1-like repeats of SEL1L Processing of X-ray diffraction data collected in oscillation mode PHENIX: a comprehensive Python-based system for macromolecular structure solution Automated MAD and MIR structure solution Features and development of Coot Author Contributions H.J. and H.L. performed cloning, expression, and protein purification. H.J., S.H. and C.L. crystallized, collected X-ray data, and solved the protein structure. H.J., H.S., E.S. and Y.J performed biochemical experiments. H.J., T.P. and C.L. wrote the manuscript. T.P. and C.L. designed and supervised the project. All the authors discussed the results, commented on the manuscript, and approved the manuscript. Crystal Structure of SEL1Lcent. (A) The diagram shows the domain structure of Mus musculus SEL1L, as defined by proteolytic mapping and sequence/structure analysis. The 11 SLR motifs were divided into three groups (SLR-N, SLR-M, and SLR-C) due to the presence of linker sequences that are not predicted SLR motifs. Putative N-glycosylation sites are indicated by black triangles. We determined the crystal structure of the SLR-M, residues 348–533. (B) Ribbon diagram of the biological unit of the SEL1Lcent, viewed along the two-fold NCS axis. The crystal structure was determined by SAD phasing using selenium as the anomalous scatterer and refined to 2.6 Å resolution (Table 1). (C) SEL1Lcent ribbon diagram rotated 90° around a horizontal axis relative to (B). (D) One protomer of the SEL1Lcent dimer. This view is rotated about 90° anticlockwise from the bottom copy in (B), along the two-fold NCS axis. Starting from the N-terminus, SEL1Lcent has five SLR motifs comprising ten α helices. Each SLR motif (from 5 to 9) is indicated in a different color. (E) Evolutionary conservation of surface residues in SEL1Lcent, calculated using ConSurf, from a structure-based alignment of 135 SEL1L sequences. The surface is colored from red (high) to white (poor) according to the degree of conservation in the SEL1L phylogenetic orthologs. The ribbon diagram of the counterpart protomer is drawn to show the orientation of the SEL1Lcent dimer. Dimer Interface of SEL1Lcent. (A) The diagram on the left shows the SEL1Lcent dimer viewed along the two-fold symmetry axis. Three distinct contact regions are indicated with labeled boxes. The close-up view on the right shows the residues of SEL1Lcent that contribute to dimer formation via the three contact interfaces. Oxygen and nitrogen atoms are shown as red and blue, respectively. The yellow dotted lines indicate intermolecular hydrogen bonds between two protomers of SEL1Lcent. (B) Size-exclusion chromatography (SEC) analysis of the wild-type and dimeric interface SEL1Lcent mutants to compare the oligomeric states of the proteins. The standard molecular masses for the SEC experiments (top) were obtained from the following proteins: aldolase, 158 kDa; cobalbumin, 75 kDa; ovalbumin, 44 kDa; and carbonic anhydrase, 29 kDa. Chromatography was performed on a Superdex 200 column with a buffer containing 25 mM Tris, 150 mM NaCl, and 5 mM DTT (pH 7.5). The elution fractions, indicated by the gray shading, were run on SDS-PAGE and are shown below the gel-filtration elution profile. The schematic diagrams representing the protein constructs used in the SEC are shown on the left of each SDS-PAGE profile. Domain Swapping for Dimerization of SEL1Lcent. (A) Sequence alignment of the SLR motifs in SEL1L. The 11 SLR motifs were aligned based on the present crystal structure of SEL1Lcent. The sequences of SEL1Lcent included in the crystal structure are highlighted by the blue box. The secondary structure elements are indicated above the sequences, with helices depicted as cylinders. Residues that are conserved in at least 7 out of 11 sequences are red. The GG sequence in SLR motif 9, which creates the hinge for domain swapping (see text), is shaded yellow. Stars below the sequences indicate the specific residues that commonly appear in SLRs. (B) Structure alignment of five SLR motifs in SEL1Lcent is shown to highlight the unusual geometry of SLR motif 9. Each SLR motif is shown in a different color. The arrow indicates the direction of the helical axes. In SLR motif 9, the axes for the two helices are almost parallel, while the other SLR motifs adopt an α-hairpin structure. (C) Stereo view shows that the Gly 512 and Gly 513 residues are surrounded by neighboring residues from helix 9B from the counterpart dimer. Oxygen and nitrogen atoms are colored red and blue, respectively. The Gly 512 and Gly 513 residues are colored green and red, respectively. (D) The following point mutations were generated to check the effect of the Gly 512 and Gly 513 residues in terms of generating the hinge of SLR motif 9: G512A, G513A, G512A/G513A, and G512K/G513K. Size-exclusion chromatography was conducted as described in Fig. 2B. The standard molecular masses are shown at the top as in Fig. 2B. SEL1L forms self-oligomer mediated by the SEL1Lcent domain in vivo. (A) HEK293T cells were transfected with the indicated plasmid constructs and the lysates were immunoprecipitated with an anti-FLAG antibody followed by western blot analysis using an anti-HA antibody. The full-length SEL1L-FLAG was co-immunoprecipitated with the full-length SEL1L-HA. Also, SEL1Lcent was co-immunoprecipitated with the full-length SEL1L while the SLR motif 9 deletion failed to do so. (B) The HEK293T cells were transfected with the indicated plasmid constructs and the cell lysate and culture media were analyzed by western blot analysis and immunoprecipitation respectively. The SEL1L348–497 fragment was secreted to the culture media but the SEL1Lcent was retained in the ER. (C) SEL1Lcent-FLAG-KDEL and SEL1L348–497-FLAG-KDEL localized to the ER. The nuclei were stained with DAPI in blue. The ER was visualized with the anti-calnexin antibody in green. The SEL1L fragments were stained in red. (D) HEK293T cells were transfected with the indicated plasmid constructs and the lysates were immunoprecipitated with an anti-HA antibody followed by Western blot analysis using an anti-FLAG antibody. The full-length SEL1L forms self-oligomers and the SEL1Lcent-FLAG-KDEL was co-immunoprecipitated with full-length SEL1L-HA. The red asterisk indicates the expected signal for SEL1L348–497-FLAG-KDEL. SEL1L348–497-FLAG-KDEL did not co-immunoprecipitate with full-length SEL1L-HA. The white asterisks indicate non-specific bands. (E) SEL1Lcent-HA-KDEL competitively inhibited self-oligomerization of full-length SEL1L. The indicated plasmid constructs were transfected and immunoprecipitation assay was performed using an anti-FLAG antibody followed by western blot analysis using an anti-HA antibody. The red rectangle indicates competitively inhibited SEL1L self-oligomer formation by the increasing doses of SEL1Lcent-HA-KDEL. (F) L521A point mutant in SEL1Lcent did not inhibit the self-association of SEL1L. Comparison of SLR in SEL1L with TPR or Other SLR-Containing Proteins. (A) Ribbon diagram showing superimposition of an isolated TPR motif from Cdc23 and an SLR motif from SEL1Lcent (left), and SLR motifs in HcpC and SEL1Lcent (right). The SEL1L, Cdc23, and HcpC are colored magenta, green and cyan, respectively. Black arrows indicate the helical axes. The red arrow indicates disulfide bonds in the HcpC, and Cys residues involved in disulfide bonding are shown by a yellow line. (B) Ribbon representation showing superimposition of Cdc23 and SEL1Lcent (left) or HcpC and SEL1Lcent (right) to compare the overall organization of the α-solenoid domain. Both SEL1Lcent schematics are identically oriented for comparison. The Cα atoms of the residues in each α-solenoid domain are superimposed with a root-mean-squared deviation of 3.3 Å for Cdc23 and SEL1Lcent (left), and 2.5 Å for HcpC and SEL1Lcent (right). SEL1Lcent, Cdc23, and HcpC are colored as in (A). (C) Ribbon diagram showing the overall structure of Cdc23N-term (left) and SEL1Lcent (right) to compare their similarities regarding dimer formation through domain swapping. The view is along the two-fold axis. The Role of SLR-C in ERAD machinery and Model for the Organization of Proteins in Membrane-Associated ERAD Components. (A) Schematic diagram shows three HRD1 fragment constructs used in the GST pull-down experiment. (B) Pull-down experiments to examine the interactions between HRD luminal loops and certain SLR motifs of SEL1L. Fragments of the luminal loop of HRD1 fused to GST were immobilized on glutathione sepharose beads and incubated with purified three clusters of SLR motifs and monomer form of SLR-M (SLR-ML521A, right panel) in SEL1L. Proteins were analyzed by 12% SDS-PAGE and Coomassie blue staining. (C) Schematic representation of the organization of metazoan ERAD components in the ER membrane. The 11 SLR motifs of SEL1L were expressed with red cylinders and grouped into three parts (SLR-N, SLR-M, and SLR-C) based on the sequence alignment across the motifs and the crystal structure presented herein. We hypothesized that the interrupted SLR motifs of SEL1L have distinct functions such that the SLR-M is important for dimer formation of the protein, and SLR-C is involved in the interaction with HRD1 in the ER lumen. The surface representation of SEL1Lcent is placed in the same orientation as that shown in the schematic model to show that the putative N-glycosylation site, residue N427 (indicated in yellow), is exposed on the surface of the protein. The yellow arrow indicates self-association among the respective components. Data Collection and Refinement Statistics.  \tSEL1Lcent\t \t \tData set:\tNative\tSe-SAD\t \tPDB accession #:\t5B26\t \t \tX-ray source\tBeamline 7A, PAL\tBeamline 7A, PAL\t \tTemperature (K)\t100\t100\t \tSpace group:\tP21\tP21\t \tCell parameters a, b, c (Å)\t29.13, 110.52, 109.81\t29.51, 110.49, 109.81\t \t \t90.00, 90.61, 90.00\t90.00, 90.74, 90.00\t \tData processing\t \tWavelength (Å)\t1.00000\t0.97923\t \tResolution (Å)\t50-2.60\t50–2.90\t \tRmerge (%)a\t6.1 (38.7)*\t9.4 (40.6)\t \tI/σ\t29.4 (4.6)\t21.0 (3.3)\t \tCompleteness (%)\t99.5 (99.3)\t99.9 (100.0)\t \tRedundancy\t4.1 (4.1)\t3.8 (3.8)\t \tMeasured reflections\t88070\t116951\t \tUnique reflections\t21479\t30823\t \tRefinement statistics\t \tData range (Å)\t30-2.60\t \t \tReflections\t21446\t \t \tNonhydrogen atoms\t5402\t \t \tWater molecules\t47\t \t \tR.m.s. ∆ bonds (Å)b\t0.010\t \t \tR.m.s. ∆ angles (°)b\t1.365\t \t \tR-factor (%)c\t20.7\t \t \tRfree (%)c,d\t27.7\t \t \tRamachandran plot, residues in\t \t \t \tMost favored regions (%)\t92.8\t \t \tAdditional allowed regions (%)\t6.5\t \t \tGenerously allowed regions (%)\t0.7\t \t \tDisallowed regions (%)\t0.0\t \t \t *Highest resolution shell is shown in parenthesis. aRmerge = 100 × ∑h∑i | Ii(h) − \u003cI(h) \u003e|/∑h \u003cI(h)\u003e, where Ii(h) is the ith measurement and \u003cI(h)\u003e is the weighted mean of all measurement of I(h) for Miller indices h. bRoot-mean-squared deviation (r.m.s. ∆) from target geometries. cR-factor = 100 × ∑|FP – FP(calc)|/∑ FP. dRfree was calculated with 5% of the 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+[{"sourceid":"4772114","sourcedb":"","project":"","target":"","text":"Structural basis for the regulation of enzymatic activity of Regnase-1 by domain-domain interactions Regnase-1 is an RNase that directly cleaves mRNAs of inflammatory genes such as IL-6 and IL-12p40, and negatively regulates cellular inflammatory responses. Here, we report the structures of four domains of Regnase-1 from Mus musculus—the N-terminal domain (NTD), PilT N-terminus like (PIN) domain, zinc finger (ZF) domain and C-terminal domain (CTD). The PIN domain harbors the RNase catalytic center; however, it is insufficient for enzymatic activity. We found that the NTD associates with the PIN domain and significantly enhances its RNase activity. The PIN domain forms a head-to-tail oligomer and the dimer interface overlaps with the NTD binding site. Interestingly, mutations blocking PIN oligomerization had no RNase activity, indicating that both oligomerization and NTD binding are crucial for RNase activity in vitro. These results suggest that Regnase-1 RNase activity is tightly controlled by both intramolecular (NTD-PIN) and intermolecular (PIN-PIN) interactions. The initial sensing of infection is mediated by a set of pattern-recognition receptors (PRRs) such Toll-like receptors (TLRs) and the intracellular signaling cascades triggered by TLRs evoke transcriptional expression of inflammatory mediators that coordinate the elimination of pathogens and infected cells. Since aberrant activation of this system leads to auto immune disorders, it must be tightly regulated. Regnase-1 (also known as Zc3h12a and MCPIP1) is an RNase whose expression level is stimulated by lipopolysaccharides and prevents autoimmune diseases by directly controlling the stability of mRNAs of inflammatory genes such as interleukin (IL)-6, IL-1β, IL-2, and IL-12p40. Regnase-1 accelerates target mRNA degradation via their 3′-terminal untranslated region (3′UTR), and also degrades its own mRNA. Regnase-1 is a member of Regnase family and is composed of a PilT N-terminus like (PIN) domain followed by a CCCH-type zinc–finger (ZF) domain, which are conserved among Regnase family members. Recently, the crystal structure of the Regnase-1 PIN domain derived from Homo sapiens was reported. The structure combined with functional analyses revealed that four catalytically important Asp residues form the catalytic center and stabilize Mg2+ binding that is crucial for RNase activity. Several CCCH-type ZF motifs in RNA-binding proteins have been reported to directly bind RNA. In addition, Regnase-1 has been predicted to possess other domains in the N- and C- terminal regions. However, the structure and function of the ZF domain, N-terminal domain (NTD) and C-terminal domain (CTD) of Regnase-1 have not been solved. Here, we performed structural and functional analyses of individual domains of Regnase-1 derived from Mus musculus in order to understand the catalytic activity in vitro. Our data revealed that the catalytic activity of Regnase-1 is regulated through both intra and intermolecular domain interactions in vitro. The NTD plays a crucial role in efficient cleavage of target mRNA, through intramolecular NTD-PIN interactions. Moreover, Regnase-1 functions as a dimer through intermolecular PIN-PIN interactions during cleavage of target mRNA. Our findings suggest that Regnase-1 cleaves its target mRNA by an NTD-activated functional PIN dimer, while the ZF increases RNA affinity in the vicinity of the PIN dimer. Results Domain structures of Regnase-1 We analyzed Rengase-1 derived from Mus musculus and solved the structures of the four domains; NTD, PIN, ZF, and CTD individually by X-ray crystallography or NMR (Fig. 1a–e). X-ray crystallography was attempted for the fragment containing both the PIN and ZF domains, however, electron density was observed only for the PIN domain (Fig. 1c), consistent with a previous report on Regnase-1 derived from Homo sapiens. This suggests that the PIN and ZF domains exist independently without interacting with each other. The domain structures of NTD, ZF, and CTD were determined by NMR (Fig. 1b,d,e). The NTD and CTD are both composed of three α helices, and structurally resemble ubiquitin conjugating enzyme E2 K (PDB ID: 3K9O) and ubiquitin associated protein 1 (PDB ID: 4AE4), respectively, according to the Dali server. Contribution of each domain of Regnase-1 to the mRNA binding activity Although the PIN domain is responsible for the catalytic activity of Regnase-1, the roles of the other domains are largely unknown. First, we evaluated a role of the NTD and ZF domains for mRNA binding by an in vitro gel shift assay (Fig. 1f). Fluorescently 5′-labeled RNA corresponding to nucleotides 82–106 of the IL-6 mRNA 3′UTR and the catalytically inactive mutant (D226N and D244N) of Regnase-1—hereafter referred to as the DDNN mutant—were utilized. Upon addition of a larger amount of Regnase-1, the fluorescence of free RNA decreased, indicating that Regnase-1 bound to the RNA. Based on the decrease in the free RNA fluorescence band, we evaluated the contribution of each domain of Regnase-1 to RNA binding. While the RNA binding ability was not significantly changed in the presence of NTD, it increased in the presence of the ZF domain (Fig. 1f,g and Supplementary Fig. 1). Direct binding of the ZF domain and RNA were confirmed by NMR spectral changes. The fitting of the titration curve of Y314 resulted in an apparent dissociation constant (Kd) of 10 ± 1.1 μM (Supplementary Fig. 2). These results indicate that not only the PIN but also the ZF domain contribute to RNA binding, while the NTD is not likely to be involved in direct interaction with RNA. Contribution of each domain of Regnase-1 to RNase activity In order to characterize the role of each domain in the RNase activity of Regnase-1, we performed an in vitro cleavage assay using fluorescently 5′-labeled RNA corresponding to nucleotides 82–106 of the IL-6 mRNA 3′UTR (Fig. 1g). Regnase-1 constructs consisting of NTD-PIN-ZF completely cleaved the target mRNA and generated the cleaved products. The apparent half-life (T1/2) of the RNase activity was about 20 minutes. Regnase-1 lacking the ZF domain generated a smaller but appreciable amount of cleaved product (T1/2 ~ 70 minutes), while those lacking the NTD did not generate cleaved products (T1/2 \u003e 90 minutes). It should be noted that NTD-PIN(DDNN)-ZF, which possesses the NTD but lacks the catalytic residues in PIN, completely lost all RNase activity (Fig. 1g, right panel), as expected, confirming that the RNase catalytic center is located in the PIN domain. Taken together with the results in the previous section, we conclude that the NTD is crucial for the RNase activity of Regnase-1 in vitro, although it does not contribute to the direct mRNA binding. Dimer formation of the PIN domains During purification by gel filtration, the PIN domain exhibited extremely asymmetric elution peaks in a concentration dependent manner (Fig. 2a). By comparison with the elution volume of standard marker proteins, the PIN domain was assumed to be in equilibrium between a monomer and a dimer in solution at concentrations in the 20–200 μM range. The crystal structure of the PIN domain has been determined in three distinct crystal forms with a space group of P3121 (form I in this study and PDB ID 3V33), P3221 (form II in this study), and P41 (PDB ID 3V32 and 3V34), respectively. We found that the PIN domain formed a head-to-tail oligomer that was commonly observed in all three crystal forms in spite of the different crystallization conditions (Supplementary Fig. 3). Mutation of Arg215, whose side chain faces to the opposite side of the oligomeric surface, to Glu preserved the monomer/dimer equilibrium, similar to the wild type. On the other hand, single mutations of side chains involved in the PIN–PIN oligomeric interaction resulted in monomer formation, judging from gel filtration (Fig. 2a,b). Wild type and monomeric PIN mutants (P212A and D278R) were also analyzed by NMR. The spectra indicate that the dimer interface of the wild type PIN domain were significantly broadened compared to the monomeric mutants (Supplementary Fig. 4). These results indicate that the PIN domain forms a head-to-tail oligomer in solution similar to the crystal structure. Interestingly, the monomeric PIN mutants P212A, R214A, and D278R had no significant RNase activity for IL-6 mRNA in vitro (Fig. 2c). The side chains of these residues point away from the catalytic center on the same molecule (Fig. 2b). Therefore, we concluded that head-to-tail PIN dimerization, together with the NTD, are required for Regnase-1 RNase activity in vitro. Domain-domain interaction between the NTD and the PIN domain While the NTD does not contribute to RNA binding (Fig. 1f,g, and Supplementary Fig. 1), it increases the RNase activity of Regnase-1 (Fig. 1h). In order to gain insight into the molecular mechanism of the NTD-mediated enhancement of Regnase-1 RNase activity, we further investigated the domain-domain interaction between the NTD and the PIN domain using NMR. We used the catalytically inactive monomeric PIN mutant possessing both the DDNN and D278R mutations to avoid dimer formation of the PIN domain. The NMR signals from the PIN domain (residues V177, F210-T211, R214, F228-L232, and F234-S236) exhibited significant chemical shift changes upon addition of the NTD (Fig. 3a). Likewise, upon addition of the PIN domain, NMR signals derived from R56, L58-G59, and V86-H88 in the NTD exhibited large chemical shift changes and residues D53, F55, K57, Y60-S61, V68, T80-G83, L85, and G89 of the NTD as well as side chain amide signals of N79 exhibited small but appreciable chemical shift changes (Fig. 3b and Supplementary Fig. 5). These results clearly indicate a direct interaction between the PIN domain and the NTD. Based on the titration curve for the chemical shift changes of L58, the apparent Kd between the isolated NTD and PIN was estimated to be 110 ± 5.8 μM. Considering the fact that the NTD and PIN domains are attached by a linker, the actual binding affinity is expected much higher in the native protein. Mapping the residues with chemical shift changes reveals the putative PIN/NTD interface, which includes a helix that harbors catalytic residues D225 and D226 on the PIN domain (Fig. 3a). Interestingly, the putative binding site for the NTD overlaps with the PIN-PIN dimer interface, implying that NTD binding can “terminate” PIN-PIN oligomerization (Fig. 2b). An in silico docking of the NTD and PIN domains using chemical shift restraints provided a model consistent with the NMR experiments (Fig. 3c). Residues critical for Regnase-1 RNase activity To gain insight into the residues critical for Regnase-1 RNase activity, each basic or aromatic residue located around the catalytic site of the PIN oligomer was mutated to alanine, and the oligomerization and RNase activity were investigated (Fig. 4). From the gel filtration assays, all mutants except R214A formed dimers, suggesting that any lack of RNase activity in the mutants, except R214A, was directly due to mutational effects of the specific residues and not to abrogation of dimer formation. The W182A, R183A, and R214A mutants markedly lost cleavage activity for IL-6 mRNA as well as for Regnase-1 mRNA. The K184A, R215A, and R220A mutants moderately but significantly decreased the cleavage activity for both target mRNAs. The importance of K219 and R247 was slightly different for IL-6 and Regnase-1 mRNA; both K219 and R247 were more important in the cleavage of IL-6 mRNA than for Regnase-1 mRNA. The other mutated residues—K152, R158, R188, R200, K204, K206, K257, and R258—were not critical for RNase activity. The importance of residues W182 and R183 can readily be understood in terms of the monomeric PIN structure as they are located near to the RNase catalytic site; however, the importance of residue K184, which points away from the active site is more easily rationalized in terms of the oligomeric structure, in which the “secondary” chain’s residue K184 is positioned near the “primary” chain’s catalytic site (Fig. 4). In contrast, R214 is important for oligomerization of the PIN domain and the “secondary” chain’s residue R214 is also positioned near the “primary” chain’s active site within the dimer interface. It should be noted that the putative-RNA binding residues K184 and R214 are unique to Regnase-1 among PIN domains. Molecular mechanism of target mRNA cleavage by the PIN dimer Our mutational experiments indicated that the observed dimer is functional and that the role of the secondary PIN domain is to position Regnase-1-unique RNA binding residues near the active site of the primary PIN domain. If this model is correct, then we reasoned that a catalytically inactive PIN and a PIN lacking the putative RNA-binding residues ought to be inactive in isolation but become active when mixed together. In order to test this hypothesis, we performed in vitro cleavage assays using combinations of Regnase-1 mutants that had no or decreased RNase activities by themselves (Fig. 5). One group consisted of catalytically active PIN domains with mutation of basic residues found in the previous section to confer decreased RNase activity (Fig. 4). These were paired with a DDNN mutant that had no RNase activity by itself. When any members of the two groups are mixed, two kinds of heterodimers can be formed: one is composed of a DDNN primary PIN and a basic residue mutant secondary PIN and is expected to exhibit no RNase activity; the other is composed of a basic residue mutant primary PIN and a DDNN secondary PIN and is predicted to rescue RNase activity (Fig. 5a). When we compared the fluorescence intensity of uncleaved IL-6 mRNA, basic residue mutants W182A, K184A, R214A, and R220A were rescued upon addition of the DDNN mutant (Fig. 5b). Consistently, when we compared the fluorescence intensity of the uncleaved Regnase-1 mRNA, basic residue mutants K184A and R214A were rescued upon addition of the DDNN mutant (Fig. 5c). Rescue of K184A and R214A by the DDNN mutant was also confirmed by a significant increase in the cleaved products. This is particularly significant because the side chains of K184 and R214 in the primary PIN are oriented away from their own catalytic center, while those in the secondary PIN face toward the catalytic center of the primary PIN. R214 is an important residue for dimer formation as shown in Fig. 2, therefore, R214A in the secondary PIN cannot dimerize. According to the proposed model, an R214A PIN domain can only form a dimer when the DDNN PIN acts as the secondary PIN. Taken together, the rescue experiments above support the proposed model in which the head-to-tail dimer is functional in vitro. Discussion We determined the individual domain structures of Regnase-1 by NMR and X-ray crystallography. Although the function of the CTD remains elusive, we revealed the functions of the NTD, PIN, and ZF domains. A Regnase-1 construct consisting of PIN and ZF domains derived from Mus musculus was crystallized; however, the electron density of the ZF domain was low, indicating that the ZF domain is highly mobile in the absence of target mRNA or possibly other protein-protein interactions. Our NMR experiments confirmed direct binding of the ZF domain to IL-6 mRNA with a Kd of 10 ± 1.1 μM. Furthermore, an in vitro gel shift assay indicated that Regnase-1 containing the ZF domain enhanced target mRNA-binding, but the protein-RNA complex remained in the bottom of the well without entering into the polyacrylamide gel. These results indicate that Regnase-1 directly binds to RNA and precipitates under such experimental conditions. Due to this limitation, it is difficult to perform further structural analyses of mRNA-Regnase-1 complexes by X-ray crystallography or NMR. The previously reported crystal structure of the Regnase-1 PIN domain derived from Homo sapiens is nearly identical to the one derived from Mus musculus in this study, with a backbone RMSD of 0.2 Å. The amino acid sequences corresponding to PIN (residues 134–295) are the two non-identical residues are substituted with similar amino acids. Both the mouse and human PIN domains form head-to-tail oligomers in three distinct crystal forms. Rao and co-workers previously argued that PIN dimerization is likely to be a crystallographic artifact with no physiological significance, since monomers were dominant in their analytical ultra-centrifugation experiments. In contrast, our gel filtration data, mutational analyses, and NMR spectra all indicate that the PIN domain forms a head-to-tail dimer in solution in a manner similar to the crystal structure. This inconsistency might be due to difference in the analytical methods and/or protein concentrations used in each experiment, since the oligomer formation of PIN was dependent on the protein concentration in our study. Single mutations to residues involved in the putative oligomeric interaction of PIN monomerized as expected and these mutants lost their RNase activity as well. Since the NMR spectra of monomeric mutants overlaps with those of the oligomeric forms, it is unlikely that the tertiary structure of the monomeric mutants were affected by the mutations. (Supplementary Fig. 4b,c). Based on these observations, we concluded that PIN-PIN dimer formation is critical for Regnase-1 RNase activity in vitro. Within the crystal structure of the PIN dimer, the Regnase-1 specific basic regions in both the “primary” and “secondary” PINs are located around the catalytic site of the primary PIN (Supplementary Fig. 6). Moreover, our structure-based mutational analyses showed these two Regnase-1 specific basic regions were essential for target mRNA cleavage in vitro. The cleavage assay also showed that the NTD is crucial for efficient mRNA cleavage. Moreover, we found that the NTD associates with the oligomeric surface of the primary PIN, docking to a helix that harbors its catalytic residues (Figs 2b and 3a). Taken together, this suggests that the NTD and the PIN domain compete for a common binding site. The affinity of the domain-domain interaction between two PIN domains (Kd = ~10−4 M) is similar to that of the NTD-PIN (Kd = 110 ± 5.8 μM) interactions; however, the covalent connection corresponding to residues 90–133 between the NTD and the primary PIN will greatly enhance the intramolecular domain interaction in the case of full-length Regnase-1. While further analyses are necessary to prove this point, our preliminary docking and molecular dynamics simulations indicate that NTD-binding rearranges the catalytic residues of the PIN domain toward an active conformation suitable for binding Mg2+. In this context, it is interesting that, in response to TCR stimulation, Malt1 cleaves Regnase-1 at R111 to control immune responses in vivo. This result is consistent with a model in which the NTD acts as an enhancer, and cleavage of the linker lowers enzymatic activity dramatically. Based on these structural and functional analyses of Regnase-1 domain-domain interactions, we performed docking simulations of the NTD, PIN dimer, and IL-6 mRNA. We incorporated information from the cleavage site of IL-6 mRNA in vitro is indicated by denaturing polyacrylamide gel electrophoresis (Supplementary Fig. 7a,b). The docking result revealed multiple RNA binding modes that satisfied the experimental results in vitro (Supplementary Fig. 7c,d), however, it should be noted that, in vivo, there would likely be many other RNA-binding proteins that would protect loop regions from cleavage by Regnase-1. The overall model of regulation of Regnase-1 RNase activity through domain-domain interactions in vitro is summarized in Fig. 6. In the absence of target mRNA, the PIN domain forms head-to-tail oligomers at high concentration. A fully active catalytic center can be formed only when the NTD associates with the oligomer surface of the PIN domain, which terminates the head-to-tail oligomer formation in one direction (primary PIN), and forms a functional dimer together with the neighboring PIN (secondary PIN). While further investigations on the domain-domain interactions of Regnase-1 in vivo are necessary, these intramolecular and intermolecular domain interactions of Regnase-1 appear to structurally constrain Regnase-1activity, which, in turn, enables tight regulation of immune responses. Methods Protein expression and purification The DNA fragment encoding Regnase-1 derived from Mus musculus was cloned into pGEX6p vector (GE Healthcare). All the mutants were generated by PCR-mediated site-directed mutagenesis and confirmed by the DNA sequence analyses. As a catalytically deficient mutant, both Asp226 and Asp244 at the catalytic center of PIN were mutated to Asn, which is referred to as DDNN mutant. Regnase-1 was expressed at 16 °C using the Escherichia coli RosettaTM(DE3)pLysS strain. After purification with a GST-affinity resin, an N-terminal GST tag was digested by HRV-3 C protease. NTD was further purified by gel filtration chromatography using a HiLoad 16/60 Superdex 75 pg (GE Healthcare). The other domains were further purified by cation exchange chromatography using Resource S (GE Healthcare), followed by gel filtration chromatography using a HiLoad 16/60 Superdex 75 pg (GE Healthcare). Uniformly 15N or 13C, 15N-double labeled proteins for NMR experiments were prepared by growing E. coli host in M9 minimal medium containing 15NH4Cl, unlabeled glucose and 15N CELTONE® Base Powder (CIL) or 15NH4Cl, 13C6-glucose, and13C, 15N CELTONE® Base Powder (CIL), respectively. X-ray crystallography Crystallization was performed using the sitting drop vapor diffusion method at 20 °C and two crystal forms (I and II) were obtained. In the case of form I crystals, drops (0.5 μl) of 6 mg/ml selenomethionine-labeled Regnase-1 PIN-ZF (residues 134–339 derived from Mus musculus) in 20 mM HEPES-NaOH (pH 6.8), 200 mM NaCl and 5 mM DTT were mixed with reservoir solution consisting of 1 M (NH4)2HPO4, 200 mM NaCl and 100 mM sodium citrate (pH 5.5) whereas in the case of form II crystals, drops (0.5 μl) of 6 mg/ml native Regnase-1 PIN-ZF (residues 134–339) in 20 mM HEPES-NaOH (pH 6.8), 200 mM NaCl and 5 mM DTT were mixed with reservoir solution consisting of 1.7 M NaCl and 100 mM HEPES-NaOH (pH 7.0). Diffraction data were collected at a Photon Factory Advanced Ring beamline NE3A (form I) or at a SPring-8 beamline BL41XU (form II), and were processed with HKL2000. The structure of the form I crystal was determined by the multiple anomalous dispersion (MAD) method. Nine Se sites were found using the program SOLVE; however, the electron density obtained by MAD phases calculated using SOLVE was not good enough to build a model even after density modification using the program RESOLVE. Then the program CNS was used to find additional three Se sites and calculate MAD phases using 12 Se sites. The electron density after density modification using CNS was good enough to build a model. Structure of the form II crystal was determined by the molecular replacement method using CNS and using the structure of the form I crystal as a search model. For all structures, further model building was performed manually with COOT, and TLS and restrained refinement using isotropic individual B factors was performed with REFMAC5 in the CCP4 program suite. Crystallographic parameters are summarized in Supplementary Table 1. NMR measurements All NMR experiments were carried out at 298 K on Inova 500-MHz, 600-MHz, and 800-MHz spectrometer (Agilent). The NMR data were processed using the NMRPipe, the Olivia (fermi.pharm.hokudai.ac.jp/olivia/), and the Sparky program (Sparky3, University of California, San Francisco). For structure calculation, NOE distance restraints were obtained from 3D 15N-NOESY-HSQC (100 ms mixing time for the NTD, 75 ms mixing time for the ZF domain and the CTD) and 13C-NOESY-HSQC spectra (100 ms mixing time for the NTD, 75 ms mixing time for the ZF domain and the CTD). The NMR structures were determined using the CANDID/CYANA2.1. Dihedral restraints were derived from backbone chemical shifts using TALOS. For the domain-domain interaction analyses between the NTD and the PIN domain, 1H-15N HSQC spectra of uniformly 15N-labeled proteins in the concentration of 100 μM were obtained in the presence of 3 or 6 molar equivalents of unlabeled proteins. Preparation of RNAs The fluorescently labeled RNAs at the 5′-end by 6-FAM were purchased from SIGMA-ALDORICH. The RNA sequences used in this study were shown below. IL-6 mRNA 3′UTR (82–106): 5′-UGUUGUUCUCUACGAAGAACUGACA-3′ (25 nts) Regnase-1 mRNA 3′UTR (191–211): 5′- CUGUUGAUACACAUUGUAUCU-3′ (21 nts) Electrophoretic mobility shift assay Catalytically deficient Regnase-1 proteins, containing DDNN mutations, and 5′-terminally 6-FAM labeled RNAs were incubated in the RNA-binding buffer (20 mM HEPES-NaOH (pH 6.8), 150 mM NaCl, 1 mM DTT, 10% glycerol (v/v), and 0.1% NP-40 (v/v)) at 4 °C for 30 minutes, then analyzed by non-denaturing polyacrylamide gel electrophoresis. The electrophoreses were performed at 4 °C using the 7.5% polyacrylamide (w/v) gel (monomer : bis = 29 : 1) in the electrophoresis buffer (25 mM Tris-HCl (pH 7.5) and 200 mM glycine). The fluorescence of 6-FAM labeled RNA was directly detected at the excitation wavelength of 460 nm with a fluorescence filter (Y515-Di) using a fluoroimaging analyzer (LAS-4000 (FUJIFILM)). The fluorescence intensity of each sample was quantified using ImageJ software. In vitro RNA cleavage assay Regnase-1 (2 μM) and 5′-terminally 6-FAM labeled RNA (1 μM) were incubated in the RNA-cleavage buffer (20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 5 mM MgCl2, and 1 mM DTT) at 37 °C. For the assay using combinations of Regnase-1 mutants, equimolar amounts of Regnase-1 mutants (2 μM each) were mixed with fluorescently labeled RNA (1 μM). After incubation for 30–120 minutes, the reaction was stopped by the addition of 1.5-fold volume of denaturing buffer containing 8 M urea and 100 mM EDTA, and samples were boiled. The electrophoreses were performed at room temperature using the 8 M urea containing denaturing gel with 20% polyacrylamide (w/v) (monomer : bis = 19 : 1) in 0.5 × TBE as the electrophoresis buffer. Docking calculations For docking NTD to PIN, OSCAR-star was first used to rebuild sidechains in the head-to-tail PIN dimer. Docking was carried out by surFit (http://sysimm.ifrec.osaka-u.ac.jp/docking/main/) with restraints obtained from NMR data (Fig. 3a,b) as follows. NTD: R56, L58, G59, V86, K87, H88; PIN: V177, F210, T211, R214, F228, I229, V230, K231, L232, F234, D235, S236. Top-scoring model was selected. For docking IL-6 mRNA 3′UTR to the PIN dimer, each domain of the PIN dimer structure was superimposed onto the PIN dimer of the human X-ray structure (PDB ID: 3V34) in order to graft both water molecules and Mg2+ ions to the mouse model. Each IL-6 representative structure was submitted to the HADDOCK 2.0 server, for total of 10 independent jobs. In order to be consistent with the cleavage assay, active residues consisted of all nucleotides in RNA, Mg2+ and W182, R183, K184, R188, R214, R215, K219, R220, and R247 in the protein. Docked models were selected based on the following criteria: one heavy atom within 7, 8, or 9th nucleotide from the 5′ end was \u003c5 Å from the Mg2+ ion on the primary PIN. Further classification was done manually in order to divide the selected models into two clusters. Additional Information Accession codes: The crystal structure of the Regnase-1 PIN domain has been deposited in the Protein Data Bank (accession codes: 5H9V (Form I) and 5H9W (Form II)). The chemical shift assignments of the NTD, the ZF domain, and the CTD have been deposited at Biological Magnetic Resonance Bank (accession codes: 25718, 25719, and 25720, respectively), and the coordinates for the ensemble have been deposited in the Protein Data Bank (accession codes: 2N5J, 2N5K, and 2N5L, respectively).  How to cite this article: Yokogawa, M. et al. Structural basis for the regulation of enzymatic activity of Regnase-1 by domain-domain interactions. Sci. Rep. 6, 22324; doi: 10.1038/srep22324 (2016). Supplementary Material Pathogen recognition and innate immunity. Recognition of microorganisms and activation of the immune response Genetic analysis of host resistance: Toll-like receptor signaling and immunity at large Zc3h12a is an RNase essential for controlling immune responses by regulating mRNA decay Interleukin-1-inducible MCPIP protein has structural and functional properties of RNase and participates in degradation of IL-1beta mRNA MCPIP1 down-regulates IL-2 expression through an ARE-independent pathway mRNA degradation by the endoribonuclease Regnase-1/ZC3H12a/MCPIP-1 The IκB kinase complex regulates the stability of cytokine-encoding mRNA induced by TLR-IL-1R by controlling degradation of regnase-1 A novel CCCH-zinc finger protein family regulates proinflammatory activation of macrophages MCP-1-induced protein-1, an immune regulator Structural study of MCPIP1 N-terminal conserved domain reveals a PIN-like RNase Specificity of RNA binding by CPEB: requirement for RNA recognition motifs and a novel zinc finger Interactions of CCCH zinc finger proteins with mRNA: non-binding tristetraprolin mutants exert an inhibitory effect on degradation of AU-rich element-containing mRNAs Recognition of the mRNA AU-rich element by the zinc finger domain of TIS11d Multiple modes of RNA recognition by zinc finger proteins Monocyte chemoattractant protein-1 induces a novel transcription factor that causes cardiac myocyte apoptosis and ventricular dysfunction MCP-induced protein 1 deubiquitinates TRAF proteins and negatively regulates JNK and NF-kappaB signaling Dali server: conservation mapping in 3D Malt1-induced cleavage of regnase-1 in CD4(+) helper T cells regulates immune activation Processing of X-ray Diffraction Data Collected in Oscillation Mode Automated MAD and MIR structure solution Maximum-likelihood density modification Crystallography \u0026 NMR system: A new software suite for macromolecular structure determination Features and development of Coot Refinement of macromolecular structures by the maximum-likelihood method Overview of the CCP4 suite and current developments NMRPipe: a multidimensional spectral processing system based on UNIX pipes Automated NMR structure calculation with CYANA Protein backbone angle restraints from searching a database for chemical shift and sequence homology Fast and accurate prediction of protein side-chain conformations Author Contributions F.I. supervised the overall project. M.Y., T.T., Y.E., D.M.S., O.T., S.A. and F.I. designed the research; M.Y. and T.T. performed the research; M.Y., T.T., N.N.N., H.K., K.Y., D.M.S. and F.I. analyzed the data; and M.Y., N.N.N., H.K., K.Y., D.M.S. and F.I. wrote the paper. All authors reviewed the manuscript. Structural and functional analyses of Regnase-1. (a) Domain architecture of Regnase-1. (b) Solution structure of the NTD. (c) Crystal structure of the PIN domain. Catalytic Asp residues were shown in sticks. (d) Solution structure of the ZF domain. Three Cys residues and one His residue responsible for Zn2+-binding were shown in sticks. (e) Solution structure of the CTD. All the structures were colored in rainbow from N-terminus (blue) to C-terminus (red). (f) In vitro gel shift binding assay between Regnase-1 and IL-6 mRNA. Fluorescence intensity of the free IL-6 in each sample was indicated as the percentage against that in the absence of Regnase-1. (g) Binding of Regnase-1 and IL-6 mRNA was plotted. The percentage of the bound IL-6 was calculated based on the fluorescence intensities of the free IL-6 quantified in (f). (h) In vitro cleavage assay of Regnase-1 to IL-6 mRNA. Fluorescence intensity of the uncleaved IL-6 mRNA was indicated as the percentage against that in the absence of Regnase-1. Head-to-tail oligomer formation of the PIN domain is crucial for the RNase activity of Regnase-1. (a) Gel filtration analyses of the PIN domain. Elution volumes of the standard marker proteins were indicated by arrows at the upper part. (b) Dimer structure of the PIN domain. Two PIN molecules in the crystal were colored white and green, respectively. Catalytic residues and mutated residues were shown in sticks. Residues important for the oligomeric interaction were colored red, while R215 that was dispensable for the oligomeric interaction was colored blue. (c) RNase activity of monomeric mutants for IL-6 mRNA was analyzed. Domain-domain interaction between the NTD and the PIN domain. (a) NMR analyses of the NTD-binding to the PIN domain. The residues with significant chemical shift changes were labeled in the overlaid spectra (left) and colored red on the surface and ribbon structure of the PIN domain (right). Pro and the residues without analysis were colored black and gray, respectively. (b) NMR analyses of the PIN-binding to the NTD. The residues with significant chemical shift changes were labeled in the overlaid spectra (left) and colored red, yellow, or green on the surface and ribbon structure of the NTD. S62 was colored gray and excluded from the analysis, due to low signal intensity. (c) Docking model of the NTD and the PIN domain. The NTD and the PIN domain are shown in cyan and white, respectively. Residues in close proximity (\u003c5 Å) to each other in the docking structure were colored yellow. Catalytic residues of the PIN domain are shown in sticks, and the residues that exhibited significant chemical shift changes in (a,b) were labeled. Critical residues in the PIN domain for the RNase activity of Regnase-1. (a) In vitro cleavage assay of basic residue mutants for IL-6 mRNA. The results indicate mean ± SD of four independent experiments. (b)\nIn vitro cleavage assay of basic residue mutants for Regnase-1 mRNA. The results indicate mean ± SD of three independent experiments. The fluorescence intensity of the uncleaved mRNA was quantified and the results were mapped on the PIN dimer structure. Mutated basic residues were shown in sticks and those with significantly reduced RNase activities were colored red or yellow. Heterodimer formation by combination of the Regnase-1 basic residue mutants and the DDNN mutant restored the RNase activity. (a) Cartoon representation of the concept of the experiment. (b) In vitro cleavage assay of Regnase-1 for IL-6 mRNA. (c) In vitro cleavage assay of Regnase-1 for Regnase-1 mRNA. The results indicate mean ± SD of three independent experiments. The fluorescence intensity of the uncleaved mRNA was quantified and the results were mapped on the PIN dimer. The mutations whose RNase activities were not increased in the presence of DDNN mutant were colored in blue on the primary PIN. The mutations whose RNase activities were restored in the presence of DDNN mutant were colored in red or yellow on the primary PIN. Schematic representation of regulation of the Regnase-1 catalytic activity through the domain-domain 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diff --git a/annotated_BioC_JSON/PMC4773095_ann.json b/annotated_BioC_JSON/PMC4773095_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..7ec64cbddee2966dde2f999b3d91b29b28496093
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4773095_ann.json
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+[{"sourceid":"4773095","sourcedb":"","project":"","target":"","text":"Structure of the Response Regulator NsrR from Streptococcus agalactiae, Which Is Involved in Lantibiotic Resistance Lantibiotics are antimicrobial peptides produced by Gram-positive bacteria. Interestingly, several clinically relevant and human pathogenic strains are inherently resistant towards lantibiotics. The expression of the genes responsible for lantibiotic resistance is regulated by a specific two-component system consisting of a histidine kinase and a response regulator. Here, we focused on a response regulator involved in lantibiotic resistance, NsrR from Streptococcus agalactiae, and determined the crystal structures of its N-terminal receiver domain and C-terminal DNA-binding effector domain. The C-terminal domain exhibits a fold that classifies NsrR as a member of the OmpR/PhoB subfamily of regulators. Amino acids involved in phosphorylation, dimerization, and DNA-binding were identified and demonstrated to be conserved in lantibiotic resistance regulators. Finally, a model of the full-length NsrR in the active and inactive state provides insights into protein dimerization and DNA-binding. Introduction The dramatic rise in antibiotic resistance has posed a major threat to the treatment of infectious diseases. This has led to the search for novel antibiotics that can be used as pharmaceuticals against human pathogenic bacteria. One of the potential antibiotic alternatives are lantibiotics. Lantibiotics are small antimicrobial peptides (30–50 amino acids in length), which are produced by several Gram-positive bacterial strains. They are post-translationally modified and contain specific lanthionine/methyl-lanthionine rings, which are crucial for their high antimicrobial activity. Lantibiotics are for example highly effective against various Gram-positive, human pathogenic bacteria including Streptococcus pneumoniae and several methicillin-resistant Staphylococcus aureus (MRSA) strains. The high potency of lantibiotics for medical usage has already been noticed, and several lantibiotics are already included in clinical trials. Their high potency is highlighted by the fact that, although being extensively used in food industry, resistance has not been described so far. Nisin is the most prominent member of the lantibiotic family and is able to inhibit cell growth, penetrates the membranes of various Gram-positive bacteria, and is characterized by five specific (methyl-)lanthionine rings, which are crucial for stability and activity in the nanomolar range. Thus, the lantibiotic producer strains have an inbuilt self-protection mechanism (immunity) to prevent cell death caused due to the action of its cognate lantibiotic. This immunity system consists of a membrane–associated lipoprotein (usually referred to as LanI) and/or an ABC transporter (termed as LanFEG and comprising three subunits). Although some lantibiotics such as Pep5, epicidin, epilancin, and lactocin S only require LanI for immunity, other lantibiotics with a dual mode of action involving pore formation and lipid II binding such as nisin, subtilin, epidermin, gallidermin, and lacticin 3147 require additionally the presence of LanFEG. Examples for LanFEG are NisI and NisFEG of the nisin system, SpaI and SpaFEG conferring immunity towards subtilin, and PepI constituting the immunity system of Pep5 producing strains. Structural data are reported for the immunity proteins NisI from Lactococcus lactis, SpaI from Bacillus subtilis and MlbQ from the lantibiotic NAI-107 producer strain Microbispora ATCC PTA-5024. Recently, gene clusters were identified in certain clinically relevant human pathogenic strains such as Streptococcus agalactiae, S. aureus, and others that confer inherent resistance against specific lantibiotics such as nisin and resemble the genetic architecture of the lantibiotic immunity genes found in the producing strains. Within these resistance operons, genes encoding for a membrane-associated protease and an ABC transporter were identified. Expression of these proteins provides resistance against lantibiotics. Recently, the structure of SaNSR from S. agalactiae was solved which provides resistance against nisin by a protease activity. Furthermore, the upregulation of these genes is mediated by a specific two-component system (TCS) similar to the one found in lantibiotic producing strains, consisting of a sensor histidine kinase (HK) and a response regulator (RR), apparently mediate the expression of the resistance proteins: HK senses the external lantibiotic and, upon receiving the stimuli, auto-phosphorylates at a conserved histidine residue within the cytosol; this high-energetic phosphoryl group is then transferred to the associated RR inducing a conformational change there, which activates the RR to evoke the cellular response. Bacteria have the ability to sense and survive various environmental stimuli through adaptive responses, which are regulated by TCSs. These processes include drug resistance, quorum-sensing, phosphate uptake, sporulation, and osmoregulation. The absence of TCSs within mammals makes them unique targets for novel antimicrobial drugs. The expression of the lantibiotic-resistance genes via TCS is generally regulated by microorganism-specific lantibiotics, which act via external stimuli. Some examples of TCS are: BraRS in S. aureus which is induced by bacitracin, nisin and nukacin-ISK-1 resistance, BceRS in Bacillus spp. that is induced by actagardine and mersacidin resistance, LcrRS in Streptococcus mutans induced by nukacin-ISK-1 and lacticin 481 and LisRK of Listeria monocytogenes induced by nisin resistance. Furthermore, multiple lantibiotics can induce the TCS CprRK from Clostridium difficile, leading to the expression of the genes localized on the cpr operon, resulting in resistance against several lantibiotics of which nisin, gallidermin, subtilin, and mutacin 1140 are some examples. Interestingly, the histidine kinase contains two-transmembrane helices but lacks an extracellular sensory domain, and are therefore known as ‘intramembrane-sensing’ histidine kinases. It has been suggested that in addition to conferring general resistance against lantibiotics, the BceAB-type transporters assist in signalling as via the presence of a large extracellular domain within the transmembrane segment indicated by experimental evidence from various systems. The recently discovered nsr gene cluster of the human pathogen S. agalactiae encodes for the resistance protein NSR and the ABC transporter NsrFP, both conferring resistance against nisin. Homologous operons have been identified in various human pathogenic strains such as Staphylococcus epidermis and Streptococcus ictaluri based on the high sequence identity of NSR and NsrFP. In this gene cluster, the TCS NsrRK is responsible for the expression of the nsr and nsrFP genes. The similarity of the TCS within all the described nisin resistance operons suggests an expression specifically induced by nisin. Thus, NsrRK might be a useful target to combat inherently pathogenic lantibiotic-resistant strains. Generally, RRs consist of two distinct structural domains, a receiver domain (RD) and an effector domain (ED), that are separated from each other by a flexible linker. RDs contain a highly conserved aspartate residue, which acts as a phosphoryl acceptor that becomes phosphorylated by the kinase domain of the histidine kinase upon reception of an external signal. The ED is thereby activated and binds to the designated promoters, thus initiating transcription of the target genes. The RRs are classified into different subfamilies depending on the three-dimensional structure of their EDs. The OmpR/PhoB subfamily is the largest subgroup of RRs and comprises approximately 40% of all response regulators in bacteria. All their members are characterized by a winged helix-turn-helix (wHTH) motif. Although numerous structures of the single domains are known, only a few structures of full-length OmpR/PhoB-type RRs have been determined: RegX3 (PDB code: 2OQR), MtrA (PDB code: 2GWR), PrrA (PDB code: 1YS6) and PhoP (PDB code: 3R0J) from Mycobacterium tuberculosis; DrrB (PDB code: 1P2F) and DrrD (PDB code: 1KGS) from Thermotoga maritima; and KdpE from Escherichia coli (PDB code: 4KNY). The various structures of RRs reveal that in addition to being in either “inactive” or “active” state, the RRs can also exist in two distinct conformations: “open” and “closed”. MtrA and PrrA exhibit a very compact, closed structure with the DNA-binding sequence, called recognition helix, of the ED being inaccessible to DNA. The structures of DrrD and DrrB exist in an open conformation, here the recognition helix is fully exposed, suggesting that RRs are flexible in solution and can adopt multiple conformations. Here, we describe the crystal structures of the N-terminal RD and the C-terminal ED of the lantibiotic resistance-associated RR NsrR from S. agalactiae. NsrR is part of the nisin resistance operon. The expression of the genes of this operon is induced by a TCS consisting of the HK NsrK and the RR NsrR. Based on the crystal structures of both the domains, modeling was employed to shed light on the putative DNA-bound state of full-length NsrR. Materials and Methods Cloning, expression and purification NsrR NsrR was constructed, expressed, and purified as described previously. In brief, the nsrR gene (accession no. HG939456.1) from S. agalactiae COH1 was ligated into the expression vector pET24a allowing expression in E. coli with a His6-tag introduced at the C-terminus. The resulting plasmid pET24a-NsrR was transformed into E. coli BL21 (DE3) for expression. A single transformed colony was inoculated into 20 ml LB media containing 30 μg/ml kanamycin. The culture was grown for 14 h at 310 K with shaking at 200 rpm. 4 l LB media with 30 μg/ml kanamycin were inoculated with the overnight culture at an OD600 of 0.05 and grown at 310 K with shaking at 170 rpm until an OD600 of 0.3 was reached. Subsequently, temperature was lowered to 291 K, and cells were further grown until an OD600 of 0.8 was reached before inducing the expression by addition of 1 mM IPTG. Cells were further grown for 15 h and harvested by centrifugation at 8000 rpm for 20 min at 277 K. The harvested cell pellet was re-suspended in 10 ml of buffer A (50 mM Tris pH 8.0, 50 mM NaCl, 2 mM PMSF and 10% (v/v) glycerol) and 10 mg of DNase (Deoxyribonuclease I from bovine pancreas, Sigma Aldrich) was added. Cells were lysed using a cell disruptor (Constant Cell Disruption Systems, United Kingdom) at 2.6 × 105 kPa. The lysate was centrifuged at 42000 rpm for 60 min using a Ti60 rotor to remove non-lysed cells and cell debris. 20 mM imidazole was added to the cleared lysate prior to applying it onto a Ni2+ loaded Hi-Trap HP Chelating column (GE Healthcare) pre-equilibrated with buffer B (20 mM Tris pH 8.0, 250 mM NaCl and 20 mM imidazole, 2 mM PMSF). The column was washed with six column volumes of buffer B. Protein was eluted with a linear gradient of imidazole from 20 mM to 400 mM in buffer B. The fractions containing NsrR were pooled and concentrated up to 8 mg/ml in an Amicon centrifugal filter concentrator with a 10 kDa cut-off membrane (Millipore). The concentrated protein was further purified by size exclusion chromatography using a Superdex 200 GL 10/300 column (GE Healthcare), equilibrated with buffer C (25 mM Tris pH 9.0, 50 mM NaCl, 2 mM PMSF). The eluted protein fractions were pooled and concentrated to 11 mg/ml as described above. The purity of the protein was analyzed with 15% SDS-PAGE using colloidal Coomassie blue staining. Crystallization of NsrR Crystals were obtained by using 1 μl of protein solution (concentration of 6.0 mg/ml) mixed with 1 μl of reservoir solution using the hanging-drop vapor diffusion method at 285 K. The reservoir solution contained PEG 20000 (11, 13, 15, 17 and 21% (w/v)) and 0.1 M MES pH (6.0, 6.5, 7.0 and 7.5). Crystals were obtained after three weeks and grew to their maximum dimensions within one month. Two different crystal forms, rectangular plate-shaped crystals and thin plates, were observed in the same drop. Both crystals forms were transferred into a buffer containing the reservoir solution plus 30% (v/v) ethylene glycol for 5 min prior to flash cooling using liquid nitrogen. For phasing, 20 mM tetra-chloro platinate IV (Hampton Research) was added to the crystallization drop, and the rectangular plate-shaped crystals were soaked for 30 min. The crystals with no obvious optical damage were harvested and flash-cooled in liquid nitrogen following the procedure above. Data collection Initially crystals were screened for quality at beamline P13 (DESY, EMBL Hamburg). All X-ray diffraction data were collected at beamline ID23eh1 of the European Synchrotron Radiation Facility (ESRF), Grenoble. All data sets were processed and scaled using XDS and XSCALE software package. Data sets from both native crystal forms were collected at 100 K. A single-wavelength anomalous dispersion (SAD) dataset from a single heavy-atom derivatized crystal (rectangular plate-shaped crystal) was collected at 1.0714 Å at 100 K. Diffraction data up to 1.7 Å was used for heavy atom localization and subsequent phasing. Structure determination of NsrR The structure of the thin plate-shaped crystals was solved by molecular replacement using the structure of the receiver domain of PhoB (PDB entry: 1B00) as a model to phase the native data set at 1.8 Å resolution. The model generated was refined manually in COOT followed by iterative cycles of refinement using the program phenix.refine. Manual adjustments between the refinement cycles were performed with the program COOT and Ramachandran validation was done using MolProbity. The SAD dataset of the rectangular plate-shaped crystal was used for phasing via the Auto-Rickshaw server. The initial model was further built and refined manually using COOT and phenix.refine from the Phenix package with iterative cycles of refinement. This model was used to phase the native data set of the rectangular plate-shaped crystals at a resolution of 1.6 Å. Data collection and refinement statistics are listed in Table 1 and all images of the models were prepared using PyMOL. Data collection, phasing, and refinement statistics for the receiver and effector domains of NsrR. \tNsrR-RD (native)\tNsrR-ED (native)\tNsrR-ED (SAD dataset)\t \tData collection\t\t\t\t \tSpace group\tP 21 21 2\tP 21 21 2\tP 21 21 2\t \tCell dimensions\t\t\t\t \t    a, b, c (Å)\t57.0 107.1 39.4\t56.3 60.4 56.8\t56.3 60.6 56.7\t \t    α, β, γ (°)\t90.0 90.0 90.0\t90.0 90.0 90.0\t90.0 90.0 90.0\t \tWavelength (λ)\t1.0688\t0.9677\t1.0714\t \tResolution (Å)\t39.48–1.80 (1.86–1.80)\t56.85–1.60 (1.65–1.60)\t100.00–1.70 (1.75–1.70)\t \tRmergea\t3.4 (33.3)\t4.8 (30.5)\t6.8 (97.0)\t \tI /σ(I)\t26.2 (5.1)\t18.2 (4.6)\t21.6 (1.7)\t \tCompleteness (%)\t98.8 (98.8)\t99.5 (99.7)\t98.8 (90.2)\t \tRedundancy\t4.8 (4.8)\t4.8 (4.9)\t11.7 (6.5)\t \tStructure Refinement\t\t\t\t \tResolution (Å)\t39.48–1.80 (1.86–1.80)\t56.85–1.60 (1.65–1.60)\t\t \tNo. of reflections\t109201 (10602)\t124810 (12438)\t\t \tCC1/2\t0.999 (0.924)\t0.999 (0.923)\t\t \tRworkb / Rfreeb\t0.17 (0.20)/ 0.22 (0.27)\t0.18 (0.22)/ 0.22 (0.27)\t\t \tNo. of atoms\t2027\t1843\t\t \t    Macromolecules\t1894\t1580\t\t \t    Ligand/ion\t20\t8\t\t \t    Water\t113\t255\t\t \tB-factors (Å2)\t28.3\t21.7\t\t \t    Macromolecules\t27.7\t20.2\t\t \t    Ligand/ion\t34.0\t38.6\t\t \t    Solvent\t36.9\t30.4\t\t \tR.m.s. deviations\t\t\t\t \t    Bond lengths (Å)\t0.007\t0.008\t\t \t    Bond angles (°)\t1.11\t1.18\t\t \tRamachandran plot (%)\t\t\t\t \t    Favored\t99.0\t97.0\t\t \t    Allowed\t1.0\t2.48\t\t \t    Outliers\t0.0\t0.52\t\t \t Values in parentheses are for the highest resolution shell. a Rmerge is defined as Rsym = ∑hkl∑i|Ii(hkl) − ⟨I(hkl)⟩|/∑hkl∑iIi(hkl) and b RF as Rf = ∑hkl‖Fobs|−|Fcalc‖/∑hkl|Fobs| Accession numbers Coordinates and structure factors have been deposited in the PDB with accession numbers 5DCL (NsrR-RD) and 5DCM (NsrR-ED). Results and Discussion NsrR was expressed and purified as described, resulting in a homogenous protein as observed by size exclusion chromatography (Fig 1A), with a yield of 2 mg per liter of cell culture. By calibrating the column with proteins of known molecular weight the NsrR full length protein elutes as a dimer. The purified NsrR protein has a theoretical molecular mass of 27.7 kDa and was \u003e98% pure as assessed by SDS-PAGE (Fig 1B, indicated by *). Surprisingly, over time NsrR degraded into two distinct fragments as visible on SDS-PAGE analysis using the same purified protein sample after one week (Fig 1C, indicated by ** and ***, respectively). This was also observed by size exclusion chromatography where a peak at an elution time of 18 min appeared (Fig 1A). Both bands were subjected to mass spectrometry analysis. The analysis revealed that the larger fragment (**) represents the N-terminal receiver domain (residues 1–119; referred to as NsrR-RD) whereas the smaller fragment (***) contained the C-terminal DNA-binding effector domain of NsrR (residues 129–243 including 21 amino acids derived from the expression tag; referred to as NsrR-ED) (Fig 1C). Residues 120–128 form the linker connecting the RD and ED. Such a cleavage of the full-length RR into two specific domains is not unusual and has been previously reported for other RRs as well. Mass spectrometry analysis did not reveal the presence of any specific protease in the purified NsrR sample. Furthermore, addition of a protease inhibitor, such as PMSF (Phenylmethylsulfonyl fluoride) and AEBSF {4-(2-Aminoethyl) benzenesulfonyl fluoride hydrochloride}, even at high concentrations, did not inhibit proteolysis (data not shown). Purification of NsrR and SDS PAGE analysis of purified NsrR directly and one week after purification. (a) Elution profile of size-exclusion chromatography step of NsrR. The y-axis represents the UV absorption of the protein at 280 nm, while the x-axis represents the elution volume. a, b, c refer to the protein standards dextran blue (2,000 kDa), BSA (67 kDa), and lysozyme (14.3 kDa), respectively. The bold line represents the chromatogram of freshly purified NsrR while the dashed line shows the chromatogram of the same NsrR protein after one week. (b) Freshly purified NsrR protein, and (c) NsrR protein after one week. Lanes: M represents the PAGE Ruler Unstained Ladder; 1: NsrR after a two-step purification; 2: NsrR one week after purification. * corresponds to full-length NsrR protein at 27 kDa, while ** and *** correspond to the NsrR-RD and NsrR-ED domain at around 13 kDa, respectively. Since formation of the crystals took around one month, it is not surprising that this cleavage also occurred in the crystallization drop. NsrR was crystallized yielding two crystal forms, which were distinguishable by visual inspection. Initially, we tried to solve the structure of NsrR by molecular replacement, which was not successful. Therefore, we tried heavy atom phasing using a platinum compound. This succeeded for the rectangular plate-shaped crystals. After the structure was solved, it became evident that these crystals contained two monomers of the ED of NsrR in the asymmetric unit. We also tried to solve the structure of the thin plate-shaped crystals with this template, but the resulting model generated was not sufficient. Therefore, we thought that these crystals contained the N-terminal domain of NsrR and successfully phased this dataset using molecular replacement with the N-terminal domain of PhoB (PDB code: 1B00; as a template. This approach revealed that this crystal form indeed contained two monomers of the RD of NsrR in the asymmetric unit. Since both crystals forms were obtained in the same drop it is not surprising that, when dissolving several crystals and performing subsequent mass-spectrometry to identify the protein in the crystals, it yielded peptide fragments throughout the NsrR sequence. In summary, the two crystal forms contained one of the two domains, respectively, such that both domains were successfully crystallized. We determined the crystal structures of NsrR-RD and NsrR-ED separately. However, a part of the linker region (residues 120–128; 120RRSQQFIQQ128; underlined are the amino acid residues not visible in either domain) could not be traced in the electron density. Overall structure of the N-terminal NsrR receiver domain (NsrR-RD) The structure of the NsrR-RD was determined at a resolution of 1.8 Å (Table 1). The Rwork and Rfree values after refinement were 0.17 and 0.22, respectively. Ramachandran validation revealed that all residues (100%, 236 amino acids) were in the preferred or allowed regions. The structure contained many ethylene glycol molecules arising from the cryo-protecting procedure. Data collection and refinement statistics are listed in Table 1. The asymmetric unit contains two copies of NsrR-RD. Although the entire N-terminal receiver domain is composed of residues Met1-Leu119, only residues Asn4 to Arg121 of chain A (including residues Arg120 and Arg121 of the linker) and Gln5 to Ser122 of chain B (including residues Arg120 until Ser122 of the linker) could be traced in the electron density of NsrR-RD. For Asn85, Asp86, and Glu87 of chain A, poor electron density was observed for the side chains and, thus, these side chains were deleted during refinement and are not present in the final structure. Since the two monomers of NsrR-RD were virtually identical (rmsd of 0.6 Å over 116 Cα atoms for the two monomers). Therefore, the overall structure is described for monomer A only. NsrR-RD structurally adopts a αβ doubly-wound fold previously observed in OmpR/PhoB type regulators. Five β-strands (β1-β5) are arranged in a parallel fashion constituting the central core of the structure, which is surrounded by two α-helices (α1 and α5) on one and three helices (α2, α3, α4) on the other side (Fig 2). The NsrR-RD structure shows a β1-α1-β2-α2-β3-α3-β4-α4-β5-α5 topology as also observed for other RRs. Structure of NsrR-RD. Cartoon representation of the helices (α1 – α5) and β-sheets (β1 - β5). Structural areas with the highest variations to the receiver domains of DrrB (pink, 1P2F), MtrA (grey, 2GWR), and PhoB (blue, 1B00) are marked in separate boxes. Comparison with structures of other receiver domains NsrR belongs to the OmpR/PhoB family of RRs. The receiver domain of NsrR was superimposed with other structurally characterized receiver domains from the OmpR/PhoB family, such as DrrB, KdpE, MtrA, and the crystal structure of only the receiver domain of PhoB. The rmsd of the overlays and the corresponding PDB codes used are highlighted in Table 2. Superimposition of the structures revealed that helix α4 is slightly rotated outward in NsrR-RD (Fig 2). In receiver domains of response regulators, helix α4 has been shown to be a crucial part of the dimerization interface. Furthermore, helix α4 in NsrR is shorter than in other RRs. The first helical turn is unwound and adopts an unstructured region (see Fig 2). A slightly outward rotation or unwinding of helix α4 has been observed in the structures of other RD of regulators. For example, the structure of BaeR and RegX3 displayed a completely unwound helix α4. In the structure of DrrD, helix α4 is only partially displaced. In the receiver domain of NsrR, helix α4 is also partially displaced but in a different direction (S1 Fig). Inspection of the crystal contacts revealed no major interactions in this region that could have influenced the orientation of helix α4. Furthermore, NsrR is crystallized as a monomer, and investigation of the symmetry-related molecules did not reveal a functional dimer within the crystal. This could explain the flexibility and thereby the different orientation of helix α4 in NsrR. The structures of the RD and ED domains of NsrR aligned to other response regulators. The rmsd values of the superimpositions of the structures of NsrR-RD and NsrR-ED with the available structures of members of the OmpR/PhoB subfamily are highlighted. *Seq ID (%) corresponds to the full-length protein sequence. Protein\tPDB\tZ-score\tRMSD (Å)\tNumber of residues (total number of residues)\tSeq. ID (%)*\tReference\t \tReceiver domain\t \tKdpE\t4KNY\t18.8\t1.9\t117 (222)\t28\t\t \tYycF\t2ZWM\t18.3\t1.7\t115 (120)\t35\t\t \tYycF\t3F6P\t18.1\t1.7\t114 (120)\t35\t\t \tDivK\t1M5T\t18.1\t1.9\t116 (123)\t27\t\t \tKdpE\t1ZH2\t18.0\t1.9\t115 (120)\t28\t\t \tPhoB\t1B00\t17.0\t1.9\t113 (122)\t30\t\t \tEffector domain\t \tPhoB\t1GXQ\t13.7\t1.7\t92 (105)\t30\t\t \tPhoP\t2PMU\t13.4\t1.7\t87 (93)\t30\t\t \tPhoB\t2Z33\t13.3\t1.8\t92 (104)\t30\t\t \tPhoB (DNA bound)\t1GXP\t13.3\t2.0\t92 (101)\t30\t\t \tSaeR\t4IXA\t13.0\t2.1\t94 (102)\t29\tNot available\t \tRstA\t4NHJ\t11.8\t1.9\t85 (101)\t29\t\t \tKdpE\t4KNY\t11.5\t2.6\t86 (222)\t28\t\t \tFull-length Response Regulators\t \t\tPDB code\tN-terminal rmsd (Å)\tC-terminal rmsd (Å)\tDNA bound\t\tReference\t \tDrrB\t1P2F\t2.1\t2.3\tNo\t\t\t \tDrrD\t1KGS\t2.1\t1.9\tNo\t\t\t \tKdpE\t4KNY\t1.9\t2.6\tYes\t\t\t \tMtrA\t2GWR\t2.1\t2.0\tNo\t\t\t \tPrrA\t1YS6\t2.0\t2.2\tNo\t\t\t \tRegX3\t2OQR\t2.3\t2.1\tNo\t\t\t \tBaeR\t4B09\t2.1\t2.1\tNo\t\t\t \tVraR\t4GVP\t2.3\t2.6\tNo\t\t\t \t Based on the Dali server, the NsrR-RD domain is structurally closely related to KdpE (PDB code: 4KNY) from E. coli, displaying a sequence identity of 28%. This structural homology is also reflected by the low rmsd of 1.9 Å over 117 Cα atoms after superimposition of the receiver domains of NsrR and KdpE (Table 2). Furthermore, the orientation of the helix α4 in NsrR is close to that present in KdpE (S1 Fig). Active site residues and dimerization All RRs contain a highly conserved aspartate residue in the active site (Fig 3; shown in red). Phosphorylation of this aspartate residue induces a conformational change leading to the activation of the effector domain that binds DNA and regulates the transcription of target genes. This site of phosphorylation is conserved throughout the family of response regulators, including the lantibiotic resistance-associated RRs such as BraR from L. monocytogenes, BceR from Bacillus subtilis, CprR from C. difficile, GraR from S. aureus, LcrR from S. mutans, LisR, and VirR from L. monocytogenes (Fig 3). Sequence alignment of NsrR protein with other response regulators. A sequence alignment of NsrR with RRs belonging to the OmpR/PhoB subfamily (marked in grey) and RRs involved in lantibiotic resistance (black) is shown. The active site aspartate residue (highlighted in red), the residues forming the acidic pocket surrounding it (highlighted in pink), the switch residues (highlighted in blue), the conserved lysine residue (highlighted in green), the highly conserved residues of the linker region (colored in purple), the residues involved in dimer interface of receiver domain (highlighted in yellow), residues involved in interdomain interactions (shown in orange boxes and in cyan) and the residues involved in interaction with DNA (colored in blue) are shown. The linker region of the known structures is underlined within the sequence. The putative phosphorylation site of NsrR is Asp55, which is localized at the end of strand β3 (Fig 3, shown in red; Fig 4) and lies within an acidic environment composed of the side chains of Glu12 and Asp13 (Fig 3, highlighted in pink). This pocket is similar to the acidic active site observed within most structures of RRs such as PhoB from E. coli, PhoP from M. tuberculosis, and DivK from Caulobacter crescentus. In NsrR, Glu12, Asp13, and Asp55 are in close proximity of a highly conserved Lys104 residue (highlighted in green in Fig 3). Location of the highly conserved Asp55 and inactive state conformation of the key switch residues, Ser82 and Phe101 in NsrR-RD. NsrR (represented in yellow) displays a geometry representing the inactive state as deduced from the inactive state structure of PhoB (shown in brown, PDB code 1B00) (a). The inactive conformation of NsrR differs from the active state structure of PhoB (light blue, PDB code 1ZES) (b) in the orientation of the corresponding switch residues, Ser82 and Phe101, which adopt a conformation pointing away from the active site (Asp55 in NsrR). A divalent metal ion is usually bound in this acidic environment and is essential for phosphorylation and de-phosphorylation of RRs. In some RRs like CheY, Mg2+ is observed in the structure, bound near the phosphorylation site. In the KdpE regulator from E. coli that is involved in osmoregulation, a divalent calcium ion is present. However, the structure of NsrR-RD did not contain any divalent ion. Instead, a water molecule is present, which interacts with Glu12 of the acidic pocket, Lys104, and another water molecule in the vicinity. Within the β4-α4 loop and in β5 of the RD of RRs, specific amino acids are crucial for signal transduction from the RD to the ED via conformational changes that are a consequence of phosphorylation of the RD. These amino acids are Ser/Thr and Phe/Tyr located at the end of β4 and before β5, respectively, and designated as “signature switch residues”. As seen in the alignment (Fig 3, highlighted in blue), these signature residues (Ser/Thr and Phe/Tyr) are highly conserved in the lantibiotic resistance-associated RRs. The orientation of the side chains of these residues determines whether the RD is in an active or inactive state. In the inactive state, the phenylalanine or tyrosine residue faces away from the active site, and the corresponding serine or threonine residue adopts an outward-facing conformation as well (Fig 4A). In contrast, the switch residues face towards the active site in the active state conformation (Fig 4B). By sequence alignment with other lantibiotic resistance-associated RRs, these “signature switch residues” are identified as Ser82 and Phe101 in NsrR (see above). Although some RRs such as KdpE, BraR, BceR, GraR, and VirR contain a serine residue as the first switch residue, the others possess a threonine instead. Furthermore, the second switch residue is mostly a tyrosine, with NsrR, BraR, and BceR being the only exceptions containing a phenylalanine at that position. A comparison of the NsrR-RD structure with the available structures of PhoB (Fig 4) in the active (PDB code: 1ZES) and inactive (PDB code: 1B00) states demonstrates that Ser82 (NsrR-RD) is oriented away from the active site Asp55, and that Phe101 is also in an outward conformation suggesting an inactive state of the NsrR-RD (Fig 4A). As mentioned above, RRs contain a phosphorylation-activated switch and normally exist in equilibrium between the active and inactive conformations. Phosphorylation shifts the equilibrium towards the active conformation and induces the formation of rotationally symmetric dimers on the α4-β5-α5 interface of RDs. It has been suggested that dimerization is crucial for DNA-binding of RRs of the OmpR/PhoB subfamily. The RD domain of NsrR was crystallized with two separate monomers in the asymmetric unit. Therefore, we performed a DALI search and focused on RD domains that were structurally determined as functional dimers. In this context, the dimer of full-length KdpE from E. coli (Z-score 18.8, rmsd 1.9 Å over 117 Cα atoms) (PDB code: 4KNY) and the structure of the functional dimer of the RD of KdpE from E. coli (PDB code: 1ZH2) represent the most structurally related structures. We aligned NsrR-RD on both monomers of the RD of KdpE. Since helix α4 of NsrR-RD is orientated slightly different when compared with other structures of RDs (Fig 2), helix α4 and the N-terminal loop of one monomer were clashing with the second monomer (S2A Fig). Therefore, helix α4 and the N-terminal loop were shifted to the position of KdpE by primarily modifying backbone torsion angles in the region immediately C-terminal to helix α4. Afterwards, helix α4 and the adjacent loops were energy minimized with the MAB force field as implemented in the program Moloc; all other atoms of NsrR-RD were kept fixed. The result is highlighted in S2B Fig. The energy minimized structure of NsrR-RD was then superimposed on the dimeric structure of KdpE. The putative functional dimer of NsrR-RD is depicted in Fig 5. The dimeric interface is formed by α4-β5-α5 of RD (Fig 5A), as previously observed in other RRs. In KdpE, a network of salt bridges and other electrostatic interactions stabilize the interface within a single monomer as well as between the monomers. Majority of these interactions involve residues that are highly conserved within the OmpR/PhoB subfamily of RRs. In addition, the dimeric interface of KdpE is characterized by hydrophobic patch formed by residues Ile88 (α4), Leu91 (α4), Ala110 (α5), and Val114 (α5). Structurally, a similar set of residues is also found in NsrR: Leu94 (α4), Val110 (α5) and Ala113 (α5), respectively (depicted as spheres in Fig 5B), which are conserved to some extent on sequence level (highlighted in yellow; Fig 3). Functional dimer orientation of the RDs of NsrR. Dimeric structure of the RD of NsrR aligned to the structure of KdpE (PDB code 1ZH2, not shown). (a) The two monomers of NsrR as functional dimers are represented in a cartoon representation displayed in cyan and yellow colors. (b) Zoom-in of the dimeric interface mediated by α4-β5-α5. The monomer-monomer interactions are facilitated by hydrophobic residues (displayed as spheres), inter- and intra-domain interactions (displayed as sticks). The layout is adopted from. Conserved intermolecular electrostatic interactions further stabilize the monomer-monomer interaction of KdpE and are formed between Asp97 (β5) and Arg111 (α5), Asp96 (α4–β5 loop) and Arg118 (α5), and Asp92 (α4) and Arg113 (α5). Some of these interactions can also be identified in the dimeric model of NsrR-RD. Here, Asp100 (β5) and Lys114 (α5) form an interaction within one monomer, and an intermolecular interaction can be observed between Asn95 (α4) of one monomer with Thr116 (α5) of the other monomer (Fig 3, shown in cyan). Asp99 (α4–β5 loop; Fig 3, shown in cyan) points toward the side chain of Arg121. This interaction is also observed in KdpE (Asp96 (α4–β5 loop) and Arg118 (α5)). In KdpE, Arg111 is additionally stabilized by another intra-molecular salt bridge with Glu107 (α5). Interestingly, in NsrR-RD this amino acid corresponds to Val110 (highlighted in yellow in Fig 3). As observed in this alignment, the above-mentioned arginine residue (Arg111 in KdpE) is either an arginine or a lysine residue (Lys114 in NsrR) in all RRs used in the alignment (Fig 3, shown in cyan). Interestingly, whenever an arginine is present at this position (Arg111 in KdpE), a glutamate (Glu107 in KdpE) is present as well, presumably stabilizing the arginine side chain. However, when a lysine is present at this position, the glutamate is exchanged to a hydrophobic residue contributing to the hydrophobic patch described above. Additionally, it has been shown for PhoB from E. coli and PhoP from B. subtilis that mutating the corresponding residues involved in dimerisation (residues Asp100, Val110 and Lys114 in NsrR) results in monomeric form of response regulator which has lost the ability to dimerize as well as display reduced DNA binding capabilities. Overall Structure of C-terminal DNA-binding effector domain of NsrR The structure of NsrR-ED from S. agalactiae was determined using experimental phases from a single-wavelength anomalous dispersion dataset from the rectangular plate-shaped crystal derivatized with platinum at a resolution of 1.6 Å in space group P21212. The Rwork and Rfree values after refinement were 0.18 and 0.22, respectively. Ramachandran validation was done using MolProbity. Almost all residues (99.48%, 193 amino acids) were in the preferred or allowed regions, while 0.52% (1 amino acid) were localized in the disallowed region. The latter is Glu128 (last residue of the linker region) of chain B that is involved in crystal contacts and, therefore, likely adopts an unfavorable conformation. The structure contained a few ethylene glycol molecules introduced by the cryo-protecting procedure. The data collection and refinement statistics are listed in Table 1. The C-terminal effector DNA-binding domain of NsrR is about 13 kDa in size and consists of residues 129–243 (including 21 amino acid residues of the expression tag). Monomer A contains residue 129–224 and monomer B contain residues 128–225. For Asp147 of chain A and Glu174 of chain B, poor electron density was observed for the side chains and, thus, these side chains were removed during refinement. The asymmetric unit contains two copies of NsrR-ED related by two-fold rotational symmetry. An overlay revealed that both monomers display high similarity in their overall structure with an rmsd of 0.5 Å over 95 Cα atoms. We therefore describe for the overall structure only monomer A. The ED domain of NsrR consists of six β-strands and three α-helices in a β6-β7-β8-β9-α6-α7-α8-β10-β11 topology (the secondary structure elements are counted in continuation of those of the RD). The effector domain starts with a 4-stranded antiparallel β-sheet, followed by three α-helices and eventually ends in a C-terminal β-hairpin (Fig 6). The two β-sheets sandwich the three α-helices. Structure of the C-terminal effector domain of NsrR. Cartoon representation of the C-terminal effector domain of NsrR (green; recognition helix in cyan). The structural areas with the highest variations compared to the effector domains of DrrB (pink, 1P2F), MtrA (grey, 2GWR), and PhoB (blue, 1GXQ) are marked. The transactivation loop of MtrA is missing in the structure, therefore, the two termini are connected by a dashed line. The characteristic feature of the OmpR/PhoB subfamily of RRs is a winged helix-turn-helix (wHTH) fold that is adopted by the α7-loop-α8 segment in full-length and single effector domain structures of RRs of this subfamily. The structure of NsrR-ED also contains such a wHTH motif built up by helices α7 and α8 (Fig 6). The second helix of the wHTH motif is important for DNA-binding and, therefore, is termed “recognition helix” (shown in cyan in Fig 6). Furthermore, a helix within the HTH motif, named “positioning helix”, is important for proper orientation and positioning of the loop between these two helices and is referred to as “transactivation loop” (also called α-loop; Fig 6). In the structure of NsrR-ED, helix α8 is identified as the recognition helix, α7 as the positioning helix, and the loop region between helices α7-α8 as transactivation loop as observed in other RRs (Fig 6). The 16-residue long, solvent-exposed recognition helix α8 of NsrR-ED contains four positively charged residues that can potentially interact with DNA. These are Arg198, Arg200, Lys201, and Lys202. When comparing the sequence of NsrR with PhoB, KdpE, and MtrA, the alignment (Fig 3, colored in blue) emphasizes the variations at these positions, except for Arg200, which is conserved throughout the lantibiotic resistance RRs. Additionally, Lys202 is also highly conserved throughout the family of RRs except PhoB, clearly reflecting differences in the sequences of DNA to be bound. Comparison with structures of other effector domains We performed a DALI search to identify structurally related proteins to NsrR-ED. Here the structure of the effector domain of PhoB from E. coli (PDB code: 1GXQ) (Z-score of 13.7) is structurally the most closely related. Similar to the PhoB effector domain, a 9-residues long loop (amino acid 182–189) is also present in the structure of NsrR-ED that connects helices α7 and α8. The rmsd between the three helices of the effector domain (including the two helices forming the wHTH motif) of PhoB and NsrR-ED is 1.6 Å over 47 Cα atoms, clearly indicating that NsrR belongs to the OmpR/PhoB family of RRs. Therefore, we superimposed the Cα traces of the effector domain of NsrR (NsrR-ED) with other previously determined effector domains from the OmpR/PhoB family such as DrrB, MtrA and of only the effector domain structure of PhoB from E. coli. Overall, the structures are very similar with rmsd’s ranging from 1.7 to 2.6 Å (Table 2). The highest variations (Fig 6) are visible in in the loop regions α7-α8, which corresponds to the transactivation loop. Interestingly, this region also shows low sequence conservation (Fig 3). In many RRs this transactivation loop along with the recognition helix α8, form inter-domain contacts in the inactive state and are only exposed upon activation of the RRs via a conformational change where the N- and C-terminal domains move away from each other. Linker region The linkers that connect the RDs and EDs in response regulators are highly variable with respect to both length and sequence. The exact boundaries of these linkers are difficult to predict from sequence alignments in the absence of structural information of the distinct RR. Linker lengths in OmpR/PhoB proteins of unknown structure have been estimated by comparing the number of residues between conserved landmark residues in the regulatory and effector domains to those from structurally characterized family members. Such analysis has indicated that linker lengths vary from 5 to 20 residues. Similar to the OmpR/PhoB family, the lantibiotic resistance-associated family of response regulators also displays diverse linker regions, which are recognized in sequence alignments by the introduction of gaps (Fig 3). Interestingly, two arginine residues (Arg120 and Arg121 in NsrR; Fig 3, shown in purple) at the end of the RDs seem to be strictly conserved throughout the family of response regulators in both the OmpR/PhoB and lantibiotic resistance-associated RRs, indicating a conserved similarity. As seen in the structures of MtrA and KdpE, this arginine residue residing at the end of α5 participates in the active state dimer interface of the RD through a salt bridge interaction with an aspartate residue. This aspartate residue is identified in NsrR as Asp99 (see above). Arginine 121 of NsrR points towards this Asp99 residue however, the distance for a salt bridge interaction is too large. Although we aimed at crystallizing full-length NsrR, this endeavor failed due to proteolytic cleavage within the linker region during the time period of crystallization. Nonetheless, the structures of NsrR-RD and NsrR-ED together provide the required structural knowledge to predict the linker region that joins the receiver and effector domains. The linker region of NsrR consists of approximately nine residues (Fig 3), comprising 120RRSQQFIQQ128 (underlined residues are neither present in the structure of RD nor in ED of NsrR) and contains two positively charged amino acids. DNA-binding mode of NsrR using a full-length model Since the structures of both domains of NsrR were determined, we used this structural information together with the available crystal structures of related proteins to create a model of the full-length NsrR in its active and inactive state. To achieve this, we first carefully analyzed the outcome of the Dali search for each domain and identified structurally highly similar proteins (based on Z-scores and rmsd values) and choose the full-length structures previously reported. This resulted in a list of possible templates for modeling the full-length structure of NsrR (Table 2). In solution, RRs exist in equilibrium between the active and inactive state, which is shifted towards the active state upon phosphorylation of the ED. This results in oligomerization of the RR and a higher affinity towards DNA. Based on the above-mentioned criteria, the structure of MtrA from M. tuberculosis, crystallized in an inactive and non-phosphorylated state, seemed best suited for modeling purposes. Furthermore, the linker between the two domains of MtrA contains nine amino acids and is of similar length as the linker of NsrR. We aligned the NsrR-RD and -ED to the corresponding MtrA domains and evaluated the structure. This mimics the closed inactive conformation of NsrR (Fig 7A; the missing linker is represented as dotted line). Model of full-length NsrR in its inactive state and active state. The RD domain of NsrR is highlighted in yellow and the ED domain in green with the “recognition helix” colored in cyan. (a) Inactive state conformation: Both domains of NsrR were aligned to the structure of MtrA (not shown), which adopts a closed inactive conformation, to obtain a model of full-length NsrR. Phe101 and Asp187 stabilize this closed conformation. The missing linker is represented by a dotted line. (b) Active state conformation: A model of full-length NsrR in active conformation based on the alignment of both the domains of NsrR to the structure of DNA bound structure of KdpE (PDB code: 4KNY), adopting an active open conformation, where the other molecule of NsrR is shown in shades of blue with the recognition helix colored in green. In MtrA, the two domains interact via the α4-β5-α5 interface of the receiver domain and the end of α7, α7-α8 loop and α8 of the effector domain. Both interfaces have been shown to form functionally important contact areas in the active state within members of the OmpR/PhoB subfamily. In our model of full-length NsrR, a similar orientation between the domains is observed, contributing to the inter-domain interactions. The inactive conformation of MtrA is supported by the orientation of the side chain of Tyr102, which points away from the active Asp56 residue, while the side chain of Tyr102 interacts with Asp190 of the RD of MtrA, thereby stabilizing its closed conformation. In the model of NsrR, similar amino acids are present, Phe101 (switch residue) and Asp188 (Fig 3, represented by orange boxes) forming a likewise similar network of interaction. Next, we were interested in the active conformation of the NsrR protein adopting an active “open” conformation in the dimeric state. We compared and aligned the NsrR-RD and ED on the dimeric structure of KdpE that was solved in the DNA-bound state (Fig 7B). Also the linker region of KdpE is of similar length as of NsrR, which suggests that the distance in the DNA-bound state between the RD and ED of NsrR will be similar to that in the KdpE active dimer. We superimposed the ED of NsrR with the DNA-binding domain of KdpE resulting in a reasonably well-aligned structure (rmsd of 2.6Å over 86 Cα atoms; Table 2). As a result, a highly positive groove is created by the two ED domains of NsrR which likely represents the DNA binding site as observed in KdpE. A prediction of the putative promoter sequence that NsrR binds via the BPROM online server was performed (S3 Fig). A promoter region was identified upstream of the nsr operon. However, the regulation of the predicted promoter and the DNA binding by NsrR has to be confirmed. Conclusion In numerous pathogenic bacteria such as S. agalactiae, S. aureus, and C. difficile that apparently do not produce a lantibiotic, a gene cluster is present to provide resistance against lantibiotics such as nisin, nukacin ISK-1, lacticin 481 gallidermin, actagardine, or mersacidin. The regulation of the expression of these genes is mediated by two-component systems. The structure of the response regulator NsrR from S. agalactiae presented in this study is the first structural information available for the subgroup of lantibiotic resistance-associated RRs. Supporting Information References Bacteriocins—a viable alternative to antibiotics? Engineering of a novel thioether bridge and role of modified residues in the lantibiotic Pep5 Lantibiotics: biosynthesis and biological activities of uniquely modified peptides from gram-positive bacteria Bacterial lantibiotics: strategies to improve therapeutic potential Lantibiotics: promising candidates for future applications in health care The therapeutic potential of lantibiotics The C‐terminus of nisin is important for the ABC transporter NisFEG to confer immunity in Lactococcus lactis Specific Binding of Nisin to the Peptidoglycan Precursor Lipid II Combines Pore Formation and Inhibition of Cell Wall Biosynthesis for Potent Antibiotic Activity Regulation of nisin biosynthesis and immunity in Lactococcus lactis 6F3 Characterization of the nisin gene cluster nisABTCIPR of Lactococcus lactis Role of the single regulator MrsR1 and the two-component system MrsR2/K2 in the regulation of mersacidin production and immunity Lantibiotics: how do producers become self-protected? Biosynthesis and mode of action of lantibiotics Immunity to lantibiotics NMR resonance assignments for the class II GTP binding RNA aptamer in complex with GTP The First structure of a lantibiotic immunity protein, SpaI from Bacillus subtilis, reveals a novel fold Distinct mechanisms contribute to immunity in the lantibiotic NAI-107 producer strain Microbispora ATCC PTA-5024 NSR from Streptococcus agalactiae confers resistance against nisin and is encoded by a conserved nsr operon Involvement of the Novel Two-Component NsrRS and LcrRS Systems in Distinct Resistance Pathways against Nisin A and Nukacin ISK-1 in Streptococcus mutans GraXSR proteins interact with the VraFG ABC transporter to form a five-component system required for cationic antimicrobial peptide sensing and resistance in Staphylococcus aureus Structural basis of lantibiotic recognition by the nisin resistance protein from Streptococcus agalactiae Lantibiotic Resistance Quorum sensing by peptide pheromones and two-component signal-transduction systems in Gram-positive bacteria Two-component signal transduction Two-component signal transduction as a target for microbial anti-infective therapy Bacitracin and nisin resistance in Staphylococcus aureus: a novel pathway involving the BraS/BraR two-component system (SA2417/SA2418) and both the BraD/BraE and VraD/VraE ABC transporters The BceRS two-component regulatory system induces expression of the bacitracin transporter, BceAB, in Bacillus subtilis Identification and disruption of lisRK, a genetic locus encoding a two-component signal transduction system involved in stress tolerance and virulence in Listeria monocytogenes Coevolution of ABC transporters and two-component regulatory systems as resistance modules against antimicrobial peptides in Firmicutes bacteria Intramembrane-sensing histidine kinases: a new family of cell envelope stress sensors in Firmicutes bacteria Bacitracin sensing in Bacillus subtilis Peptide antibiotic sensing and detoxification modules of Bacillus subtilis The BceABRS four-component system regulates the bacitracin-induced cell envelope stress response in Streptococcus mutans Bacterial response regulators: versatile regulatory strategies from common domains Structural relationships in the OmpR family of winged-helix transcription factors The structure of a full-length response regulator from Mycobacterium tuberculosis in a stabilized three-dimensional domain-swapped, activated state Domain orientation in the inactive response regulator Mycobacterium tuberculosis MtrA provides a barrier to activation The structural basis of signal transduction for the response regulator PrrA from Mycobacterium tuberculosis Structure of the response regulator PhoP from Mycobacterium tuberculosis reveals a dimer through the receiver domain Structural analysis of the domain interface in DrrB, a response regulator of the OmpR/PhoB subfamily Evidence of intradomain and interdomain flexibility in an OmpR/PhoB homolog from Thermotoga maritima An asymmetric heterodomain interface stabilizes a response regulator–DNA complex Overexpression, purification and crystallization of the response regulator NsrR involved in nisin resistance Fast and sensitive colloidal coomassie G-250 staining for proteins in polyacrylamide gels The ID23-1 structural biology beamline at the ESRF Xds Three-dimensional crystal structure of the transcription factor PhoB receiver domain PHENIX: a comprehensive Python-based system for macromolecular structure solution Features and development of Coot MolProbity: all-atom structure validation for macromolecular crystallography On the combination of molecular replacement and single-wavelength anomalous diffraction phasing for automated structure determination Delano WL (2002) The PyMOL molecular graphics system. Response regulator YycF essential for bacterial growth: X-ray crystal structure of the DNA-binding domain and its PhoB-like DNA recognition motif Tandem DNA Recognition by PhoB, a Two-Component Signal Transduction Transcriptional Activator Structural analysis and solution studies of the activated regulatory domain of the response regulator ArcA: a symmetric dimer mediated by the α4-β5-α5 face Mechanism of activation for transcription factor PhoB suggested by different modes of dimerization in the inactive and active states The dimeric form of the unphosphorylated response regulator BaeR Dali server: conservation mapping in 3D Basic local alignment search tool The Clostridium difficile cpr locus is regulated by a noncontiguous two-component system in response to type A and B lantibiotics Identification of a genetic locus responsible for antimicrobial peptide resistance in Clostridium difficile Mutated response regulator graR is responsible for phenotypic conversion of Staphylococcus aureus from heterogeneous vancomycin-intermediate resistance to vancomycin-intermediate resistance Interaction of the GraRS two-component system with the VraFG ABC transporter to support vancomycin-intermediate resistance in Staphylococcus aureus VirR, a response regulator critical for Listeria monocytogenes virulence Crystallographic and biochemical studies of DivK reveal novel features of an essential response regulator in Caulobacter crescentus Response regulation in bacterial chemotaxis Acetyl phosphate and the activation of two-component response regulators Structure of the magnesium-bound form of CheY and mechanism of phosphoryl transfer in bacterial chemotaxis The three-dimensional structure of two mutants of the signal transduction protein CheY suggest its molecular activation mechanism Histidine kinases and response regulator proteins in two-component signaling systems Crystal Structures of CheY Mutants Y106W and T87I/Y106W CheY Activation Correlates with Movement of Residue 106 Proposed signal transduction role for conserved CheY residue Thr87, a member of the response regulator active-site quintet Functional Dynamics of Response Regulators Using NMR Relaxation Techniques A common dimerization interface in bacterial response regulators KdpE and TorR MAB, a generally applicable molecular force field for structure modelling in medicinal chemistry Probing the roles of the two different dimers mediated by the receiver domain of the response regulator PhoB Residue R113 is essential for PhoP dimerization and function: a residue buried in the asymmetric PhoP dimer interface determined in the PhoPN three-dimensional crystal structure The DNA-binding domain of OmpR: crystal structures of a winged helix transcription factor Advances and pitfalls of protein structural alignment Automatic annotation of microbial genomes and metagenomic sequences Preliminary crystallographic studies of the regulatory domain of response regulator YycF from an essential two-component signal transduction system. Acta Crystallographica Section F Structure of the DNA-binding domain of the response regulator PhoP from Mycobacterium tuberculosis Water-mediated interactions between DNA and PhoB DNA-binding/transactivation domain: NMR-restrained molecular dynamics in explicit water environment Structural dynamics of the two-component response regulator RstA in recognition of promoter DNA element Phosphorylation-dependent conformational changes and domain rearrangements in Staphylococcus aureus VraR 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diff --git a/annotated_BioC_JSON/PMC4774019_ann.json b/annotated_BioC_JSON/PMC4774019_ann.json
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index 0000000000000000000000000000000000000000..aaeedbb44d270da89a2e3031d365a57a28fd0011
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+++ b/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 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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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+[{"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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+[{"sourceid":"4784909","sourcedb":"","project":"","target":"","text":"The Structural Basis of Coenzyme A Recycling in a Bacterial Organelle Bacterial Microcompartments (BMCs) are proteinaceous organelles that encapsulate critical segments of autotrophic and heterotrophic metabolic pathways; they are functionally diverse and are found across 23 different phyla. The majority of catabolic BMCs (metabolosomes) compartmentalize a common core of enzymes to metabolize compounds via a toxic and/or volatile aldehyde intermediate. The core enzyme phosphotransacylase (PTAC) recycles Coenzyme A and generates an acyl phosphate that can serve as an energy source. The PTAC predominantly associated with metabolosomes (PduL) has no sequence homology to the PTAC ubiquitous among fermentative bacteria (Pta). Here, we report two high-resolution PduL crystal structures with bound substrates. The PduL fold is unrelated to that of Pta; it contains a dimetal active site involved in a catalytic mechanism distinct from that of the housekeeping PTAC. Accordingly, PduL and Pta exemplify functional, but not structural, convergent evolution. The PduL structure, in the context of the catalytic core, completes our understanding of the structural basis of cofactor recycling in the metabolosome lumen. This study describes the structure of a novel phosphotransacylase enzyme that facilitates the recycling of the essential cofactor acetyl-CoA within a bacterial organelle and discusses the properties of the enzyme's active site and how it is packaged into the organelle. Author Summary In metabolism, molecules with “high-energy” bonds (e.g., ATP and Acetyl~CoA) are critical for both catabolic and anabolic processes. Accordingly, the retention of these bonds during biochemical transformations is incredibly important. The phosphotransacylase (Pta) enzyme catalyzes the conversion between acyl-CoA and acyl-phosphate. This reaction directly links an acyl-CoA with ATP generation via substrate-level phosphorylation, producing short-chain fatty acids (e.g., acetate), and also provides a path for short-chain fatty acids to enter central metabolism. Due to this key function, Pta is conserved across the bacterial kingdom. Recently, a new type of phosphotransacylase was described that shares no evolutionary relation to Pta. This enzyme, PduL, is exclusively associated with organelles called bacterial microcompartments, which are used to catabolize various compounds. Not only does PduL facilitate substrate level phosphorylation, but it also is critical for cofactor recycling within, and product efflux from, the organelle. We solved the structure of this convergent phosphotransacylase and show that it is completely structurally different from Pta, including its active site architecture. We also discuss features of the protein important to its packaging in the organelle. Introduction Bacterial Microcompartments (BMCs) are organelles that encapsulate enzymes for sequential biochemical reactions within a protein shell. The shell is typically composed of three types of protein subunits, which form either hexagonal (BMC-H and BMC-T) or pentagonal (BMC-P) tiles that assemble into a polyhedral shell. The facets of the shell are composed primarily of hexamers that are typically perforated by pores lined with highly conserved, polar residues that presumably function as the conduits for metabolites into and out of the shell. The vitamin B12-dependent propanediol-utilizing (PDU) BMC was one of the first functionally characterized catabolic BMCs; subsequently, other types have been implicated in the degradation of ethanolamine, choline, fucose, rhamnose, and ethanol, all of which produce different aldehyde intermediates (Table 1). More recently, bioinformatic studies have demonstrated the widespread distribution of BMCs among diverse bacterial phyla and grouped them into 23 different functional types. The reactions carried out in the majority of catabolic BMCs (also known as metabolosomes) fit a generalized biochemical paradigm for the oxidation of aldehydes (Fig 1). This involves a BMC-encapsulated signature enzyme that generates a toxic and/or volatile aldehyde that the BMC shell sequesters from the cytosol. The aldehyde is subsequently converted into an acyl-CoA by aldehyde dehydrogenase, which uses NAD+ and CoA as cofactors. These two cofactors are relatively large, and their diffusion across the protein shell is thought to be restricted, necessitating their regeneration within the BMC lumen. NAD+ is recycled via alcohol dehydrogenase, and CoA is recycled via phosphotransacetylase (PTAC) (Fig 1). The final product of the BMC, an acyl-phosphate, can then be used to generate ATP via acyl kinase, or revert back to acyl-CoA by Pta for biosynthesis. Collectively, the aldehyde and alcohol dehydrogenases, as well as the PTAC, constitute the common metabolosome core. General biochemical model of aldehyde-degrading BMCs (metabolosomes) illustrating the common metabolosome core enzymes and reactions. Substrates and cofactors involving the PTAC reaction are shown in red; other substrates and enzymes are shown in black, and other cofactors are shown in gray. Characterized and predicted catabolic BMC (metabolosome) types that represent the aldehyde-degrading paradigm (for definition of types see Kerfeld and Erbilgin). Name\tPTAC Type\tSequestered Aldehyde\t \tPDU*\tPduL\tpropionaldehyde\t \tEUT1\tPTA_PTB\tacetaldehyde\t \tEUT2\tPduL\tacetaldehyde\t \tETU\tNone\tacetaldehyde\t \tGRM1/CUT\tPduL\tacetaldehyde\t \tGRM2\tPduL\tacetaldehyde\t \tGRM3*,4\tPduL\tpropionaldehyde\t \tGRM5/GRP\tPduL\tpropionaldehyde\t \tPVM*\tPduL\tlactaldehyde\t \tRMM1,2\tNone\tunknown\t \tSPU\tPduL\tunknown\t \t * PduL from these functional types of metabolosomes were purified in this study. The activities of core enzymes are not confined to BMC-associated functions: aldehyde and alcohol dehydrogenases are utilized in diverse metabolic reactions, and PTAC catalyzes a key biochemical reaction in the process of obtaining energy during fermentation. The concerted functioning of a PTAC and an acetate kinase (Ack) is crucial for ATP generation in the fermentation of pyruvate to acetate (see Reactions 1 and 2). Both enzymes are, however, not restricted to fermentative organisms. They can also work in the reverse direction to activate acetate to the CoA-thioester. This occurs, for example, during acetoclastic methanogenesis in the archaeal Methanosarcina species.  Reaction 1: acetyl-S-CoA + Pi ←→ acetyl phosphate + CoA-SH (PTAC)  Reaction 2: acetyl phosphate + ADP ←→ acetate + ATP (Ack) The canonical PTAC, Pta, is an ancient enzyme found in some eukaryotes and archaea, and widespread among the bacteria; 90% of the bacterial genomes in the Integrated Microbial Genomes database contain a gene encoding the PTA_PTB phosphotransacylase (Pfam domain PF01515). Pta has been extensively characterized due to its key role in fermentation. More recently, a second type of PTAC without any sequence homology to Pta was identified. This protein, PduL (Pfam domain PF06130), was shown to catalyze the conversion of propionyl-CoA to propionyl-phosphate and is associated with a BMC involved in propanediol utilization, the PDU BMC. Both pduL and pta genes can be found in genetic loci of functionally distinct BMCs, although the PduL type is much more prevalent, being found in all but one type of metabolosome locus: EUT1 (Table 1). Furthermore, in the Integrated Microbial Genomes Database, 91% of genomes that encode PF06130 also encode genes for shell proteins. As a member of the core biochemical machinery of functionally diverse aldehyde-oxidizing metabolosomes, PduL must have a certain level of substrate plasticity (see Table 1) that is not required of Pta, which has generally been observed to prefer acetyl-CoA. PduL from the PDU BMC of Salmonella enterica favors propionyl-CoA over acetyl-CoA, and it is likely that PduL orthologs in functionally diverse BMCs would have substrate preferences for other CoA derivatives. Another distinctive feature of BMC-associated PduL homologs is an N-terminal encapsulation peptide (EP) that is thought to “target” proteins for encapsulation by the BMC shell. EPs are frequently found on BMC-associated proteins and have been shown to interact with shell proteins. EPs have also been observed to cause proteins to aggregate, and this has recently been suggested to be functionally relevant as an initial step in metabolosome assembly, in which a multifunctional protein core is formed, around which the shell assembles. Of the three common metabolosome core enzymes, crystal structures are available for both the alcohol and aldehyde dehydrogenases. In contrast, the structure of PduL, the PTAC found in the vast majority of catabolic BMCs, has not been determined. This is a major gap in our understanding of metabolosome-encapsulated biochemistry and cofactor recycling. Structural information will be essential to working out how the core enzymes and their cofactors assemble and organize within the organelle lumen to enhance catalysis. Moreover, it will be useful for guiding efforts to engineer novel BMC cores for biotechnological applications. The primary structure of PduL homologs is subdivided into two PF06130 domains, each roughly 80 residues in length. No available protein structures contain the PF06130 domain, and homology searches using the primary structure of PduL do not return any significant results that would allow prediction of the structure. Moreover, the evident novelty of PduL makes its structure interesting in the context of convergent evolution of PTAC function; to-date, only the Pta active site and catalytic mechanism is known. Here we report high-resolution crystal structures of a PduL-type PTAC in both CoA- and phosphate-bound forms, completing our understanding of the structural basis of catalysis by the metabolosome common core enzymes. We propose a catalytic mechanism analogous but yet distinct from the ubiquitous Pta enzyme, highlighting the functional convergence of two enzymes with completely different structures and metal requirements. We also investigate the quaternary structures of three different PduL homologs and situate our findings in the context of organelle biogenesis in functionally diverse BMCs. Results Structure Determination of PduL We cloned, expressed, and purified three different PduL homologs from functionally distinct BMCs (Table 1): from the well-studied pdu locus in S. enterica Typhimurium LT2 (sPduL), from the recently characterized pvm locus in Planctomyces limnophilus (pPduL), and from the grm3 locus in Rhodopseudomonas palustris BisB18 (rPduL). While purifying full-length sPduL, we observed a tendency to aggregation as described previously, with a large fraction of the expressed protein found in the insoluble fraction in a white, cake-like pellet. Remarkably, after removing the N-terminal putative EP (27 amino acids), most of the sPduLΔEP protein was in the soluble fraction upon cell lysis. Similar differences in solubility were observed for pPduL and rPduL when comparing EP-truncated forms to the full-length protein, but none were quite as dramatic as for sPduL. We confirmed that all homologs were active (S1a and S1b Fig). Among these, we were only able to obtain diffraction-quality crystals of rPduL after removing the N-terminal putative EP (33 amino acids, also see Fig 2a) (rPduLΔEP). Truncated rPduLΔEP had comparable enzymatic activity to the full-length enzyme (S1a Fig). Structural overview of R. palustris PduL from the grm3 locus. (a) Primary and secondary structure of rPduL (tubes represent α-helices, arrows β-sheets and dashed line residues disordered in the structure. Blocks of ten residues are shaded alternatively black/dark gray. The first 33 amino acids are present only in the wildtype construct and contains the predicted EP alpha helix, α0); the truncated rPduLΔEP that was crystallized begins with M-G-V. Coloring is according to structural domains (domain 1 D36-N46/Q155-C224, blue; loop insertion G61-E81, grey; domain 2 R47-F60/E82-A154, red). Metal coordination residues are highlighted in light blue and CoA contacting residues in magenta, residues contacting the CoA of the other chain are also outlined. (b) Cartoon representation of the structure colored by domains and including secondary structure numbering. The N-and C-termini are in close proximity. Coenzyme A is shown in magenta sticks and Zinc (grey) as spheres. We collected a native dataset from rPduLΔEP crystals diffracting to a resolution of 1.54 Å (Table 2). Using a mercury-derivative crystal form diffracting to 1.99 Å (Table 2), we obtained high quality electron density for model building and used the initial model to refine against the native data to Rwork/Rfree values of 18.9/22.1%. There are two PduL molecules in the asymmetric unit of the P212121 unit cell. We were able to fit all of the primary structure of PduLΔEP into the electron density with the exception of three amino acids at the N-terminus and two amino acids at the C-terminus (Fig 2a); the model is of excellent quality (Table 2). A CoA cofactor as well as two metal ions are clearly resolved in the density (for omit maps of CoA see S2 Fig). Data collection and refinement statistics \tPduL native\tPduL mercury derivative\tPduL phosphate soaked\t \tData collection\t\t\t\t \tSpace group\tP 21 21 21\tP 21 21 21\tP 21 21 21\t \tCell dimensions\t\t\t\t \ta, b, c (Å)\t57.7, 56.4, 150.4\t55.6, 57.7, 150.2\t57.1, 58.8, 136.7\t \tα, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t \tResolution (Å)\t31.4 − 1.54 (1.60 − 1.54)*\t35.3 − 1.99 (2.07 − 1.99)\t39.2 − 2.10 (2.21 − 2.10)\t \tRmerge\t0.169 (1.223)\t0.084 (0.299)\t0.122 (0.856)\t \tI/σ(I)\t12.9 (1.7)\t22.1 (7.1)\t12.6 (2.0)\t \tCompleteness (%)\t99.4 (94.4)\t99.3 (93.3)\t100 (99.9)\t \tRedundancy\t13.9 (12.1)\t7.2 (7.0)\t6.5 (6.1)\t \tRefinement\t\t\t\t \tResolution (Å)\t31.4 − 1.54 (1.60 − 1.54)*\t\t39.2 − 2.10 (2.18 − 2.1)\t \tNo. reflections\t72,698\t\t27,554\t \tRwork/Rfree (%)\t18.9 (30.7) / 22.1 (34.7)\t\t17.5 (24.2) / 22.6 (30.0)\t \tNo. atoms\t3,453\t\t3,127\t \tProtein\t2,841\t\t2,838\t \tLigand/ion\t100\t\t24\t \tWater\t512\t\t265\t \tB-factors\t22.8\t\t34.7\t \tProtein\t21.5\t\t24.3\t \tLigand/ion\t21.9\t\t40.6\t \tWater\t30.3\t\t37.9\t \tR.m.s deviations\t\t\t\t \tBond lengths (Å)\t0.006\t\t0.013\t \tBond angles (°)\t1.26\t\t1.30\t \tRamachandran Plot\t\t\t\t \tfavored (%)\t99\t\t99\t \tallowed (%)\t1\t\t1\t \tdisallowed (%)\t0\t\t0\t \t *Highest resolution shell is shown in parentheses. Structurally, PduL consists of two domains (Fig 2, blue/red), each a beta-barrel that is capped on both ends by short α-helices. β-Barrel 1 consists of the N-terminal β strand and β strands from the C-terminal half of the polypeptide chain (β1, β10-β14; residues 37–46 and 155–224). β-Barrel 2 consists mainly of the central segment of primary structure (β2, β5–β9; residues 47–60 and 82–154) (Fig 2, red), but is interrupted by a short two-strand beta sheet (β3-β4, residues 61–81). This β-sheet is involved in contacts between the two domains and forms a lid over the active site. Residues in this region (Gln42, Pro43, Gly44), covering the active site, are strongly conserved (Fig 3). This structural arrangement is completely different from the functionally related Pta, which is composed of two domains, each consisting of a central flat beta sheet with alpha-helices on the top and bottom. Primary structure conservation of the PduL protein family. Sequence logo calculated from the multiple sequence alignment of PduL homologs (see Materials and Methods), but not including putative EP sequences. Residues 100% conserved across all PduL homologs in our dataset are noted with an asterisk, and residues conserved in over 90% of sequences are noted with a colon. The sequences aligning to the PF06130 domain (determined by BLAST) are highlighted in red and blue. The position numbers shown correspond to the residue numbering of rPduL; note that some positions in the logo represent gaps in the rPduL sequence. There are two PduL molecules in the asymmetric unit forming a butterfly-shaped dimer (Fig 4c). Consistent with this, results from size exclusion chromatography of rPduLΔEP suggest that it is a dimer in solution (Fig 5e). The interface between the two chains buries 882 Å2 per monomer and is mainly formed by α-helices 2 and 4 and parts of β-sheets 12 and 14, as well as a π–π stacking of the adenine moiety of CoA with Phe116 of the adjacent chain (Fig 4c). The folds of the two chains in the asymmetric unit are very similar, superimposing with a rmsd of 0.16 Å over 2,306 aligned atom pairs. The peripheral helices and the short antiparallel β3–4 sheet mediate most of the crystal contacts. Details of active site, dimeric assembly, and sequence conservation of PduL. (a,b) Proposed active site of PduL with relevant residues shown as sticks in atom coloring (nitrogen blue, oxygen red, sulfur yellow), zinc as grey colored spheres and coordinating ordered water molecules in red. Distances between atom centers are indicated in Å. (a) Coenzyme A containing, (b) phosphate-bound structure. (c) View of the dimer in the asymmetric unit from the side, domains 1 and 2 colored as in Fig 2 and the two chains differentiated by blue/red versus slate/firebrick. The bottom panel shows a top view down the 2-fold axis as indicated by the arrow in the top panel. The asterisk and double arrow marks the location of the π–π interaction between F116 and the CoA base of the other dimer chain. (d) Surface representation of the structure with indicated conservation (red: high, white: intermediate, yellow: low). Size exclusion chromatography of PduL homologs. (a)–(c): Chromatograms of sPduL (a), rPduL (b), and pPduL (c) with (orange) or without (blue) the predicted EP, post-nickel affinity purification, applied over a preparative size exclusion column (see Materials and Methods). (d)–(f): Chromatograms of sPduL (d), rPduL (e), and pPduL (f) post-preparative size exclusion chromatography with different size fractions separated, applied over an analytical size exclusion column (see Materials and Methods). All chromatograms are cropped to show only the linear range of separation based on standard runs, shown in black squares with a dashed linear trend line. All y-axes are arbitrary absorbance units except the right-hand axes for panels (a) and (d), which is the log10(molecular weight) of the standards. Active Site Properties CoA and the metal ions bind between the two domains, presumably in the active site (Figs 2b and 4a). To identify the bound metals, we performed an X-ray fluorescence scan on the crystals at various wavelengths (corresponding to the K-edges of Mn, Fe, Co, Ni, Cu, and Zn). There was a large signal at the zinc edge, and we tested for the presence of zinc by collecting full data sets before and after the Zn K-edge (1.2861 and 1.2822 Å, respectively). The large differences between the anomalous signals confirm the presence of zinc at both metal sites (S3 Fig). The first zinc ion (Zn1) is in a tetrahedral coordination state with His48, His50, Glu109, and the CoA sulfur (Fig 4a). The second (Zn2) is in octahedral coordination by three conserved histidine residues (His157, His159 and His204) as well as three water molecules (Fig 4a). The nitrogen atom coordinating the zinc is the Nε in each histidine residue, as is typical for this interaction. When the crystals were soaked in a sodium phosphate solution for 2 d prior to data collection, the CoA dissociates, and density for a phosphate molecule is visible at the active site (Table 2, Fig 4b). The phosphate-bound structure aligns well with the CoA-bound structure (0.43 Å rmsd over 2,361 atoms for the monomer, 0.83 Å over 5,259 aligned atoms for the dimer). The phosphate contacts both zinc atoms (Fig 4b) and replaces the coordination by CoA at Zn1; the coordination for Zn2 changes from octahedral with three bound waters to tetrahedral with a phosphate ion as one of the ligands (Fig 4b). Conserved Arg103 seems to be involved in maintaining the phosphate in that position. The two zinc atoms are slightly closer together in the phosphate-bound form (5.8 Å vs 6.3 Å), possibly due to the bridging effect of the phosphate. An additional phosphate molecule is bound at a crystal contact interface, perhaps accounting for the 14 Å shorter c-axis in the phosphate-bound crystal form (Table 2). Oligomeric States of PduL Orthologs Are Influenced by the EP Interestingly, some of the residues important for dimerization of rPduL, particularly Phe116, are poorly conserved across PduL homologs associated with functionally diverse BMCs (Figs 4c and 3), suggesting that they may have alternative oligomeric states. We tested this hypothesis by performing size exclusion chromatography on both full-length and truncated variants (lacking the EP, ΔEP) of sPduL, rPduL, and pPduL. These three homologs are found in functionally distinct BMCs (Table 1). Therefore, they are packaged with different signature enzymes and different ancillary proteins. It has been proposed that the catabolic BMCs may assemble in a core-first manner, with the luminal enzymes (signature enzyme, aldehyde, and alcohol dehydrogenases and the BMC PTAC) forming an initial bolus, or prometabolosome, around which a shell assembles. Given the diversity of signature enzymes (Table 1), it is plausible that PduL orthologs may adopt different oligomeric states that reflect the differences in the proteins being packaged with them in the organelle lumen. We found that not only did the different orthologs appear to assemble into different oligomeric states, but that quaternary structure was dependent on whether or not the EP was present. Full-length sPduL was unstable in solution—precipitating over time—and eluted throughout the entire volume of a size exclusion column, indicating it was nonspecifically aggregating. However, when the putative EP (residues 1–27) was removed (sPduL ΔEP), the truncated protein was stable and eluted as a single peak (Fig 5a) consistent with the size of a monomer (Fig 5d, blue curve). In contrast, both full-length rPduL and pPduL appeared to exist in two distinct oligomeric states (Fig 5b and 5c respectively, orange curves), one form of the approximate size of a dimer and the second, a higher molecular weight oligomer (~150 kDa). Upon deletion of the putative EP (residues 1–47 for rPduL, and 1–20 for pPduL), there was a distinct change in the elution profiles (Fig 5b and 5c respectively, blue curves). pPduLΔEP eluted as two smaller forms, possibly corresponding to a trimer and a monomer. In contrast, rPduLΔEP eluted as one smaller oligomer, possibly a dimer. We also analyzed purified rPduL and rPduLΔEP by size exclusion chromatography coupled with multiangle light scattering (SEC-MALS) for a complementary approach to assessing oligomeric state. SEC-MALS analysis of rPdulΔEP is consistent with a dimer (as observed in the crystal structure) with a weighted average (Mw) and number average (Mn) of the molar mass of 58.4 kDa +/− 11.2% and 58.8 kDa +/− 10.9%, respectively (S4a Fig). rPduL full length runs as Mw = 140.3 kDa +/− 1.2% and Mn = 140.5 kDa +/− 1.2%. This corresponds to an oligomeric state of six subunits (calculated molecular weight of 144 kDa). Collectively, these data strongly suggest that the N-terminal EP of PduL plays a role in defining the quaternary structure of the protein. Discussion The hallmark attribute of an organelle is that it serves as a discrete subcellular compartment functioning as an isolated microenvironment distinct from the cytosol. In order to create and preserve this microenvironment, the defining barrier (i.e., lipid bilayer membrane or microcompartment shell) must be selectively permeable. The BMC shell not only sequesters specific enzymes but also their cofactors, thereby establishing a private cofactor pool dedicated to the encapsulated reactions. In catabolic BMCs, CoA and NAD+ must be continually recycled within the organelle (Fig 1). Homologs of the predominant cofactor utilizer (aldehyde dehydrogenase) and NAD+ regenerator (alcohol dehydrogenase) have been structurally characterized, but until now structural information was lacking for PduL, which recycles CoA in the organelle lumen. Curiously, while the housekeeping Pta could provide this function, and indeed does so in the case of one type of ethanolamine-utilizing (EUT) BMC, the evolutionarily unrelated PduL fulfills this function for the majority of metabolosomes using a novel structure and active site for convergent evolution of function. The Tertiary Structure of PduL Is Formed by Discontinuous Segments of Primary Structure The structure of PduL consists of two β-barrel domains capped by short alpha helical segments (Fig 2b). The two domains are structurally very similar (superimposing with a rmsd of 1.34 Å (over 123 out of 320/348 aligned backbone atoms, S5a Fig). However, the amino acid sequences of the two domains are only 16% identical (mainly the RHxH motif, β2 and β10), and 34% similar. Our structure reveals that the two assigned PF06130 domains (Fig 3) do not form structurally discrete units; this reduces the apparent sequence conservation at the level of primary structure. One strand of the domain 1 beta barrel (shown in blue in Fig 2) is contributed by the N-terminus, while the rest of the domain is formed by the residues from the C-terminal half of the protein. When aligned by structure, the β1 strand of the first domain (Fig 2a and 2b, blue) corresponds to the final strand of the second domain (β9), effectively making the domains continuous if the first strand was transplanted to the C-terminus. Refined domain assignment based on our structure should be able to predict domains of PF06130 homologs much more accurately. The closest structural homolog of the PduL barrel domain is a subdomain of a multienzyme complex, the alpha subunit of ethylbenzene dehydrogenase (S5b Fig, rmsd of 2.26 Å over 226 aligned atoms consisting of one beta barrel and one capping helix). In contrast to PduL, there is only one barrel present in ethylbenzene dehydrogenase, and there is no comparable active site arrangement. The PduL signature primary structure, two PF06130 domains, occurs in some multidomain proteins, most of them annotated as Acks, suggesting that PduL may also replace Pta in variants of the phosphotransacetylase-Ack pathway. These PduL homologs lack EPs, and their fusion to Ack may have evolved as a way to facilitate substrate channeling between the two enzymes. Implications for Metabolosome Core Assembly For BMC-encapsulated proteins to properly function together, they must be targeted to the lumen and assemble into an organization that facilitates substrate/product channeling among the different catalytic sites of the signature and core enzymes. The N-terminal extension on PduL homologs may serve both of these functions. The extension shares many features with previously characterized EPs: it is present only in homologs associated with BMC loci, and it is predicted to form an amphipathic α-helix. Moreover, its removal affects the oligomeric state of the protein. EP-mediated oligomerization has been observed for the signature and core BMC enzymes; for example, full-length propanediol dehydratase and ethanolamine ammonia-lyase (signature enzymes for PDU and EUT BMCs) subunits are also insoluble, but become soluble upon removal of the predicted EP. sPduL has also previously been reported to localize to inclusion bodies when overexpressed; we show here that this is dependent on the presence of the EP. This propensity of the EP to cause proteins to form complexes (Fig 5) might not be a coincidence, but could be a necessary step in the assembly of BMCs. Structured aggregation of the core enzymes has been proposed to be the initial step in metabolosome assembly and is known to be the first step of β-carboxysome biogenesis, where the core enzyme Ribulose Bisphosphate Carboxylase/Oxygenase (RuBisCO) is aggregated by the CcmM protein. Likewise, CsoS2, a protein in the α-carboxysome core, also aggregates when purified and is proposed to facilitate the nucleation and encapsulation of RuBisCO molecules in the lumen of the organelle. Coupled with protein–protein interactions with other luminal components, the aggregation of these enzymes could lead to a densely packed organelle core. This role for EPs in BMC assembly is in addition to their interaction with shell proteins. Moreover, the PduL crystal structures offer a clue as to how required cofactors enter the BMC lumen during assembly. Free CoA and NAD+/H could potentially be bound to the enzymes as the core assembles and is encapsulated. However, this raises an issue of stoichiometry: if the ratio of cofactors to core enzymes is too low, then the sequestered metabolism would proceed at suboptimal rates. Our PduL crystals contained CoA that was captured from the Escherichia coli cytosol, indicating that the “ground state” of PduL is in the CoA-bound form; this could provide an elegantly simple means of guaranteeing a 1:1 ratio of CoA:PduL within the metabolosome lumen. Active Site Identification and Structural Insights into Catalysis The active site of PduL is formed at the interface of the two structural domains (Fig 2b). As expected, the amino acid sequence conservation is highest in the region around the proposed active site (Fig 4d); highly conserved residues are also involved in CoA binding (Figs 2a and 3, residues Ser45, Lys70, Arg97, Leu99, His204, Asn211). All of the metal-coordinating residues (Fig 2a) are absolutely conserved, implicating them in catalysis or the correct spatial orientation of the substrates. Arg103, which contacts the phosphate (Fig 4b), is present in all PduL homologs. The close resemblance between the structures binding CoA and phosphate likely indicates that no large changes in protein conformation are involved in catalysis, and that our crystal structures are representative of the active form. The native substrate for the forward reaction of rPduL and pPduL, propionyl-CoA, most likely binds to the enzyme in the same way at the observed nucleotide and pantothenic acid moiety, but the propionyl group in the CoA-thioester might point in a different direction. There is a pocket nearby the active site between the well-conserved residues Ser45 and Ala154, which could accommodate the propionyl group (S6 Fig). A homology model of sPduL indicates that the residues making up this pocket and the surrounding active site region are identical to that of rPduL, which is not surprising, because these two homologs presumably have the same propionyl-CoA substrate. The homology model of pPduL also has identical residues making up the pocket, but with a key difference in the vicinity of the active site: Gln77 of rPduL is replaced by a tyrosine (Tyr77) in pPduL. The physiological substrate of pPduL (Table 1) is thought to be lactyl-CoA, which contains an additional hydroxyl group relative to propionyl-CoA. The presence of an aromatic residue at this position may underlie the substrate preference of the PduL enzyme from the pvm locus. Indeed, in the majority of PduLs encoded in pvm loci, Gln77 is substituted by either a Tyr or Phe, whereas it is typically a Gln or Glu in PduLs in all other BMC types that degrade acetyl- or propionyl-CoA. A comparison of the PduL active site to that of the functionally identical Pta suggests that the two enzymes have distinctly different mechanisms. The catalytic mechanism of Pta involves the abstraction of a thiol hydrogen by an aspartate residue, resulting in the nucleophilic attack of thiolate upon the carbonyl carbon of acetyl-phosphate, oriented by an arginine and stabilized by a serine —there are no metals involved. In contrast, in the rPduL structure, there are no conserved aspartate residues in or around the active site, and the only well-conserved glutamate residue in the active site is involved in coordinating one of the metal ions. These observations strongly suggest that an acidic residue is not directly involved in catalysis by PduL. Instead, the dimetal active site of PduL may create a nucleophile from one of the hydroxyl groups on free phosphate to attack the carbonyl carbon of the thioester bond of an acyl-CoA. In the reverse direction, the metal ion(s) could stabilize the thiolate anion that would attack the carbonyl carbon of an acyl-phosphate; a similar mechanism has been described for phosphatases where hydroxyl groups or hydroxide ions can act as a base when coordinated by a dimetal active site. Our structures provide the foundation for studies to elucidate the details of the catalytic mechanism of PduL. Conserved residues in the active site that may contribute to substrate binding and/or transition state stabilization include Ser127, Arg103, Arg194, Gln107, Gln74, and Gln/Glu77. In the phosphate-bound crystal structure, Ser127 and Arg103 appear to position the phosphate (Fig 4b). Alternatively, Arg103 might act as a base to render the phosphate more nucleophilic. The functional groups of Gln74, Gln/Glu77, and Arg194 are directed away from the active site in both CoA and phosphate-bound crystal structures and do not appear to be involved in hydrogen bonding with these substrates, although they could be important for positioning an acyl-phosphate. The free CoA-bound form is presumably poised for attack upon an acyl-phosphate, indicating that the enzyme initially binds CoA as opposed to acyl-phosphate. This hypothesis is strengthened by the fact that the CoA-bound crystals were obtained without added CoA, indicating that the protein bound CoA from the E. coli expression strain and retained it throughout purification and crystallization. The phosphate-bound structure indicates that in the opposite reaction direction phosphate is bound first, and then an acyl-CoA enters. The two high-resolution crystal structures presented here will serve as the foundation for mechanistic studies on this noncanonical PTAC enzyme to determine how the dimetal active site functions to catalyze both forward and reverse reactions. Functional, but Not Structural, Convergence of PduL and Pta PduL and Pta are mechanistically and structurally distinct enzymes that catalyze the same reaction, a prime example of evolutionary convergence upon a function. There are several examples of such functional convergence of enzymes, although typically the enzymes have independently evolved similar, or even identical active sites; for example, the carbonic anhydrase family. However, apparently less frequent is functional convergence that is supported by distinctly different active sites and accordingly catalytic mechanism, as revealed by comparison of the structures of Pta and PduL. One well-studied example of this is the β-lactamase family of enzymes, in which the active site of Class A and Class C enzymes involve serine-based catalysis, but Class B enzymes are metalloproteins. This is not surprising, as β-lactamases are not so widespread among bacteria and therefore would be expected to have evolved independently several times as a defense mechanism against β-lactam antibiotics. However, nearly all bacteria encode Pta, and it is not immediately clear why the Pta/PduL functional convergence should have evolved: it would seem to be evolutionarily more resourceful for the Pta-encoding gene to be duplicated and repurposed for BMCs, as is apparently the case in one type of BMC—EUT1 (Table 1). There could be some intrinsic biochemical difference between the two enzymes that renders PduL a more attractive candidate for encapsulation in a BMC—for example, PduL might be more amenable to tight packaging, or is better suited for the chemical microenvironment formed within the lumen of the BMC, which can be quite different from the cytosol. Further biochemical comparison between the two PTACs will likely yield exciting results that could answer this evolutionary question. Implications BMCs are now known to be widespread among the bacteria and are involved in critical segments of both autotrophic and heterotrophic biochemical pathways that confer to the host organism a competitive (metabolic) advantage in select niches. As one of the three common metabolosome core enzymes, the structure of PduL provides a key missing piece to our structural picture of the shared core biochemistry (Fig 1) of functionally diverse catabolic BMCs. We have observed the oligomeric state differences of PduL to correlate with the presence of an EP, providing new insight into the function of this sequence extension in BMC assembly. Moreover, our results suggest a means for Coenzyme A incorporation during metabolosome biogenesis. A detailed understanding of the underlying principles governing the assembly and internal structural organization of BMCs is a requisite for synthetic biologists to design custom nanoreactors that use BMC architectures as a template. Furthermore, given the growing number of metabolosomes implicated in pathogenesis, the PduL structure will be useful in the development of therapeutics. It is gradually being realized that the metabolic capabilities of a pathogen are also important for virulence, along with the more traditionally cited factors like secretion systems and effector proteins. The fact that PduL is confined almost exclusively to metabolosomes can be used to develop an inhibitor that blocks only PduL and not Pta as a way to selectively disrupt BMC-based metabolism, while not affecting most commensal organisms that require PTAC activity. Materials and Methods Molecular Cloning Genes for PduL homologs with and without the EP were amplified via PCR using the primers listed in S1 Table. sPduL was amplified using S. enterica Typhimurium LT2 genomic DNA, and pPduL and rPduL sequences were codon optimized and synthesized by GenScript with the 6xHis tag. All 5’ primers included EcoRI and BglII restriction sites, and all 3’ primers included a BamHI restriction site to facilitate cloning using the BglBricks strategy. 5’ primers also included the sequence TTTAAGAAGGAGATATACCATG downstream of the restriction sites, serving as a strong ribosome binding site. The 6x polyhistidine tag sequence was added to the 3’ end of the gene using the BglBricks strategy and was subcloned into the pETBb3 vector, a pET21b-based vector modified to be BglBricks compatible. Protein Purification, Size Exclusion Chromatography, and Protein Crystallization  E. coli BL21(DE3) expression strains containing the relevant PduL construct in the pETBb3 vector were grown overnight at 37°C in standard LB medium and then used to inoculate 1L of standard LB medium in 2.8 L Fernbach flasks at a 1:100 dilution, which were then incubated at 37°C shaking at 150 rpm, until the culture reached an OD600 of 0.8–1.0, at which point cultures were induced with 200 μM IPTG (isopropylthio-β-D-galactoside) and incubated at 20°C for 18 h, shaking at 150 rpm. Cells were centrifuged at 5,000 xg for 15 min, and cell pellets were frozen at –20°C. For protein purifications, cell pellets from 1–3 L cultures were resuspended in 20–30 ml buffer A (50 mM Tris-HCl pH 7.4, 300 mM NaCl) and lysed using a French pressure cell at 20,000 lb/in2. The resulting cell lysate was centrifuged at 15,000 xg. 30 mM imidazole was added to the supernatant that was then applied to a 5 mL HisTrap column (GE Healthcare Bio-Sciences, Pittsburgh, PA). Protein was eluted off the column using a gradient of buffer A from 0 mM to 500 mM imidazole over 20 column volumes. Fractions corresponding to PduL were pooled and concentrated using Amicon Ultra Centrifugal filters (EMD Millipore, Billerica, MA) to a volume of no more than 2.5 mL. The protein sample was then applied to a HiLoad 26/60 Superdex 200 preparative size exclusion column (GE Healthcare Bio-Sciences, Pittsburgh, PA) and eluted with buffer B (20 mM Tris pH 7.4, 50 mM NaCl). Where applicable, fractions corresponding to different oligomeric states were pooled separately, leaving one or two fractions in between to prevent cross contamination. Pooled fractions were concentrated to 1–20 mg/mL protein as determined by the Bradford method prior to applying on a Superdex 200 10/300 GL analytical size exclusion column (GE Healthcare Bio-Sciences, Pittsburgh, PA). Size standards used were Thyroglobulin 670 kDa, γ-globulin 158 kDa, Ovalbumin 44 kDa, and Myoglobin 17 kDa. For light scattering, the proteins were measured in a Protein Solutions Dynapro dynamic light scattering instrument with an acquisition time of 5 s, averaging 10 acquisitions at a constant temperature of 25°C. The radii were calculated assuming a globular particle shape. Size exclusion chromatography coupled with SEC-MALS was performed on full-length rPduL and rPduL-ΔEP similar to Luzi et al. 2015. A Wyatt DAWN Heleos-II 18-angle light scattering instrument was used in tandem with a GE AKTA pure FPLC with built in UV detector, and a Wyatt Optilab T-Rex refractive index detector. Detector 16 of the DAWN Heleos-II was replaced with a Wyatt Dynapro Nanostar QELS detector for dynamic light scattering. A GE Superdex S200 10/300 GL column was used, with 125–100 μl of protein sample at 1 mg/ml concentration injected, and the column run at 0.5 ml/min in 20 mM Tris, 50 mM NaCl, pH 7.4. Each detector of the DAWN-Heleos-II was plotted with the Zimm model in the Wyatt ASTRA software to calculate the molar mass. The molar mass was measured at each collected data point across the peaks at ~1 point per 8 μl eluent. Both the Mw and Mn of the molar mass calculations, as well as percent deviations, were also determined using Wyatt software program ASTRA. For preparing protein for crystallography, expression cells were grown as above, except were induced with 50 μM IPTG. Harvested cells were resuspended in buffer B and lysed using a French Press. Cleared lysate was applied on a 5 ml HisTrap HP column (GE Healthcare) and washed with buffer A containing 20 mM imidazole. Pdul-His was eluted with 2 CV buffer B containing 300 mM imidazole, concentrated and then applied on a HiLoad 26/60 Superdex 200 (GE Healthcare) column equilibrated in buffer B for final cleanup. Protein was then concentrated to 20–30 mg/ml for crystallization. Crystals were obtained from sitting drop experiments at 22°C, mixing 3 μl of protein solution with 3 μl of reservoir solution containing 39%–35% MPD. Crystals were flash frozen in liquid nitrogen after being adding 5 μl of a reservoir solution. For heavy atom derivatives, 0.2 μl of 100 mM Thiomerosal (Hampton Research) was added to the crystallization drop 36 h prior to freezing. For phosphate soaks, 5 μl reservoir and 1.5 μl 200 mM sodium phosphate solution (pH 7.0) were added 2 d prior to flash freezing. PTAC Activity Assay Enzyme reactions were performed in a 2 mL cuvette containing 50 mM Tris-HCl pH 7.5, 0.2 mM 5,5'-dithiobis-2-nitrobenzoic acid (DTNB; Ellman’s reagent), 0.1 mM acyl-CoA, and 0.5 μg purified PTAC, unless otherwise noted. To initiate the reaction, 5 mM NaH2PO4 was added, the cuvette was inverted to mix, and the absorbance at 412 nm was measured every 2 s over the course of four minutes in a Nanodrop 2000c, in the cuvette holder. 14,150 M-1cm-1 was used as the extinction coefficient of DTNB to determine the specific activity. PduL Sequence Analysis A multiple sequence alignment of 228 PduL sequences associated with BMCs and 20 PduL sequences not associated with BMCs was constructed using MUSCLE. PduL sequences associated with BMCs were determined from Dataset S1 of Reference, and those not associated with BMCs were determined by searching for genomes that encoded PF06130 but not PF03319 nor PF00936 in the IMG database. The multiple sequence alignment was visualized in Jalview, and the nonconserved N- and C-terminal amino acids were deleted. This trimmed alignment was used to build the sequence logo using WebLogo. Diffraction Data Collection, Structure Determination and Visualization Diffraction data were collected at the Advanced Light Source at Lawrence Berkeley National Laboratory beamline 5.0.2 (100 K, 1.0000 Å wavelength for native data, 1.0093 Å for mercury derivative, 1.2861 Å for Zn pre-edge and 1.2822 Å for Zn peak). Diffraction data were integrated with XDS and scaled with SCALA (CCP4). The structure of PduL was solved using phenix.autosol, which found 11 heavy atom sites and produced density suitable for automatic model building. The model was refined with phenix.refine, with refinement alternating with model building using 2Fo-Fc and Fo-Fc maps visualized in COOT. Statistics for diffraction data collection, structure determination and refinement are summarized in Table 2. Figures were prepared using pymol (www.pymol.org) and Raster3D. Homology Modeling Models of S. enterica Typhimurium LT2 and P. limnophilus PduL were generated with Modeller using the align2d and model-default scripts. Supporting Information Abbreviations Ack acetate kinase BMC Bacterial Microcompartment EP encapsulation peptide EUT ethanolamine-utilizing Mn number average Mw weighted average PDU propanediol-utilizing Pta phosphotransacylase PTAC phosphotransacylase RuBisCO Ribulose Bisphosphate Carboxylase/Oxygenase SEC-MALS multiangle light scattering References Bacterial microcompartments and the modular construction of microbial metabolism A taxonomy of bacterial microcompartment loci constructed by a novel scoring method The PduL phosphotransacylase is used to recycle coenzyme A within the Pdu microcompartment PduL is an Evolutionarily Distinct Phosphotransacylase Involved in B12-dependent 1,2-propanediol degradation by Salmonella enterica serovar typhimurium LT2 Protein structures forming the shell of primitive bacterial organelles Substrate channels revealed in the trimeric Lactobacillus reuteri bacterial microcompartment shell protein PduB The Propanediol Utilization (pdu) Operon of Salmonella enterica serovar Typhimurium LT2 Includes Genes Necessary for Formation of Polyhedral Organelles Involved in coenzyme B12-Dependent 1, 2-Propanediol Degradation The Distribution of Polyhedral Bacterial Microcompartments Suggests Frequent Horizontal Transfer and Operon Reassembly Using comparative genomics to uncover new kinds of protein-based metabolic organelles in bacteria Ethanolamine utilization in Salmonella typhimurium  PduP is a coenzyme-a-acylating propionaldehyde dehydrogenase associated with the polyhedral bodies involved in B12-dependent 1,2-propanediol degradation by Salmonella enterica serovar Typhimurium LT2 Evidence that a metabolic microcompartment contains and recycles private cofactor pools The PduQ enzyme is an alcohol dehydrogenase used to recycle NAD+ internally within the Pdu microcompartment of Salmonella enterica  Activation of acetate by Methanosarcina thermophila. Purification and characterization of phosphotransacetylase Enzymology of the fermentation of acetate to methane by Methanosarcina thermophila Acetate formation in the energy metabolism of parasitic helminths and protists IMG: the Integrated Microbial Genomes database and comparative analysis system Pfam: the protein families database The Pfam protein families database The purification and properties of phosphotransacetylase Phosphotransacetylase from Bacillus subtilis: purification and physiological studies Fluorometric determination of glycolytic intermediates and adenylates during sequential changes in replacement culture of Aspergillus niger Bacterial Microcompartment Assembly: The Key Role of Encapsulation Peptides Interactions between the termini of lumen enzymes and shell proteins mediate enzyme encapsulation into bacterial microcompartments Elucidating the essential role of the conserved carboxysomal protein CcmN reveals a common feature of bacterial microcompartment assembly The N-Terminal Regions of β and γ Subunits Lower the Solubility of Adenosylcobalamin-Dependent Diol Dehydratase Crystal structures of ethanolamine ammonia-lyase complexed with coenzyme B12 analogs and substrates Bacterial microcompartments moving into a synthetic biological world Streamlined construction of the cyanobacterial CO2-fixing organelle via protein domain fusions Structural and functional studies suggest a catalytic mechanism for the phosphotransacetylase from Methanosarcina thermophila Characterization of a planctomycetal organelle: a novel bacterial microcompartment for the aerobic degradation of plant saccharides Geometry of interaction of metal ions with histidine residues in protein structures The PduL Phosphotransacylase Is Used To Recycle Coenzyme A within the Pdu Microcompartment Crystal structure of ethylbenzene dehydrogenase from Aromatoleum aromaticum Advances in Understanding Carboxysome Assembly in Prochlorococcus and Synechococcus Implicate CsoS2 as a Critical Component Biogenesis of a Bacterial Organelle: The Carboxysome Assembly Pathway Solution structure of a bacterial microcompartment targeting peptide and its application in the construction of an ethanol bioreactor Enzymatic mechanisms of phosphate and sulfate transfer The active site architecture of Pisum sativum beta-carbonic anhydrase is a mirror image of that of alpha-carbonic anhydrases Carbonic anhydrase: new insights for an ancient enzyme The 3-D structure of a zinc metallo-beta-lactamase from Bacillus cereus reveals a new type of protein fold Three decades of beta-lactamase inhibitors Structural basis of the oxidative activation of the carboxysomal gamma-carbonic anhydrase, CcmM The bacterial carbon-fixing organelle is formed by shell envelopment of preassembled cargo  Salmonella enterica typhimurium colonizing the lumen of the chicken intestine grows slowly and upregulates a unique set of virulence and metabolism genes Identification of Listeria monocytogenes Genes Contributing to Intracellular Replication by Expression Profiling and Mutant Screening Ethanolamine controls expression of genes encoding components involved in interkingdom signaling and virulence in enterohemorrhagic Escherichia coli O157:H7 Identification of novel genes in genomic islands that contribute to Salmonella typhimurium replication in macrophages  Enterococcus faecalis mutations affecting virulence in the Caenorhabditis elegans model host Redefining Virulence of Bacterial Pathogens A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding Subunit disassembly and inhibition of TNFalpha by a semi-synthetic bicyclic peptide MUSCLE: multiple sequence alignment with high accuracy and high throughput Jalview Version 2—a multiple sequence alignment editor and analysis workbench WebLogo: a sequence logo generator XDS Overview of the CCP4 suite and current developments Towards automated crystallographic structure refinement with phenix.refine Coot: model-building tools for molecular graphics Raster3D: photorealistic molecular graphics Comparative protein structure modeling of genes and 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diff --git a/annotated_BioC_JSON/PMC4786784_ann.json b/annotated_BioC_JSON/PMC4786784_ann.json
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+[{"sourceid":"4786784","sourcedb":"","project":"","target":"","text":"An extended U2AF65–RNA-binding domain recognizes the 3′ splice site signal How the essential pre-mRNA splicing factor U2AF65 recognizes the polypyrimidine (Py) signals of the major class of 3′ splice sites in human gene transcripts remains incompletely understood. We determined four structures of an extended U2AF65–RNA-binding domain bound to Py-tract oligonucleotides at resolutions between 2.0 and 1.5 Å. These structures together with RNA binding and splicing assays reveal unforeseen roles for U2AF65 inter-domain residues in recognizing a contiguous, nine-nucleotide Py tract. The U2AF65 linker residues between the dual RNA recognition motifs (RRMs) recognize the central nucleotide, whereas the N- and C-terminal RRM extensions recognize the 3′ terminus and third nucleotide. Single-molecule FRET experiments suggest that conformational selection and induced fit of the U2AF65 RRMs are complementary mechanisms for Py-tract association. Altogether, these results advance the mechanistic understanding of molecular recognition for a major class of splice site signals.  The pre-mRNA splicing factor U2AF65 recognizes 3′ splice sites in human gene transcripts, but the details are not fully understood. Here, the authors report U2AF65 structures and single molecule FRET that reveal mechanistic insights into splice site recognition. The differential skipping or inclusion of alternatively spliced pre-mRNA regions is a major source of diversity for nearly all human gene transcripts. The splice sites are marked by relatively short consensus sequences and are regulated by additional pre-mRNA motifs (reviewed in ref.). At the 3′ splice site of the major intron class, these include a polypyrimidine (Py) tract comprising primarily Us or Cs, which is preceded by a branch point sequence (BPS) that ultimately serves as the nucleophile in the splicing reaction and an AG-dinucleotide at the 3′ splice site junction. Disease-causing mutations often compromise pre-mRNA splicing (reviewed in refs), yet a priori predictions of splice sites and the consequences of their mutations are challenged by the brevity and degeneracy of known splice site sequences. High-resolution structures of intact splicing factor–RNA complexes would offer key insights regarding the juxtaposition of the distinct splice site consensus sequences and their relationship to disease-causing point mutations. The early-stage pre-mRNA splicing factor U2AF65 is essential for viability in vertebrates and other model organisms (for example, ref.). A tightly controlled assembly among U2AF65, the pre-mRNA, and partner proteins sequentially identifies the 3′ splice site and promotes association of the spliceosome, which ultimately accomplishes the task of splicing. Initially U2AF65 recognizes the Py-tract splice site signal. In turn, the ternary complex of U2AF65 with SF1 and U2AF35 identifies the surrounding BPS and 3′ splice site junctions. Subsequently U2AF65 recruits the U2 small nuclear ribonucleoprotein particle (snRNP) and ultimately dissociates from the active spliceosome. Biochemical characterizations of U2AF65 demonstrated that tandem RNA recognition motifs (RRM1 and RRM2) recognize the Py tract (Fig. 1a). Milestone crystal structures of the core U2AF65 RRM1 and RRM2 connected by a shortened inter-RRM linker (dU2AF651,2) detailed a subset of nucleotide interactions with the individual U2AF65 RRMs. A subsequent NMR structure characterized the side-by-side arrangement of the minimal U2AF65 RRM1 and RRM2 connected by a linker of natural length (U2AF651,2), yet depended on the dU2AF651,2 crystal structures for RNA interactions and an ab initio model for the inter-RRM linker conformation. As such, the molecular mechanisms for Py-tract recognition by the intact U2AF65–RNA-binding domain remained unknown. Here, we use X-ray crystallography and biochemical studies to reveal new roles in Py-tract recognition for the inter-RRM linker and key residues surrounding the core U2AF65 RRMs. We use single-molecule Förster resonance energy transfer (smFRET) to characterize the conformational dynamics of this extended U2AF65–RNA-binding domain during Py-tract recognition. Results Cognate U2AF65–Py-tract recognition requires RRM extensions The RNA affinity of the minimal U2AF651,2 domain comprising the core RRM1–RRM2 folds (U2AF651,2, residues 148–336) is relatively weak compared with full-length U2AF65 (Fig. 1a,b; Supplementary Fig. 1). Historically, this difference was attributed to the U2AF65 arginine–serine rich domain, which contacts pre-mRNA–U2 snRNA duplexes outside of the Py tract. We noticed that the RNA-binding affinity of the U2AF651,2 domain was greatly enhanced by the addition of seven and six residues at the respective N and C termini of the minimal RRM1 and RRM2 (U2AF651,2L, residues 141–342; Fig. 1a). In a fluorescence anisotropy assay for binding a representative Py tract derived from the well-characterized splice site of the adenovirus major late promoter (AdML), the RNA affinity of U2AF651,2L increased by 100-fold relative to U2AF651,2 to comparable levels as full-length U2AF65 (Fig. 1b; Supplementary Fig. 1a–d). Likewise, both U2AF651,2L and full-length U2AF65 showed similar sequence specificity for U-rich stretches in the 5′-region of the Py tract and promiscuity for C-rich regions in the 3′-region (Fig. 1c, Supplementary Fig. 1e–h). U2AF65-bound Py tract comprises nine contiguous nucleotides To investigate the structural basis for cognate U2AF65 recognition of a contiguous Py tract, we determined four crystal structures of U2AF651,2L bound to Py-tract oligonucleotides (Fig. 2a; Table 1). By sequential boot strapping (Methods), we optimized the oligonucleotide length, the position of a Br-dU, and the identity of the terminal nucleotide (rU, dU and rC) to achieve full views of U2AF651,2L bound to contiguous Py tracts at up to 1.5 Å resolution. The protein and oligonucleotide conformations are nearly identical among the four new U2AF651,2L structures (Supplementary Fig. 2a). The U2AF651,2L RRM1 and RRM2 associate with the Py tract in a parallel, side-by-side arrangement (shown for representative structure iv in Fig. 2b,c; Supplementary Movie 1). An extended conformation of the U2AF65 inter-RRM linker traverses across the α-helical surface of RRM1 and the central β-strands of RRM2 and is well defined in the electron density (Fig. 2b). The extensions at the N terminus of RRM1 and C terminus of RRM2 adopt well-ordered α-helices. Both RRM1/RRM2 extensions and the inter-RRM linker of U2AF651,2L directly recognize the bound oligonucleotide. We compare the global conformation of the U2AF651,2L structures with the prior dU2AF651,2 crystal structure and U2AF651,2 NMR structure in the Supplementary Discussion and Supplementary Fig. 2. The discovery of nine U2AF65-binding sites for contiguous Py-tract nucleotides was unexpected. Based on dU2AF651,2 structures, we originally hypothesized that the U2AF65 RRMs would bind the minimal seven nucleotides observed in these structures. Surprisingly, the RRM2 extension/inter-RRM linker contribute new central nucleotide-binding sites near the RRM1/RRM2 junction and the RRM1 extension recognizes the 3′-terminal nucleotide (Fig. 2c; Supplementary Movie 1). The U2AF651,2L structures characterize ribose (r) nucleotides at all of the binding sites except the seventh and eighth deoxy-(d)U, which are likely to lack 2′-hydroxyl contacts based on the RNA-bound dU2AF651,2 structure. Qualitatively, a subset of the U2AF651,2L-nucleotide-binding sites (sites 1–3 and 7–9) share similar locations to those of the dU2AF651,2 structures (Supplementary Figs 2c,d and 3). Yet, only the U2AF651,2L interactions at sites 1 and 7 are nearly identical to those of the dU2AF651,2 structures (Supplementary Fig. 3a,f). In striking departures from prior partial views, the U2AF651,2L structures reveal three unanticipated nucleotide-binding sites at the centre of the Py tract, as well as numerous new interactions that underlie cognate recognition of the Py tract (Fig. 3a–h). U2AF65 inter-RRM linker interacts with the Py tract The U2AF651,2L RRM2, the inter-RRM linker and RRM1 concomitantly recognize the three central nucleotides of the Py tract, which are likely to coordinate the conformational arrangement of these disparate portions of the protein. Residues in the C-terminal region of the U2AF65 inter-RRM linker comprise a centrally located binding site for the fifth nucleotide on the RRM2 surface and abutting the RRM1/RRM2 interface (Fig. 3d). The backbone amide of the linker V254 and the carbonyl of T252 engage in hydrogen bonds with the rU5-O4 and -N3H atoms. In the C-terminal β-strand of RRM1, the side chains of K225 and R227 donate additional hydrogen bonds to the rU5-O2 lone pair electrons. The C-terminal region of the inter-RRM linker also participates in the preceding rU4-binding site, where the V254 backbone carbonyl and D256 carboxylate position the K260 side chain to hydrogen bond with the rU4-O4 (Fig. 3c). Otherwise, the rU4 nucleotide packs against F304 in the signature ribonucleoprotein consensus motif (RNP)-2 of RRM2. At the opposite side of the central fifth nucleotide, the sixth rU6 nucleotide is located at the inter-RRM1/RRM2 interface (Fig. 3e; Supplementary Movie 1). This nucleotide twists to face away from the U2AF65 linker and instead inserts the rU6-uracil into a sandwich between the β2/β3 loops of RRM1 and RRM2. The rU6 base edge is relatively solvent exposed; accordingly, the rU6 hydrogen bonds with U2AF65 are water mediated apart from a single direct interaction by the RRM1-N196 side chain. We tested the contribution of the U2AF651,2L interactions with the new central nucleotide to Py-tract affinity (Fig. 3i; Supplementary Fig. 4a,b). Mutagenesis of either V254 in the U2AF65 inter-RRM linker to proline or RRM1–R227 to alanine, which remove the hydrogen bond with the fifth uracil-O4 or -O2, reduced the affinities of U2AF651,2L for the representative AdML Py tract by four- or five-fold, respectively. The energetic penalties due to these mutations (ΔΔG 0.8–0.9 kcal mol−1) are consistent with the loss of each hydrogen bond with the rU5 base and support the relevance of the central nucleotide interactions observed in the U2AF651,2L structures. U2AF65 RRM extensions interact with the Py tract The N- and C-terminal extensions of the U2AF65 RRM1 and RRM2 directly contact the bound Py tract. Rather than interacting with a new 5′-terminal nucleotide as we had hypothesized, the C-terminal α-helix of RRM2 instead folds across one surface of rU3 in the third binding site (Fig. 3b). There, a salt bridge between the K340 side chain and nucleotide phosphate, as well as G338-base stacking and a hydrogen bond between the backbone amide of G338 and the rU3-O4, secure the RRM2 extension. Indirectly, the additional contacts with the third nucleotide shift the rU2 nucleotide in the second binding site closer to the C-terminal β-strand of RRM2. Consequently, the U2AF651,2L-bound rU2-O4 and -N3H form dual hydrogen bonds with the K329 backbone atoms (Fig. 3a), rather than a single hydrogen bond with the K329 side chain as in the prior dU2AF651,2 structure (Supplementary Fig. 3b). At the N terminus, the α-helical extension of U2AF65 RRM1 positions the Q147 side chain to bridge the eighth and ninth nucleotides at the 3′ terminus of the Py tract (Fig. 3f–h). The Q147 residue participates in hydrogen bonds with the -N3H of the eighth uracil and -O2 of the ninth pyrimidine. The adjacent R146 guanidinium group donates hydrogen bonds to the 3′-terminal ribose-O2′ and O3′ atoms, where it could form a salt bridge with a phospho-diester group in the context of a longer pre-mRNA. Consistent with loss of a hydrogen bond with the ninth pyrimidine-O2 (ΔΔG 1.0 kcal mol−1), mutation of the Q147 to an alanine reduced U2AF651,2L affinity for the AdML Py tract by five-fold (Fig. 3i; Supplementary Fig. 4c). We compare U2AF65 interactions with uracil relative to cytosine pyrimidines at the ninth binding site in Fig. 3g,h and the Supplementary Discussion. Versatile primary sequence of the U2AF65 inter-RRM linker The U2AF651,2L structures reveal that the inter-RRM linker mediates an extensive interface with the second α-helix of RRM1, the β2/β3 strands of RRM2 and the N-terminal α-helical extension of RRM1. Altogether, the U2AF65 inter-RRM linker residues (R228–K260) bury 2,800 Å2 of surface area in the U2AF651,2L holo-protein, suggestive of a cognate interface compared with 1,900 Å2 for a typical protein–protein complex. The path of the linker initiates at P229 following the core RRM1 β-strand, in a kink that is positioned by intra-molecular stacking among the consecutive R228, Y232 and P234 side chains (Fig. 4a, lower right). A second kink at P236, coupled with respective packing of the L235 and M238 side chains on the N-terminal α-helical RRM1 extension and the core RRM1 α2-helix, reverses the direction of the inter-RRM linker towards the RRM1/RRM2 interface and away from the RNA-binding site. In the neighbouring apical region of the linker, the V244 and V246 side chains pack in a hydrophobic pocket between two α-helices of the core RRM1. The adjacent V249 and V250 are notable for their respective interactions that connect RRM1 and RRM2 at this distal interface from the RNA-binding site (Fig. 4a, top). A third kink stacks P247 and G248 with Y245 and re-orients the C-terminal region of the linker towards the RRM2 and bound RNA. At the RNA surface, the key V254 that recognizes the fifth uracil is secured via hydrophobic contacts between its side chain and the β-sheet surface of RRM2, chiefly the consensus RNP1-F304 residue that stacks with the fourth uracil (Fig. 4a, lower left). Few direct contacts are made between the remaining residues of the linker and the U2AF65 RRM2; instead, the C-terminal conformation of the linker appears primarily RNA mediated (Fig. 3c,d). We investigated whether the observed contacts between the RRMs and linker were critical for RNA binding by structure-guided mutagenesis (Fig. 4b). We titrated these mutant U2AF651,2L proteins into fluorescein-labelled AdML Py-tract RNA and fit the fluorescence anisotropy changes to obtain the apparent equilibrium affinities (Supplementary Fig. 4d–h). We introduced glycine substitutions to maximally reduce the buried surface area without directly interfering with its hydrogen bonds between backbone atoms and the base. First, we replaced V249 and V250 at the RRM1/RRM2 interface and V254 at the bound RNA site with glycine (3Gly). However, the resulting decrease in the AdML RNA affinity of the U2AF651,2L-3Gly mutant relative to wild-type protein was not significant (Fig. 4b). In parallel, we replaced five linker residues (S251, T252, V253, V254 and P255) at the fifth nucleotide-binding site with glycines (5Gly) and also found that the RNA affinity of the U2AF651,2L-5Gly mutant likewise decreased only slightly relative to wild-type protein. A more conservative substitution of these five residues (251–255) with an unrelated sequence capable of backbone-mediated hydrogen bonds (STVVP\u003eNLALA) confirmed the subtle impact of this versatile inter-RRM sequence on affinity for the AdML Py tract. Finally, to ensure that these selective mutations were sufficient to disrupt the linker/RRM contacts, we substituted glycine for the majority of buried hydrophobic residues in the inter-RRM linker (including M144, L235, M238, V244, V246, V249, V250, S251, T252, V253, V254, P255; called 12Gly). Despite 12 concurrent mutations, the AdML RNA affinity of the U2AF651,2L-12Gly variant was reduced by only three-fold relative to the unmodified protein (Fig. 4b), which is less than the penalty of the V254P mutation that disrupts the rU5 hydrogen bond (Fig. 3d,i). To test the interplay of the U2AF65 inter-RRM linker with its N- and C-terminal RRM extensions, we constructed an internal linker deletion of 20-residues within the extended RNA-binding domain (dU2AF651,2L). We found that the affinity of dU2AF651,2L for the AdML RNA was significantly reduced relative to U2AF651,2L (four-fold, Figs 1b and 4b; Supplementary Fig. 4i). Yet, it is well known that the linker deletion in the context of the minimal RRM1–RRM2 boundaries has no detectable effect on the RNA affinities of dU2AF651,2 compared with U2AF651,2 (refs; Figs 1b and 4b; Supplementary Fig. 4j). The U2AF651,2L structures suggest that an extended conformation of the truncated dU2AF651,2 inter-RRM linker would suffice to connect the U2AF651,2L RRM1 C terminus to the N terminus of RRM2 (24 Å distance between U2AF651,2L R227-Cα–H259-Cα atoms), which agrees with the greater RNA affinities of dU2AF651,2 and U2AF651,2 dual RRMs compared with the individual U2AF65 RRMs. However, stretching of the truncated dU2AF651,2L linker to connect the RRM termini is expected to disrupt its nucleotide interactions. Likewise, deletion of the N-terminal RRM1 extension in the shortened constructs would remove packing interactions that position the linker in a kinked turn following P229 (Fig. 4a), consistent with the lower RNA affinities of dU2AF651,2L, dU2AF651,2 and U2AF651,2 compared with U2AF651,2L. To further test cooperation among the U2AF65 RRM extensions and inter-RRM linker for RNA recognition, we tested the impact of a triple Q147A/V254P/R227A mutation (U2AF651,2L-3Mut) for RNA binding (Fig. 4b; Supplementary Fig. 4d). Notably, the Q147A/V254P/R227A mutation reduced the RNA affinity of the U2AF651,2L-3Mut protein by 30-fold more than would be expected based on simple addition of the ΔΔG's for the single mutations. This difference indicates that the linearly distant regions of the U2AF65 primary sequence, including Q147 in the N-terminal RRM1 extension and R227/V254 in the N-/C-terminal linker regions at the fifth nucleotide site, cooperatively recognize the Py tract. Altogether, we conclude that the conformation of the U2AF65 inter-RRM linker is key for recognizing RNA and is positioned by the RRM extension but otherwise relatively independent of the side chain composition. The non-additive effects of the Q147A/V254P/R227A triple mutation, coupled with the context-dependent penalties of an internal U2AF65 linker deletion, highlights the importance of the structural interplay among the U2AF65 linker and the N- and C-terminal extensions flanking the core RRMs. Importance of U2AF65–RNA contacts for pre-mRNA splicing We proceeded to test the importance of new U2AF65–Py-tract interactions for splicing of a model pre-mRNA substrate in a human cell line (Fig. 5; Supplementary Fig. 5). As a representative splicing substrate, we utilized a well-characterized minigene splicing reporter (called pyPY) comprising a weak (that is, degenerate, py) and strong (that is, U-rich, PY) polypyrimidine tracts preceding two alternative splice sites (Fig. 5a). When transfected into HEK293T cells containing only endogenous U2AF65, the PY splice site is used and the remaining transcript remains unspliced. When co-transfected with an expression plasmid for wild-type U2AF65, use of the py splice site significantly increases (by more than five-fold) and as documented converts a fraction of the unspliced to spliced transcript. The strong PY splice site is insensitive to added U2AF65, suggesting that endogenous U2AF65 levels are sufficient to saturate this site (Supplementary Fig. 5b). We introduced the triple mutation (V254P/R227A/Q147A) that significantly reduced U2AF651,2L association with the Py tract (Fig. 4b) in the context of full-length U2AF65 (U2AF65-3Mut). Co-transfection of the U2AF65-3Mut with the pyPY splicing substrate significantly reduced splicing of the weak ‘py' splice site relative to wild-type U2AF65 (Fig. 5b,c). We conclude that the Py-tract interactions with these residues of the U2AF65 inter-RRM linker and RRM extensions are important for splicing as well as for binding a representative of the major U2-class of splice sites. Sparse inter-RRM contacts underlie apo-U2AF65 dynamics The direct interface between U2AF651,2L RRM1 and RRM2 is minor, burying 265 Å2 of solvent accessible surface area compared with 570 Å2 on average for a crystal packing interface. A handful of inter-RRM hydrogen bonds are apparent between the side chains of RRM1-N155 and RRM2-K292, RRM1-N155 and RRM2-D272 as well as the backbone atoms of RRM1-G221 and RRM2-D273 (Fig. 4c). This minor U2AF65 RRM1/RRM2 interface, coupled with the versatile sequence of the inter-RRM linker, highlighted the potential role for inter-RRM conformational dynamics in U2AF65-splice site recognition. Paramagnetic resonance enhancement (PRE) measurements previously had suggested a predominant back-to-back, or ‘closed' conformation of the apo-U2AF651,2 RRM1 and RRM2 in equilibrium with a minor ‘open' conformation resembling the RNA-bound inter-RRM arrangement. Yet, small-angle X-ray scattering (SAXS) data indicated that both the minimal U2AF651,2 and longer constructs comprise a highly diverse continuum of conformations in the absence of RNA that includes the ‘closed' and ‘open' conformations. To complement the static portraits of U2AF651,2L structure that we had determined by X-ray crystallography, we used smFRET to characterize the probability distribution functions and time dependence of U2AF65 inter-RRM conformational dynamics in solution. The inter-RRM dynamics of U2AF65 were followed using FRET between fluorophores attached to RRM1 and RRM2 (Fig. 6a,b, Methods). The positions of single cysteine mutations for fluorophore attachment (A181C in RRM1 and Q324C in RRM2) were chosen based on inspection of the U2AF651,2L structures and the ‘closed' model of apo-U2AF651,2. Criteria included (i) residue locations that are distant from and hence not expected to interfere with the RRM/RNA or inter-RRM interfaces, (ii) inter-dye distances (50 Å for U2AF651,2L–Py tract and 30 Å for the closed apo-model) that are expected to be near the Förster radius (Ro) for the Cy3/Cy5 pair (56 Å), where changes in the efficiency of energy transfer are most sensitive to distance, and (iii) FRET efficiencies that are calculated to be significantly greater for the ‘closed' apo-model as opposed to the ‘open' RNA-bound structures (by ∼30%). The FRET efficiencies of either of these structurally characterized conformations also are expected to be significantly greater than elongated U2AF65 conformations that lack inter-RRM contacts. Double-cysteine variant of U2AF651,2 was modified with equimolar amount of Cy3 and Cy5. Only traces that showed single photobleaching events for both donor and acceptor dyes and anti-correlated changes in acceptor and donor fluorescence were included in smFRET data analysis. Hence, molecules that were conjugated to two donor or two acceptor fluorophores were excluded from analysis. We first characterized the conformational dynamics spectrum of U2AF65 in the absence of RNA (Fig. 6c,d; Supplementary Fig. 7a,b). The double-labelled U2AF651,2LFRET(Cy3/Cy5) protein was tethered to a slide via biotin-NTA/Ni+2 resin. Virtually no fluorescent molecules were detected in the absence of biotin-NTA/Ni+2, which demonstrates the absence of detectable non-specific binding of U2AF651,2LFRET to the slide. The FRET distribution histogram built from more than a thousand traces of U2AF651,2LFRET(Cy3/Cy5) in the absence of ligand showed an extremely broad distribution centred at a FRET efficiency of ∼0.4 (Fig. 6d). Approximately 40% of the smFRET traces showed apparent transitions between multiple FRET values (for example, Fig. 6c). Despite the large width of the FRET-distribution histogram, the majority (80%) of traces that showed fluctuations sampled only two distinct FRET states (for example, Supplementary Fig. 7a). Approximately 70% of observed fluctuations were interchanges between the ∼0.65 and ∼0.45 FRET values (Supplementary Fig. 7b). We cannot exclude a possibility that tethering of U2AF651,2LFRET(Cy3/Cy5) to the microscope slide introduces structural heterogeneity into the protein and, thus, contributes to the breadth of the FRET distribution histogram. However, the presence of repetitive fluctuations between particular FRET values supports the hypothesis that RNA-free U2AF65 samples several distinct conformations. This result is consistent with the broad ensembles of extended solution conformations that best fit the SAXS data collected for U2AF651,2 as well as for a longer construct (residues 136–347). We conclude that weak contacts between the U2AF65 RRM1 and RRM2 permit dissociation of these RRMs in the absence of RNA. U2AF65 conformational selection and induced fit by bound RNA We next used smFRET to probe the conformational selection of distinct inter-RRM arrangements following association of U2AF65 with the AdML Py-tract prototype. Addition of the AdML RNA to tethered U2AF651,2LFRET(Cy3/Cy5) selectively increases a fraction of molecules showing an ∼0.45 apparent FRET efficiency, suggesting that RNA binding stabilizes a single conformation, which corresponds to the 0.45 FRET state (Fig. 6e,f). To assess the possible contributions of RNA-free conformations of U2AF65 and/or structural heterogeneity introduced by tethering of U2AF651,2LFRET(Cy3/Cy5) to the slide to the observed distribution of FRET values, we reversed the immobilization scheme. We tethered the AdML RNA to the slide via a biotinylated oligonucleotide DNA handle and added U2AF651,2LFRET(Cy3/Cy5) in the absence of biotin-NTA resin (Fig. 6g,h; Supplementary Fig. 7c–g). A 0.45 FRET value was again predominant, indicating a similar RNA-bound conformation and structural dynamics for the untethered and tethered U2AF651,2LFRET(Cy3/Cy5). We examined the effect on U2AF651,2L conformations of purine interruptions that often occur in relatively degenerate human Py tracts. We introduced an rArA purine dinucleotide within a variant of the AdML Py tract (detailed in Methods). Insertion of adenine nucleotides decreased binding affinity of U2AF65 to RNA by approximately five-fold. Nevertheless, in the presence of saturating concentrations of rArA-interrupted RNA slide-tethered U2AF651,2LFRET(Cy3/Cy5) showed a prevalent ∼0.45 apparent FRET value (Fig. 6i,j), which was also predominant in the presence of continuous Py tract. Therefore, RRM1-to-RRM2 distance remains similar regardless of whether U2AF65 is bound to interrupted or continuous Py tract. The inter-fluorophore distances derived from the observed 0.45 FRET state agree with the distances between the α-carbon atoms of the respective residues in the crystal structures of U2AF651,2L bound to Py-tract oligonucleotides. It should be noted that inferring distances from FRET values is prone to significant error because of uncertainties in the determination of fluorophore orientation factor κ2 and Förster radius R0, the parameters used in distance calculations. Nevertheless, the predominant 0.45 FRET state in the presence of RNA agrees with the Py-tract-bound crystal structure of U2AF651,2L. Importantly, the majority of traces (∼70%) of U2AF651,2LFRET(Cy3/Cy5) bound to the slide-tethered RNA lacked FRET fluctuations and predominately exhibited a ∼0.45 FRET value (for example, Fig. 6g). The remaining ∼30% of traces for U2AF651,2LFRET(Cy3/Cy5) bound to the slide-tethered RNA showed fluctuations between distinct FRET values. The majority of traces that show fluctuations began at high (0.65–0.8) FRET value and transitioned to a ∼0.45 FRET value (Supplementary Fig. 7c–g). Hidden Markov modelling analysis of smFRET traces suggests that RNA-bound U2AF651,2L can sample at least two other conformations corresponding to ∼0.7–0.8 and ∼0.3 FRET values in addition to the predominant conformation corresponding to the 0.45 FRET state. Although a compact conformation (or multiple conformations) of U2AF651,2L corresponding to ∼0.7–0.8 FRET values can bind RNA, on RNA binding, these compact conformations of U2AF651,2L transition into a more stable structural state that corresponds to ∼0.45 FRET value and is likely similar to the side-by-side inter-RRM-arrangement of the U2AF651,2L crystal structures. Thus, the sequence of structural rearrangements of U2AF65 observed in smFRET traces (Supplementary Fig. 7c–g) suggests that a ‘conformational selection' mechanism of Py-tract recognition (that is, RNA ligand stabilization of a pre-configured U2AF65 conformation) is complemented by ‘induced fit' (that is, RNA-induced rearrangement of the U2AF65 RRMs to achieve the final ‘side-by-side' conformation), as discussed below. Discussion The U2AF65 structures and analyses presented here represent a successful step towards defining a molecular map of the 3′ splice site. Several observations indicate that the numerous intramolecular contacts, here revealed among the inter-RRM linker and RRM1, RRM2, and the N-terminal RRM1 extension, synergistically coordinate U2AF65–Py-tract recognition. Truncation of U2AF65 to the core RRM1–RRM2 region reduces its RNA affinity by 100-fold. Likewise, deletion of 20 inter-RRM linker residues significantly reduces U2AF65–RNA binding only when introduced in the context of the longer U2AF651,2L construct comprising the RRM extensions, which in turn position the linker for RNA interactions. Notably, a triple mutation of three residues (V254P, Q147A and R227A) in the respective inter-RRM linker, N- and C-terminal extensions non-additively reduce RNA binding by 150-fold. Altogether, these data indicate that interactions among the U2AF65 RRM1/RRM2, inter-RRM linker, N-and C-terminal extensions are mutually inter-dependent for cognate Py-tract recognition. The implications of this finding for U2AF65 conservation and Py-tract recognition are detailed in the Supplementary Discussion. Recently, high-throughput sequencing studies have shown that somatic mutations in pre-mRNA splicing factors occur in the majority of patients with myelodysplastic syndrome (MDS). MDS-relevant mutations are common in the small U2AF subunit (U2AF35, or U2AF1), yet such mutations are rare in the large U2AF65 subunit (also called U2AF2)—possibly due to the selective versus nearly universal requirements of these factors for splicing. A confirmed somatic mutation of U2AF65 in patients with MDS, L187V, is located on a solvent-exposed surface of RRM1 that is distinct from the RNA interface (Fig. 7a). This L187 surface is oriented towards the N terminus of the U2AF651,2L construct, where it is expected to abut the U2AF35-binding site in the context of the full-length U2AF heterodimer. Likewise, an unconfirmed M144I mutation reported by the same group corresponds to the N-terminal residue of U2AF651,2L, which is separated by only ∼20 residues from the U2AF35-binding site. As such, we suggest that the MDS-relevant U2AF65 mutations contribute to MDS progression indirectly, by destabilizing a relevant conformation of the conjoined U2AF35 subunit rather than affecting U2AF65 functions in RNA binding or spliceosome recruitment per se. Our smFRET results agree with prior NMR/PRE evidence for multi-domain conformational selection as one mechanistic basis for U2AF65–RNA association (Fig. 7b). The ‘induced fit' versus ‘conformational selection' models are the prevailing views of the mechanisms underlying bio-molecular interactions (reviewed in ref.). In the former, ligand binding promotes a subsequent conformational change in the protein, whereas in the latter, the ligand selects a protein conformation from a pre-existing ensemble and thereby shifts the population towards that state. An ∼0.45 FRET value is likely to correspond to the U2AF65 conformation visualized in our U2AF651,2L crystal structures, in which the RRM1 and RRM2 bind side-by-side to the Py-tract oligonucleotide. The lesser 0.65–0.8 and 0.2–0.3 FRET values in the untethered U2AF651,2LFRET(Cy3/Cy5) experiment could correspond to respective variants of the ‘closed', back-to-back U2AF65 conformations characterized by NMR/PRE data, or to extended U2AF65 conformations, in which the intramolecular RRM1/RRM2 interactions have dissociated the protein is bound to RNA via single RRMs. An increased prevalence of the ∼0.45 FRET value following U2AF65–RNA binding, coupled with the apparent absence of transitions in many ∼0.45-value single molecule traces (for example, Fig. 6e), suggests a population shift in which RNA binds to (and draws the equilibrium towards) a pre-configured inter-RRM proximity that most often corresponds to the ∼0.45 FRET value. Notably, our smFRET results reveal that U2AF65–Py-tract recognition can be characterized by an ‘extended conformational selection' model (Fig. 7b). In this recent model for macromolecular interactions, the pure ‘conformational selection' and ‘induced fit' scenarios represent the limits of a mechanistic spectrum and may compete or occur sequentially. Examples of ‘extended conformational selection' during ligand binding have been characterized for a growing number of macromolecules (for example, adenylate kinase, LAO-binding protein, poly-ubiquitin, maltose-binding protein and the preQ1 riboswitch, among others). Here, the majority of changes in smFRET traces for U2AF651,2LFRET(Cy3/Cy5) bound to slide-tethered RNA began at high (0.65–0.8) FRET value and transition to the predominant 0.45 FRET value (Supplementary Fig. 7c–g). These transitions could correspond to rearrangement from the ‘closed' NMR/PRE-based U2AF65 conformation in which the RNA-binding surface of only a single RRM is exposed and available for RNA binding, to the structural state seen in the side-by-side, RNA-bound crystal structure. As such, the smFRET approach reconciles prior inconsistencies between two major conformations that were detected by NMR/PRE experiments and a broad ensemble of diverse inter-RRM arrangements that fit the SAXS data for the apo-protein. Similar interdisciplinary structural approaches are likely to illuminate whether similar mechanistic bases for RNA binding are widespread among other members of the vast multi-RRM family. The finding that U2AF65 recognizes a nine base pair Py tract contributes to an elusive ‘code' for predicting splicing patterns from primary sequences in the post-genomic era (reviewed in ref.). Based on (i) similar RNA affinities of U2AF65 and U2AF651,2L, (ii) indistinguishable conformations among four U2AF651,2L structures in two different crystal packing arrangements and (iii) penalties of structure-guided mutations in RNA binding and splicing assays, we suggest that the extended inter-RRM regions of the U2AF651,2L structures underlie cognate Py-tract recognition by the full-length U2AF65 protein. Further research will be needed to understand the roles of SF1 and U2AF35 subunits in the conformational equilibria underlying U2AF65 association with Py tracts. Moreover, structural differences among U2AF65 homologues and paralogues may regulate splice site selection. Ultimately, these guidelines will assist the identification of 3′ splice sites and the relationship of disease-causing mutations to penalties for U2AF65 association. Methods Protein expression and purification For crystallization and RNA-binding experiments, human U2AF651,2L (residues 141–342 of NCBI RefSeq NP_009210) was expressed in Escherichia coli strain BL21 Rosetta-2 as a GST-fusion protein in the vector pGEX6P-2 and purified by glutathione affinity, followed by anion exchange and gel filtration chromatography. The GST-tagged protein was bound to a GSTrap column (GE Healthcare) in 1 M NaCl, 25 mM HEPES, pH 7.4 and eluted using 150 mM NaCl, 100 mM Tris, pH 8 containing 10 mM glutathione. The GST tag was cleaved from the protein by treatment with PreScission Protease during dialysis against a buffer containing 100 mM NaCl, 25 mM HEPES, pH 8, 5% (v/v) glycerol, 5 mM DTT, 0.25 mM EDTA and 0.1 mM PMSF. Cleaved GST was separated from the U2AF651,2L by subtractive glutathione affinity chromatography in 100 mM NaCl, 25 mM Tris, pH 8, 0.2 mM TCEP followed by subtractive anion-exchange chromatography with a HiTrap Q column (GE Healthcare). The final purification step was size-exclusion chromatography on a Superdex-75 prep-grade column (GE Healthcare) that had been previously equilibrated with 100 mM NaCl, 15 mM HEPES, pH 6.8, 0.2 mM tris(2-carboxy-ethyl)phosphine (TCEP). The purified U2AF651,2L was concentrated using a Vivaspin 15 R (Sartorius) centrifugal concentrator with 10 kDa MWCO, and the protein concentration was estimated using the calculated extinction coefficient of 8,940 M−1cm−1 and absorbance at 280 nm. Shorter constructs (U2AF651,2, residues 148–336; dU2AF651,2, residues 148–237, 258–336; dU2AF651,2L, residues 141–237, 258–342) (Fig. 1a) and individual U2AF651,2L Q147A, R227A, V254P mutants used for RNA-binding experiments were purified similarly. For comparative RNA-binding experiments, full-length human U2AF65 (residues 1–475) and the U2AF35-UHM (U2AF homology motif; residues 43–146, NCBI RefSeq NP_006749) initially were expressed and purified separately as GST fusion proteins. Following GST cleavage and ion-exchange chromatography (SP-HiTrap and Q-HiTrap, respectively), U2AF65 was combined with slight excess U2AF35-UHM (in stoichiometric ratio of 1:1.2) and dialysed overnight. The final U2AF heterodimer was purified by size-exclusion chromatography using a Superdex-200 prep-grade column (GE Healthcare) pre-equilibrated with 150 mM NaCl, 25 mM HEPES, pH 6.8, 0.2 mM TCEP. Representative purified U2AF651,2L and U2AF65–U2AF35-UHM proteins are shown in Supplementary Fig. 1a. Oligonucleotide preparation High-performance liquid chromatography-purified oligonucleotides (sequences shown in Supplementary Fig. 2a) were purchased for crystallization (Integrated DNA Technologies, Inc.). The lyophilized oligonucleotides were diluted in gel filtration buffer for crystallization experiments. The 5′-fluorescein (Fl)-labelled RNAs (AdML: 5′-Fl-CCCUUUUUUUUCC-3′, Py tract of the AdML splicing substrate; 5′-4rU: 5′-Fl-CCUUUUCCCCCCC-3′; 3′-4rU: 5′-Fl-CCCCCCCUUUUCC-3′) for RNA-binding experiments (Dharmacon Research, Inc., Thermo Scientific) was deprotected according to the manufacturer's protocol, vacuum dried and resuspended in nuclease-free water. RNA and RNA–DNA concentrations were calculated using the calculated molar extinction coefficients and absorbance at 260 nm. Fluorescence anisotropy RNA-binding experiments For RNA-binding experiments, purified proteins and RNA were diluted separately \u003e100-fold in binding buffer (100 mM NaCl, 15 mM HEPES, pH 6.8, 0.2 mM TCEP, 0.1 U μl−1 Superase-In (Ambion Life Technologies)). The final RNA concentration in the cuvette was 30 nM. Volume changes during addition of the protein were \u003c10% to minimize dilution effects. The fluorescence anisotropy changes during titration were measured using a FluoroMax-3 spectrophotometer temperature controlled by a circulating water bath at 23 °C. Samples were excited at 490 nm and emission intensities recorded at 520 nm with a slit width of 5 nm. The titrations were repeated three times in succession. Each titration was fit with Graphpad Prism v4.0 to obtain the apparent equilibrium dissociation constant (KD). The apparent equilibrium affinities (KA) are the reciprocal of the KD. The average KD's or KA's and s.e.m. among the three replicates were calculated using Excel and are reported in Figs 3 and 4; Supplementary Figs 1 and 4. The P values from a two-tailed unpaired t-test were calculated using Graphpad Prism v4.0. Transfection, immunoblotting and RT-PCR analyses For transfection experiments, the full-length human U2AF65 cDNA in pCMV6-XL5 (Origene Tech. Inc., clone ID BC008740) was used (WT U2AF65) and in parallel mutated to encode the Q147A/R227A/V254P triple-mutant protein (Mut U2AF65). The pyPY minigene was a gift from M. Carmo-Fonseca (University of Lisbon, Portugal). HEK293T cells (kindly provided by Dr Lata Balakrishnan, originally purchased from ATCC, cat. no. CRL3216) were seeded into 12-well plates (2–4 × 105 cells per well) and grown as monolayers in MEM (Gibco Life Technologies) supplemented with 10% (v/v) of heat-inactivated fetal bovine serum, 1% (v/v) L-glutamine and 1% (v/v) penicillin–streptomycin. After 1 day, the cells were transiently transfected with either 0.5 μg of pyPY plasmid or a mixture of 0.5 μg of U2AF65 variant and 0.5 μg of pyPY plasmid per well using appropriately adjusted Lipofectamine 2000 (Invitrogen Life Technologies) ratio according to the manufacturer's instructions. For immunoblots of WT U2AF65 and Mut U2AF65 expression levels (Supplementary Fig. 5a), transfected or control cells were lysed in radioimmunoprecipitation assay buffer with proteinase and kinase inhibitors. Total protein (20 μg) was separated by SDS–PAGE, and transferred onto polyvinylidene difluoride membranes (Millipore Corp., Billerica, MA, USA) and immunoblotted using mouse monoclonal antibodies directed against U2AF65 (ref.) (MC3, cat. no. U4758 Sigma-Aldrich at 1:500 dilution) or as a control for comparison, GAPDH (glyceraldehyde-3-phosphate dehydrogenase; monoclonal clone 71.1, cat. no. G8795 Sigma-Aldrich at 1:5,000 dilution). Immunoblots were developed using anti-mouse horseradish peroxidase-conjugates (cat. no. U4758 Sigma-Aldrich, Co. at 1:2,500 or 1:10,000 dilutions for GAPDH and U2AF65, respectively) and detected using SuperSignal WestPico chemi-luminescent substrate (Pierce Thermo Scientific Inc.). Blots were imaged using a IS4000MM system (Carestream, Rochester, NY, USA). For size analysis, fluorescent images of the BioRad Precision Plus Dual Color Standards were overlaid directly. For reverse transcription PCR (RT-PCR), the total RNA was isolated 2 days post transfection using the Cells-to-cDNA II kit (Ambion Life Technologies). The RT-PCR reaction comprised 35 cycles (94 °C per 60 s—60 °C per 50 s—72 °C per 60 s) with forward (5′-TGAGGGGAGGTGAATGAGGAG-3′) and reverse (5′-TCCACTGGAAAGACCGCGAAG-3′) primers for the pyPY product or forward (5′-CATGTTCGTCATGGGTGTGAACCA-3′) and reverse (5′-ATGGCATGGACTGTGGTCATGAGT-3′) primers for a GAPDH control. The RT-PCR products were separated by 2% agarose gel electrophoresis and stained with ethidium bromide. The percentages of splice site use were calculated from the background corrected intensities I using the formula 100% × I(py)/[I(py)+I(PY)+I(unspliced)] for py spliced (Fig. 5b,c) or 100% × I(PY)/[I(py)+I(PY)+I(unspliced)] for PY spliced (Supplementary Fig. 5b). The band intensities of four independent biological replicates were measured using ImageQuant software. Crystallization, data collection and structure determination Before crystallization, the purified U2AF651,2L and given oligonucleotide were mixed to achieve respective final concentrations of 1.0 and 1.1 mM and incubated on ice for 20–30 min. For each oligonucleotide, sparse matrix screens of the Jancarik and Kim Crystal Screen(in hanging drop format; Hampton Research, Corp.) and JCSG-Plus (in sitting drop format; Molecular Dimensions) were used to identify initial crystallization conditions, which were obtained from the latter screen and further optimized in hanging drop format. In optimized crystallization experiments, a mixture of sample and reservoir solution (1.2:1 μl) was equilibrated against 700 μl reservoir solution at 4 °C. The oligonucleotide sequences were optimized and the structures were determined as follows: in addition to the previously characterized dU2AF651,2-binding sites for seven nucleotides, the new terminal residues of the U2AF651,2L construct were presumed to contact an additional nucleotide and the crystal packing of a central nucleotide between the RRM1/RRM2 of dU2AF651,2 was presumed to represent one nucleotide. Also considering the known proclivity for deoxy(d)U to co-crystallize with dU2AF651,2 (ref.) and for 5-bromo-dU (5BrdU) to bind a given site of dU2AF651,2 (ref.), we initially designed two 9-mer oligonucleotides (5′-ribose (r)UrUrUrUrU(5BrdU)dUrUrU and 5′-rUrUrUdUdU(5BrdU)dUrUrU) and screened for co-crystallization with U2AF651,2L. The former oligonucleotide failed to produce crystals in these screens. The latter oligonucleotide comprising central dU nucleotides produced diffracting crystals, which were frozen directly from a reservoir comprising 100 mM phosphate–citrate buffer pH 4.2, 40% Peg 300. The structure determined by molecular replacement using Phenix with a data set collected at beamline (BL) 12-2 of the Stanford Synchrotron Radiation Lightsource (SSRL; Menlo Park, CA, USA) (Table 1). The search models comprising each of the individual RRMs bound to two nucleotides were derived from the dU2AF651,2 structure (PDB ID 2G4B) (translation function Z-score equivalent 12.9, log-likelihood gain 528). For comparison, searches with the NMR structure (PDB ID 2YH1) as a search model failed to find a solution. The initial structure revealed a greater number of central nucleotide-binding sites than expected. The oligonucleotide binding register had slipped to place the BrdU in the preferred site, leave the 5′ terminal-binding sites empty, and the terminal nucleotide unbound and disordered. Subsequent oligonucleotides were designed to place BrdU in the preferred site, fill the unoccupied 5′ terminal sites, capture rU at the central sites, and compare rC at the terminal site. The U2AF651,2L protein co-crystals with oligonucleotide 5′-phosphorylated (P)-rUrUdUdUrUdU(BrdU)dU were obtained using a reservoir of 200 mM LiCl, 100 mM sodium citrate pH 4.0, 8% (w/v) polyethylene glycol (PEG) 6,000, 10% (v/v) PEG 300, 10% (v/v) dioxane with 0.1 μl of N,N-bis[3-(D-gluconamido)propyl]deoxy-cholamide (deoxy-BigCHAP) (14 mM) added to the hanging drop and cryoprotected by sequential layering with reservoir solution supplemented with increasing PEG 300 to a final concentration of 26%. Co-crystals with either 5′-(P)rUrUdUrUrU(BrdU)dUdU or 5′-(P)rUrUrUdUrUrU(BrdU)dUrC were obtained from 1 M succinate, 100 mM HEPES, pH 7.0, 1–3% (w/v) PEG monomethylether 2,000. The former was cryoprotected by coating with a 1:1 (v/v) mixture of silicon oil and Paratone-N and the latter by sequential transfer to 21% (v/v) glycerol. Data sets for flash-cooled crystals were collected at 100 K using remote access to SSRL BL12-2. Structures were determined by molecular replacement using the initial U2AF651,2L/rUrUrUdUdU(BrdU)dUrUrU structure as a search model. Consistent sets of free-R reflections were maintained (6% of the total reflections). Models were built using COOT and refined with PHENIX. No non-glycine/non-proline residues were found in the disallowed regions of the Ramachandran plots. Clash scores and Molprobity scores calculated using the program Molprobity were above average. Structure illustrations were prepared using PYMOL. Crystallographic data and refinement statistics are given in Table 1. Sample preparation for single-molecule FRET The U2AF651,2LFRET construct used for smFRET comprises the six histidine and T7 tags from the pET28a vector (Merck), a GGGS linker and U2AF65 residues 113–343. The single cysteine of human U2AF65 was replaced by alanine (C305A), which is a natural amino-acid variation among U2AF65 homologues. Single A181C and Q324C mutations were introduced in each RRM for fluorophore attachment at residues that were carefully selected to meet experimental criteria described in the Results. The U2AF651,2LFRET was purified by the same method as described above for U2AF651,2L protein and binds RNA with similar affinity as U2AF651,2L (Supplementary Fig. 6a). Before labelling, the purified U2AF651,2LFRET protein was incubated with 10 mM DTT on ice for 30 min and then buffer exchanged into Labelling Buffer (100 mM NaCl, 25 mM HEPES pH 7.0, 5 mM EDTA, 0.5 mM tris(2-carboxy-ethyl)phosphine (TCEP)) using Zeba Spin Desalting Columns 7K MWCO (Pierce, ThermoFisher Scientific). To initiate the labelling reaction, 4 μl each of cyanine (Cy)3-Maleimide and Cy5-Maleimide (Combinix, Inc.) stock solutions (10 mM in DMSO) were pre-mixed (total volume 8 μl) and then added to 200 μl of 20 μM protein (final 20:1 molar ratio of dye:protein). The labelling reaction was incubated at room temperature in the dark for 2 h and then quenched by the addition of 10 mM DTT. The labelled protein was separated from excess dye using a Zeba Spin Desalting Column followed by size exclusion chromatography using a pre-packed Superdex-75 10/300 GL (GE Healthcare) column in Labelling Buffer. Our previous experience of conjugating cysteines with maleimide derivatives of fluorophores and suggests that nonspecific modification of aminogroups of proteins with fluorescent dyes under the employed experimental conditions is negligible. Consistent with specific labelling of A181C and Q324C, the labelling efficiencies were ∼60% each for Cy3 and Cy5 as estimated using the dye extinction coefficients (ɛCy3=150,000 M−1 cm−1 at 550 nm, ɛCy5=170,000 M−1 cm−1 at 650 nm) and the calculated extinction coefficient of the U2AF651,2LFRET protein (ɛprot=8,940 M−1 cm−1 at 280 nm), and correcting for the absorbance (A) of the dyes at 280 nm (GE Healthcare, Amersham CyDye Maleimide product booklet): For smFRET experiments with a ‘strong', homogeneous Py tract, we used the prototypical AdML sequence (5′-CCUUUUUUUUCC-3′). To investigate the inter-RRM separation in the presence of a ‘weak' Py tract interrupted by purines, we compared the U2AF651,2L affinity for a purine-interrupted Py tract comprising an rUrUrUrUrU tract that is expected to bind U2AF65 RRM2/inter-RRM linker, a central rArA and an rUrUrUrCrC tract that is expected to bind RRM1. The tandem purines represent a compromise between significant inhibition of U2AF65 binding by longer A interruptions and an approximately five-fold penalty for the rArA mutation in the AdML Py tract (Supplementary Fig. 6b,c). To maintain avidity and provide flanking phosphoryl groups in case of inter-RRM adjustment, we included the 5′-C and 3′-A of parent AdML sequence, which are respective low-affinity nucleotides for binding RRM2 and RRM1 (ref.), in the final rArA-interrupted RNA oligonucleotide (5′-rCrUrUrUrUrUrArArUrUrUrCrCrA-3′). For the reversed immobilization of RNA via a complementary biotinyl-DNA primer experiment, the AdML Py-tract RNA was extended to include the DNA counterpart of downstream AdML intron/exon sequences that were complementary to the biotinyl-DNA primer. To increase separation from the slide surface, a hexaethylene glycol linker (18PEG) was inserted between the AdML Py-tract RNA and the tethered DNA duplex. The tethered oligonucleotide sequences included: 5′-rCrCrUrUrUrUrUrUrUrUrCrC/18PEG/dAdCdAdGdCdTdCdGdCdG-dGdTdTdGdAdGdGdAdCdAdA-3′ annealed to 5′-biotinyl-dTdTdGdTdCdCdTdCdAdA-dCdCdGdCdGdAdGdCdTdGdT-3' (purchased with high-performance liquid chromatography purification from Integrated DNA Technologies). Single-molecule FRET data acquisition and analysis The smFRET measurements were carried out at room temperature in 50 mM HEPES, pH 7.4, 100 mM NaCl. The imaging buffer also contained an oxygen-scavenging system (0.8 mg ml−1 glucose oxidase, 0.625% glucose, 0.02 mg ml−1 catalase), 1.5 mM Trolox (used to eliminate Cy5 blinking) and 6 mM β-mercaptoethanol. The sample chamber was assembled from quartz microscope slides and glass cover slips coated with a mixture of m-PEG and biotin-PEG and pre-treated with neutravidin (0.2 mg ml−1). Surface tethering of doubly labelled U2AF651,2LFRET(Cy3/Cy5) via its His-tag (Fig. 6c–f,i,j; Supplementary Fig. 7a,b) was achieved by pre-incubating the sample chamber with 50 nM biotinyl-NTA resin (Biotin-X NTA, Biotium), pre-loaded with three-fold excess NiSO4) for 20 min before addition of 5 nM U2AF651,2LFRET(Cy3/Cy5). After 10 min, unbound sample was removed by washing the sample chamber with imaging buffer. The AdML RNA ligand was added to the imaging buffer at a concentration of 5 μM (100-fold higher than the measured KD value), whereas the rArA-interrupted RNA was added at a concentration of 10 μM. Alternatively, to detect binding of doubly labelled U2AF651,2LFRET(Cy3/Cy5) to surface-tethered RNA ligand (Fig. 6g,h Supplementary Fig. 7c–g), 10 nM AdML RNA (pre-annealed to biotinyl-DNA primer) was incubated in the neutravidin-treated sample chamber for 20 min, and 1 nM U2AF651,2LFRET(Cy3/Cy5) was then added to the imaging buffer. Single-molecule FRET measurements were taken as previously described. An Olympus IX71 inverted microscope, equipped with a UPlanApo 60x/1.20w objective lens, a 532 nm laser (Spectra-Physics) for excitation of Cy3 dyes, and a 642 nm laser (Spectra-Physics) for excitation of Cy5 dyes was used. Total internal reflection (TIR) was obtained by a quartz prism (ESKMA Optics). Fluorescence emission was split into Cy3 and Cy5 fluorescence using a dual view imaging system DV2 (Photometrics) equipped with a 630 nm dichroic mirror and recorded via an Andor iXon+ EMCCD camera. Movies were recorded using the Single software (downloaded from Prof. Taekjip Ha's laboratory website at the University of Illinois at Urbana-Champaign, physics.illinois.edu/cplc/software), with the exposure time set at 100 ms. We typically took up to five 5-minute-long movies while imaging different sections of the slide for each sample. Before each measurement, we checked for non-specific binding by adding doubly-labeled U2Fret to the slide in the absence of neutravidin and imaging the slide. Non-specific binding was virtually absent. Collected data sets were processed with IDL and Matlab softwares, using scripts downloaded from a freely available source: physics.illinois.edu/cplc/software. Apparent FRET efficiencies (Eapp) were calculated from the emission intensities of donor (ICy3) and acceptor (ICy5) as follows: Eapp=ICy5/(ICy5+ICy3). The FRET distribution histograms were built from traces that showed single-step photobleaching in both Cy3 and Cy5 signals using a Matlab script generously provided by Prof. Peter Cornish (University of Missouri, Columbia). Anti-correlated changes in donor and acceptor intensities with constant sum of intensities indicated the presence of an energy transfer in single molecules labelled with one donor and one acceptor dye. All histograms were smoothed with a five-point window and plotted using Origin software (Origin Lab Co). Idealization of FRET trajectories was done using the hidden Markov model algorithms via HaMMy software (http://bio.physics.illinois.edu/HaMMy.asp). Transition density plots were generated from transitions detected in idealized FRET trajectories obtained by HaMMy fit of raw FRET traces via Matlab. Frequency of transitions from starting FRET efficiency value (x-axis) to ending FRET efficiency value (y-axis) was represented by a heat map. The range of FRET efficiencies from 0 to 1 was separated in 200 bins. The resulting heat map was normalized to the most populated bin in the plot; the lower- and upper-bound thresholds were set to 20% and 100% of the most populated bin, respectively. The surface contour plots were generated as follows: the individual single-molecule FRET traces (for example, Fig. 6g of the main text and Supplementary Fig. 7e,f) were post synchronized at the first time point showing non-zero (\u003e0.15) FRET efficiency, corresponding to binding. The time range (x-axis, 0–10 s) was separated into 100 bins. The FRET efficiency range (y-axis, 0–1 FRET) was separated into 100 bins. A heat map is used to represent the frequency of sampling of each FRET state over time; frequency in each bin was normalized to the most populated bin in the plot with lower- and upper-bound thresholds set at 10% and 80% of the most populated bin, respectively. Additional information Accession codes: Coordinates and structure factors have been deposited in the Protein Data Bank with accession codes 5EV1, 5EV2, 5EV3 and 5EV4 for respective U2AF651,2L-oligonucleotide structures (i)–(iv). How to cite this article: Agrawal, A. A. et al. An extended U2AF65–RNA-binding domain recognizes the 3′ splice site signal. Nat. Commun. 7:10950 doi: 10.1038/ncomms10950 (2016). Supplementary Material Alternative isoform regulation in human tissue transcriptomes  Pre-mRNA splicing in disease and therapeutics Acquired mutations that affect pre-mRNA splicing in hematologic malignancies and solid tumors Insertional mutagenesis in zebrafish rapidly identifies genes essential for early vertebrate development A factor, U2AF, is required for U2 snRNP binding and splicing complex assembly The splicing factor BBP interacts specifically with the pre-mRNA branchpoint sequence UACUAAC A cooperative interaction between U2AF65 and mBBP/SF1 facilitates branchpoint region recognition Functional recognition of the 3' splice site AG by the splicing factor U2AF35 Both subunits of U2AF recognize the 3' splice site in Caenorhabditis elegans Inhibition of msl-2 splicing by Sex-lethal reveals interaction between U2AF35 and the 3' splice site AG Distinct binding specificities and functions of higher eukaryotic polypyrimidine tract-binding proteins Cloning and domain structure of the mammalian splicing factor U2AF U2AF65 adapts to diverse pre-mRNA splice sites through conformational selection of specific and promiscuous RNA recognition motifs Structural basis of polypyrimidine tract recognition by the essential pre-mRNA splicing factor, U2AF65 Multi-domain conformational selection underlies pre-mRNA splicing regulation by U2AF Structuring of the 3' splice site by U2AF65 Interaction of U2AF65 RS region with pre-mRNA branch point and promotion of base pairing with U2 snRNA A pathway of sequential arginine-serine-rich domain-splicing signal interactions during mammalian spliceosome assembly Structure-guided U2AF65 variant improves recognition and splicing of a defective pre-mRNA Protein-protein interaction and quaternary structure Solution conformation and thermodynamic characteristics of RNA binding by the splicing factor U2AF65 Solution structures of the first and second RNA-binding domains of human U2 small nuclear ribonucleoprotein particle auxiliary factor (U2AF65) In vivo requirement of the small subunit of U2AF for recognition of a weak 3' splice site A broad range of conformations contribute to the solution ensemble of the essential splicing factor U2AF65 Transient electrostatic interactions dominate the conformational equilibrium sampled by multidomain splicing factor U2AF65: a combined NMR and SAXS study Fluorescence resonance energy transfer (FRET) and competing processes in donor-acceptor substituted DNA strands: a comparative study of ensemble and single molecule data A practical guide to single-molecule FRET Landscape of genetic lesions in 944 patients with myelodysplastic syndromes Frequent pathway mutations of splicing machinery in myelodysplasia A novel peptide recognition mode revealed by the X-ray structure of a core U2AF35/U2AF65 heterodimer Induced fit, conformational selection and independent dynamic segments: an extended view of binding events Conformational selection or induced fit: a flux description of reaction mechanism Illuminating the mechanistic roles of enzyme conformational dynamics Intrinsic motions along an enzymatic reaction trajectory A role for both conformational selection and induced fit in ligand binding by the LAO protein Conformational selection and induced fit mechanism underlie specificity in noncovalent interactions with ubiquitin A single-molecule dissection of ligand binding to a protein with intrinsic dynamics Mg(2+) shifts ligand-mediated folding of a riboswitch from induced-fit to conformational selection Splicing regulation: from a parts list of regulatory elements to an integrated splicing code Revised UV extinction coefficients for nucleoside-5'-monophosphates and unpaired DNA and RNA Targeting of U2AF65 to sites of active splicing in the nucleus Sparse matrix sampling: a screening method for crystallization of proteins Crystallization and preliminary X-ray analysis of U2AF65 variant in complex with a polypyrimidine tract analogue by use of protein engineering PHENIX: a comprehensive Python-based system for macromolecular structure solution Coot: model-building tools for molecular graphics MOLPROBITY: structure validation and all-atom contact analysis for nucleic acids and their complexes  Following movement of domain IV of elongation factor G during ribosomal translocation Analysis of single-molecule FRET trajectories using hidden Markov modeling Author contributions A.A.A. performed crystallization, refinement, molecular biology and most RNA-binding experiments. E.S. performed smFRET experiments. R.C. labelled protein and S.H. completed a subset of RNA-binding experiments. C.L.K. cryoprotected crystals, collected crystallographic data and built structures. J.L.J. performed molecular replacement and completed structure refinements. M.R.G. and C.L.K. conceived the study. C.L.K. and D.N.E. designed the experiments. C.L.K., D.N.E. and E.S. wrote the paper with input from J.L.J. and A.A.A. The intact U2AF65 RRM1/RRM2-containing domain and flanking residues are required for binding contiguous Py tracts. (a) Domain organization of full-length (fl) U2AF65 and constructs used for RNA binding and structural experiments. The N- and C-terminal residue numbers are indicated. An internal deletion (d, Δ) of residues 238–257 removes a portion of the inter-RRM linker from the dU2AF651,2 and dU2AF651,2L constructs. (b) Comparison of the apparent equilibrium affinities of various U2AF65 constructs for binding the prototypical AdML Py tract (5′-CCCUUUUUUUUCC-3′). The flU2AF65 protein includes a heterodimerization domain of the U2AF35 subunit to promote solubility and folding. The apparent equilibrium dissociation constants (KD) for binding the AdML 13mer are as follows: flU2AF65, 30±3 nM; U2AF651,2L, 35±6 nM; U2AF651,2, 3,600±300 nM. (c) Comparison of the RNA sequence specificities of flU2AF65 and U2AF651,2L constructs binding C-rich Py tracts with 4U's embedded in either the 5′- (light grey fill) or 3′- (dark grey fill) regions. The KD's for binding 5′-CCUUUUCCCCCCC-3′ are: flU2AF65, 41±2 nM; U2AF651,2L, 31±3 nM. The KD's for binding 5′-CCCCCCCUUUUCC-3′ are: flU2AF65, 414±12 nM; U2AF651,2L, 417±10 nM. Bar graphs are hatched to match the constructs shown in a. The average apparent equilibrium affinity (KA) and s.e.m. for three independent titrations are plotted. The P values from two-tailed unpaired t-tests with Welch's correction are indicated as follows: **P\u003c0.01; NS, not significant, P\u003e0.05. The purified protein and average fitted fluorescence anisotropy RNA-binding curves are shown in Supplementary Fig. 1. RRM, RNA recognition motif; RS, arginine-serine rich; UHM, U2AF homology motif; ULM, U2AF ligand motif. Structures of U2AF651,2L recognizing a contiguous Py tract. (a) Alignment of oligonucleotide sequences that were co-crystallized in the indicated U2AF651,2L structures. The regions of RRM1, RRM2 and linker contacts are indicated above by respective black and blue arrows from N- to C-terminus. For clarity, we consistently number the U2AF651,2L nucleotide-binding sites from one to nine, although in some cases the co-crystallized oligonucleotide comprises eight nucleotides and as such leaves the first binding site empty. The prior dU2AF651,2 nucleotide-binding sites are given in parentheses (site 4' interacts with dU2AF65 RRM1 and RRM2 by crystallographic symmetry). Italics, disordered in the structure. (b) Stereo views of a ‘kicked' 2|Fo|−|Fc| electron density map contoured at 1σ for the inter-RRM linker, N- and C-terminal residues (blue) or bound oligonucleotide of a representative U2AF651,2L structure (structure iv, bound to 5′-(P)rUrUrUdUrUrU(BrdU)dUrC) (magenta). (c) Cartoon diagram of this structure. Crystallographic statistics are given in Table 1 and the overall conformations of U2AF651,2L and prior dU2AF651,2/U2AF651,2 structures are compared in Supplementary Fig. 2. BrdU, 5-bromo-deoxy-uridine; d, deoxy-ribose; P-, 5′-phosphorylation; r, ribose. Representative views of the U2AF651,2L interactions with each new nucleotide of the bound Py tract. New residues of the U2AF651,2L structures are coloured a darker shade of blue, apart from residues that were tested by site-directed mutagenesis, which are coloured yellow. The nucleotide-binding sites of the U2AF651,2L and prior dU2AF651,2 structure are compared in Supplementary Fig. 3a–h. The first and seventh U2AF651,2L-binding sites are unchanged from the prior dU2AF651,2–RNA structure and are portrayed in Supplementary Fig. 3a,f. The four U2AF651,2L structures are similar with the exception of pH-dependent variations at the ninth site that are detailed in Supplementary Fig. 3i,j. The representative U2AF651,2L structure shown has the highest resolution and/or ribose nucleotide at the given site: (a) rU2 of structure iv; (b) rU3 of structure iii; (c) rU4 of structure i; (d) rU5 of structure iii; (e) rU6 of structure ii; (f) dU8 of structure iii; (g) dU9 of structure iii; (h) rC9 of structure iv. (i) Bar graph of apparent equilibrium affinities (KA) of the wild type (blue) and the indicated mutant (yellow) U2AF651,2L proteins binding the AdML Py tract (5′-CCCUUUUUUUUCC-3′). The apparent equilibrium dissociation constants (KD) of the U2AF651,2L mutant proteins are: wild type (WT), 35±6 nM; R227A, 166±2 nM; V254P, 137±10 nM; Q147A, 171±21 nM. The average KA and s.e.m. for three independent titrations are plotted. The P values from two-tailed unpaired t-tests with Welch's correction are indicated as follows: **P\u003c0.01; *P\u003c0.05; NS, not significant, P\u003e0.05. The average fitted fluorescence anisotropy RNA-binding curves are shown in Supplementary Fig. 4a–c. The U2AF65 linker/RRM and inter-RRM interactions. (a) Contacts of the U2AF65 inter-RRM linker with the RRMs. A semi-transparent space-filling surface is shown for the RRM1 (green) and RRM2 (light blue). Residues V249, V250, V254 (yellow) are mutated to V249G/V250G/V254G in the 3Gly mutant; residues S251, T252, V253, P255 (red) along with V254 are mutated to S251G/T252G/V253G/V254G/P255G in the 5Gly mutant or to S251N/T252L/V253A/V254L/P255A in the NLALA mutant; residues M144, L235, M238, V244, V246 (orange) along with V249, V250, S251, T252, V253, V254, P255 are mutated to M144G/L235G/M238G/V244G/V246G/V249G/ V250G/S251G/T252G/V253G/V254G/P255G in the 12Gly mutant. Other linker residues are coloured either dark blue for new residues in the U2AF651,2L structure or light blue for the remaining inter-RRM residues. The central panel shows an overall view with stick diagrams for mutated residues; boxed regions are expanded to show the C-terminal (bottom left) and central linker regions (top) at the inter-RRM interfaces, and N-terminal linker region contacts with RRM1 (bottom right). (b) Bar graph of apparent equilibrium affinities (KA) for the AdML Py tract (5′-CCCUUUUUUUUCC-3′) of the wild-type (blue) U2AF651,2L protein compared with mutations of the residues shown in a: 3Gly (yellow), 5Gly (red), NLALA (hatched red), 12Gly (orange) and the linker deletions dU2AF651,2 in the minimal RRM1–RRM2 region (residues 148–237, 258–336) or dU2AF651,2L (residues 141–237, 258–342). The apparent equilibrium dissociation constants (KD) of the U2AF651,2L mutant proteins are: wild type (WT), 35±6 nM; 3Gly, 47±4 nM; 5Gly, 61±3 nM; 12Gly, 88±21 nM; NLALA, 45±3 nM; dU2AF651,2L, 123±5 nM; dU2AF651,2, 5000±100 nM; 3Mut, 5630±70 nM. The average KA and s.e.m. for three independent titrations are plotted. The P values from two-tailed unpaired t-tests with Welch's correction are indicated as follows: **P\u003c0.01; *P\u003c0.05; NS, not significant, P\u003e0.05. The fitted fluorescence anisotropy RNA-binding curves are shown in Supplementary Fig. 4d–j. (c) Close view of the U2AF65 RRM1/RRM2 interface following a two-fold rotation about the x-axis relative to a. U2AF65 inter-domain residues are important for splicing a representative pre-mRNA substrate in human cells. (a) Schematic diagram of the pyPY reporter minigene construct comprising two alternative splice sites preceded by either the weak IgM Py tract (py) or the strong AdML Py tract (PY) (sequences inset). (b) Representative RT-PCR of pyPY transcripts from HEK293T cells co-transfected with constructs encoding the pyPY minigene and either wild-type (WT) U2AF65 or a triple U2AF65 mutant (3Mut) of Q147A, R227A and V254P residues. (c) A bar graph of the average percentage of the py-spliced mRNA relative to total detected pyPY transcripts (spliced and unspliced) for the corresponding gel lanes (black, no U2AF65 added; white, WT U2AF65; grey, 3Mut U2AF65). The average percentages and s.d.'s are given among four independent biological replicates. ****P\u003c0.0001 for two-tailed unpaired t-test with Welch's correction. Protein overexpression and qRT-PCR results are shown in Supplementary Fig. 5. RNA binding stabilizes the side-by-side conformation of U2AF65 RRMs. (a,b) Views of FRET pairs chosen to follow the relative movement of RRM1 and RRM2 on the crystal structure of ‘side-by-side' U2AF651,2L RRMs bound to a Py-tract oligonucleotide (a, representative structure iv) or ‘closed' NMR/PRE-based model of U2AF651,2 (b, PDB ID 2YH0) in identical orientations of RRM2. The U2AF651,2LFRET proteins were doubly labelled at A181C/Q324C such that a mixture of Cy3/Cy5 fluorophores are expected to be present at each site. (c–f,i,j) The U2AF651,2LFRET(Cy3/Cy5) protein was immobilized on the microscope slide via biotin-NTA/Ni+2 (orange line) on a neutravidin (black X)-biotin-PEG (orange triangle)-treated surface and imaged either in the absence of ligands (c,d), in the presence of 5 μM AdML Py-tract RNA (5′-CCUUUUUUUUCC-3′) (e,f), or in the presence of 10 μM adenosine-interrupted variant RNA (5′-CUUUUUAAUUUCCA-3′) (i,j). In g and h, the immobilization protocol was reversed. The untethered U2AF651,2LFRET(Cy3/Cy5) protein (1 nM) was added to AdML RNA–polyethylene-glycol-linker–DNA oligonucleotide (10 nM), which was immobilized on the microscope slide by annealing with a complementary biotinyl-DNA oligonucleotide (black vertical line). Typical single-molecule FRET traces (c,e,g,i) show fluorescence intensities from Cy3 (green) and Cy5 (red) and the calculated apparent FRET efficiency (blue). Additional traces for untethered, RNA-bound U2AF651,2LFRET(Cy3/Cy5) are shown in Supplementary Fig. 7c,d. Histograms (d,f,h,j) show the distribution of FRET values in RNA-free, slide-tethered U2AF651,2LFRET(Cy3/Cy5) (d); AdML RNA-bound, slide-tethered U2AF651,2LFRET(Cy3/Cy5) (f); AdML RNA-bound, untethered U2AF651,2LFRET(Cy3/Cy5) (h) and adenosine-interrupted RNA-bound, slide-tethered U2AF651,2LFRET(Cy3/Cy5) (j). N is the number of single-molecule traces compiled for each histogram. Schematic models of U2AF65 recognizing the Py tract. (a) Diagram of the U2AF65, SF1 and U2AF35 splicing factors bound to the consensus elements of the 3′ splice site. A surface representation of U2AF651,2L is shown bound to nine nucleotides (nt); the relative distances and juxtaposition of the branch point sequence (BPS) and consensus AG dinucleotide at the 3′ splice site are unknown. MDS-relevant mutated residues of U2AF65 are shown as yellow spheres (L187 and M144). (b) Following binding to the Py-tract RNA, a conformation corresponding to high FRET and consistent with the ‘closed', back-to-back apo-U2AF65 model resulting from PRE/NMR characterization (PDB ID 2YH0) often transitions to a conformation corresponding to ∼0.45 FRET value, which is consistent with ‘open', side-by-side RRMs such as the U2AF651,2L crystal structures. Alternatively, a conformation of U2AF65 corresponding to ∼0.45 FRET value can directly bind to RNA; RNA binding stabilizes the ‘open', side-by-side conformation and thus shifts the U2AF65 population towards the ∼0.45 FRET value. RRM1, green; RRM2, pale blue; RRM extensions/linker, blue. Crystallographic data and refinement statistics*. Structure\tU2AF651,2L with rUrUrUdUdU(BrdU)dUrUrU\tU2AF651,2L with (P)rUrUdUdUrUdU(BrdU)dU\tU2AF651,2L with (P)rUrUdUrUrU(BrdU)dUdU\tU2AF651,2L with (P)rUrUrUdUrUrU(BrdU)dUrC\t \tData collection\t(i)\t(ii)\t(iii)\t(iv)\t \tSpace group\tC2221\tC2221\tP212121\tP212121\t \tUnit cell (Å) a,b,c\t62.1, 114.2, 59.4\t61.9, 115.1, 59.5\t43.4, 62.2, 77.4\t43.5, 63.4, 77.7\t \tResolution limits (Å)\t32.46–2.04\t32.57–1.86\t38.71–1.50\t38.83–1.57\t \tCompleteness (%)\t95.5 (78.3)\t98.7 (95.9)\t98.2 (69.8)\t98.3 (71.7)\t \tRedundancy\t4.6 (4.1)\t4.3 (4.2)\t6.1 (3.0)\t6.2 (3.1)\t \tI/σ(I)\t21.2 (4.2)\t24.6 (4.6)\t38.0 (6.5)\t42.9 (6.9)\t \tRsym (%)\t3.9 (32.1)\t3.9 (30.3)\t2.4 (14.8)\t2.2 (14.9)\t \tRefinement\t \t No. reflections (work/test)\t12,124/1,055\t17,870/1,456\t31,802/1,996\t28,162/2,000\t \t Rwork/Rfree (%)\t17.3/22.8\t15.1/18.8\t15.3/18.6\t15.4/17.6\t \tNo. atoms\t \t Protein\t2,982\t3,052\t2,986\t2,978\t \t Oligonucleotide\t214\t209\t198\t255\t \t Water\t118\t203\t263\t177\t \tBond r.m.s.d.\t \t Bond lengths (Å)\t0.013\t0.010\t0.008\t0.009\t \t Bond angles (°)\t1.32\t1.1\t1.05\t1.05\t \t\u003cB\u003e factors (Å2)\t \t Protein\t46.4\t27.4\t26.3\t26.7\t \t Oligonucleotide\t61.8\t35.2\t24.5\t30.5\t \t Water\t45.2\t35.2\t30.7\t29.8\t \t All available crystallographic data was used for refinement. *A single crystal was used for each structure. Values from the highest resolution shell are given in parentheses: 2.15–2.04; 1.90–1.86; 1.53–1.50; 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diff --git a/annotated_BioC_JSON/PMC4792962_ann.json b/annotated_BioC_JSON/PMC4792962_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..f8cec9c7f553b6c6cbbce5d21c093a4b509dcd22
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+++ b/annotated_BioC_JSON/PMC4792962_ann.json
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+[{"sourceid":"4792962","sourcedb":"","project":"","target":"","text":"A unified mechanism for proteolysis and autocatalytic activation in the 20S proteasome Biogenesis of the 20S proteasome is tightly regulated. The N-terminal propeptides protecting the active-site threonines are autocatalytically released only on completion of assembly. However, the trigger for the self-activation and the reason for the strict conservation of threonine as the active site nucleophile remain enigmatic. Here we use mutagenesis, X-ray crystallography and biochemical assays to suggest that Lys33 initiates nucleophilic attack of the propeptide by deprotonating the Thr1 hydroxyl group and that both residues together with Asp17 are part of a catalytic triad. Substitution of Thr1 by Cys disrupts the interaction with Lys33 and inactivates the proteasome. Although a Thr1Ser mutant is active, it is less efficient compared with wild type because of the unfavourable orientation of Ser1 towards incoming substrates. This work provides insights into the basic mechanism of proteolysis and propeptide autolysis, as well as the evolutionary pressures that drove the proteasome to become a threonine protease.  The proteasome, an essential molecular machine, is a threonine protease, but the evolution and the components of its proteolytic centre are unclear. Here, the authors use structural biology and biochemistry to investigate the role of proteasome active site residues on maturation and activity. The 20S proteasome core particle (CP) is the key non-lysosomal protease of eukaryotic cells. Its seven different α and seven different β subunits assemble into four heptameric rings that are stacked on each other to form a hollow cylinder. While the inactive α subunits build the two outer rings, the β subunits form the inner rings. Only three out of the seven different β subunits, namely β1, β2 and β5, bear N-terminal proteolytic active centres, and before CP maturation these are protected by propeptides. In the last stage of CP biogenesis, the prosegments are autocatalytically removed through nucleophilic attack by the active site residue Thr1 on the preceding peptide bond involving Gly(-1). Release of the propeptides creates a functionally active CP that cleaves proteins into short peptides. Although the chemical nature of the substrate-binding channel and hence substrate preferences are unique to each of the distinct active β subunits, all active sites employ an identical reaction mechanism to hydrolyse peptide bonds. Nucleophilic attack of Thr1Oγ on the carbonyl carbon atom of the scissile peptide bond creates a first cleavage product and a covalent acyl-enzyme intermediate. Hydrolysis of this complex by the addition of a nucleophilic water molecule regenerates the enzyme and releases the second peptide fragment. The proteasome belongs to the family of N-terminal nucleophilic (Ntn) hydrolases, and the free N-terminal amine group of Thr1 was proposed to deprotonate the Thr1 hydroxyl group to generate a nucleophilic Thr1Oγ for peptide-bond cleavage. This mechanism, however, cannot explain autocatalytic precursor processing because in the immature active sites, Thr1N is part of the peptide bond with Gly(-1), the bond that needs to be hydrolysed. An alternative candidate for deprotonating the Thr1 hydroxyl group is the side chain of Lys33 as it is within hydrogen-bonding distance to Thr1OH (2.7 Å). In principle it could function as the general base during both autocatalytic removal of the propeptide and protein substrate cleavage. Here we provide experimental evidences for this distinct view of the proteasome active-site mechanism. Data from biochemical and structural analyses of proteasome variants with mutations in the β5 propeptide and the active site strongly support the model and deliver novel insights into the structural constraints required for the autocatalytic activation of the proteasome. Furthermore, we determine the advantages of Thr over Cys or Ser as the active-site nucleophile using X-ray crystallography together with activity and inhibition assays. Results Inactivation of proteasome subunits by T1A mutations Proteasome-mediated degradation of cell-cycle regulators and potentially toxic misfolded proteins is required for the viability of eukaryotic cells. Inactivation of the active site Thr1 by mutation to Ala has been used to study substrate specificity and the hierarchy of the proteasome active sites. Yeast strains carrying the single mutations β1-T1A or β2-T1A, or both, are viable, even though one or two of the three distinct catalytic β subunits are disabled and carry remnants of their N-terminal propeptides (Table 1). These results indicate that the β1 and β2 proteolytic activities are not essential for cell survival. By contrast, the T1A mutation in subunit β5 has been reported to be lethal or nearly so. Viability is restored if the β5-T1A subunit has its propeptide (pp) deleted but expressed separately in trans (β5-T1A pp trans), although substantial phenotypic impairment remains (Table 1). Our present crystallographic analysis of the β5-T1A pp trans mutant demonstrates that the mutation per se does not structurally alter the catalytic active site and that the trans-expressed β5 propeptide is not bound in the β5 substrate-binding channel (Supplementary Fig. 1a). The extremely weak growth of the β5-T1A mutant pp cis described by Chen and Hochstrasser compared with the inviability reported by Heinemeyer et al. prompted us to analyse this discrepancy. Sequencing of the plasmids, testing them in both published yeast strain backgrounds and site-directed mutagenesis revealed that the β5-T1A mutant pp cis is viable, but suffers from a marked growth defect that requires extended incubation of 4–5 days for initial colony formation (Table 1 and Supplementary Methods). We also identified an additional point mutation K81R in subunit β5 that was present in the allele used in ref.. This single amino-acid exchange is located at the interface of the subunits α4, β4 and β5 (Supplementary Fig. 1b) and might weakly promote CP assembly by enhancing inter-subunit contacts. The slightly better growth of the β5-T1A-K81R mutant allowed us to solve the crystal structure of a yeast proteasome (yCP) with the β5-T1A mutation, which is discussed in the following section (for details see Supplementary Note 1). Propeptide conformation and triggering of autolysis In the final steps of proteasome biogenesis, the propeptides are autocatalytically cleaved from the mature β-subunit domains. For subunit β1, this process was previously inferred to require that the propeptide residue at position (-2) of the subunit precursor occupies the S1 specificity pocket of the substrate-binding channel formed by amino acid 45 (for details see Supplementary Note 2). Furthermore, it was observed that the prosegment forms an antiparallel β-sheet in the active site, and that Gly(-1) adopts a γ-turn conformation, which by definition is characterized by a hydrogen bond between Leu(-2)O and Thr1NH (ref.). Here we again analysed the β1-T1A mutant crystallographically but in addition determined the structures of the β2-T1A single and β1-T1A-β2-T1A double mutants (Protein Data Bank (PDB) entry codes are provided in Supplementary Table 1). In subunit β1, we found that Gly(-1) indeed forms a sharp turn, which relaxes on prosegment cleavage (Fig. 1a and Supplementary Fig. 2a). However, the γ-turn conformation and the associated hydrogen bond initially proposed is for geometric and chemical reasons inappropriate and would not perfectly position the carbonyl carbon atom of Gly(-1) for nucleophilic attack by Thr1. Regarding the β2 propeptide, Thr(-2) occupies the S1 pocket but is less deeply anchored compared with Leu(-2) in β1, which might be due to the rather large β2-S1 pocket created by Gly45. Thr(-2) positions Gly(-1)O via hydrogen bonding (∼2.8 Å) in a perfect trajectory for the nucleophilic attack by Thr1Oγ (Fig. 1b and Supplementary Fig. 2b). Next, we examined the position of the β5 propeptide in the β5-T1A-K81R mutant. Surprisingly, Gly(-1) is completely extended and forces the histidine side chain at position (-2) to occupy the S2 instead of the S1 pocket, thereby disrupting the antiparallel β-sheet. Nonetheless, the carbonyl carbon of Gly(-1) would be ideally placed for nucleophilic attack by Thr1Oγ (Fig. 1c and Supplementary Fig. 2c,d). As the K81R mutation is located far from the active site (Thr1Cα–Arg81Cα: 24 Å), any influence on propeptide conformation can be excluded. Instead, the plasticity of the β5 S1 pocket caused by the rotational flexibility of Met45 might prevent stable accommodation of His(-2) in the S1 site and thus also promote its immediate release after autolysis. Processing of β-subunit precursors requires deprotonation of Thr1OH; however, the general base initiating autolysis is unknown. Remarkably, eukaryotic proteasomal β5 subunits bear a His residue in position (-2) of the propeptide (Supplementary Fig. 3a). As histidine commonly functions as a proton shuttle in the catalytic triads of serine proteases, we investigated the role of His(-2) in processing of the β5 propeptide by exchanging it for Asn, Lys, Phe and Ala. All yeast mutants were viable at 30 °C, but suffered from growth defects at 37 °C with the H(-2)N and H(-2)F mutants being most affected (Supplementary Fig. 3b and Table 1). In agreement, the chymotrypsin-like (ChT-L) activity of H(-2)N and H(-2)F mutant yCPs was impaired in situ and in vitro (Supplementary Fig. 3c). Structural analyses revealed that the propeptides of all mutant yCPs shared residual 2FO–FC electron densities. Gly(-1) and Phe/Lys(-2) were visualized at low occupancy, while Ala/Asn(-2) could not be assigned. This observation indicates a mixture of processed and unprocessed β5 subunits and partially impaired autolysis, thereby excluding any essential role of residue (-2) as the general base. Next, we examined the effect of residue (-2) on the orientation of the propeptide by creating mutants that combine the T1A (K81R) mutation(s) with H(-2)L, H(-2)T or H(-2)A substitutions. Leu(-2) is encoded in the yeast β1 subunit precursor (Supplementary Fig. 3a); Thr(-2) is generally part of β2-propeptides (Supplementary Fig. 3a); and Ala(-2) was expected to fit the β5-S1 pocket without inducing conformational changes of Met45, allowing it to accommodate ‘β1-like' propeptide positioning. As expected from β5-T1A mutants, the yeasts show severe growth phenotypes, with minor variations (Supplementary Fig. 4a and Table 1). We determined crystal structures of the β5-H(-2)L-T1A, β5-H(-2)T-T1A and the β5-H(-2)A-T1A-K81R mutants (Supplementary Table 1). For the β5-H(-2)A-T1A-K81R variant, only the residues Gly(-1) and Ala(-2) could be visualized, indicating that Ala(-2) leads to insufficient stabilization of the propeptide in the substrate-binding channel (Supplementary Fig. 4d). By contrast, the prosegments of the β5-H(-2)L-T1A and the β5-H(-2)T-T1A mutants were significantly better resolved in the 2FO–FC electron-density maps yet not at full occupancy (Supplementary Fig. 4b,c and Supplementary Table 1), suggesting that the natural propeptide bearing His(-2) is most favourable. Nevertheless, both Leu(-2) and Thr(-2) were found to occupy the S1 specificity pocket formed by Met45 (Fig. 2a,b and Supplementary Fig. 4f–h). This result proves that the naturally occurring His(-2) of the β5 propeptide does not stably fit into the S1 site. Since Gly(-1) adopts the same position in both wild-type (WT) and mutant β5 propeptides, and since in all cases its carbonyl carbon is perfectly placed for nucleophilic attack by Thr1Oγ (Fig. 2b), we propose that neither binding of residue (-2) to the S1 pocket nor formation of the antiparallel β-sheet is essential for autolysis of the propeptide. Next, we determined the crystal structure of a chimeric yCP having the yeast β1-propeptide replaced by its β5 counterpart. Although we observed fragments of 2FO–FC electron density in the β1 active site, the data were not interpretable. Bearing in mind that in contrast to Thr(-2) in β2, Leu(-2) in subunit β1 is not conserved among species (Supplementary Fig. 3a), we created a β2-T(-2)V proteasome mutant. As proven by the β2-T1A crystal structures, Thr(-2) hydrogen bonds to Gly(-1)O. Although this interaction was not observed for the β5-H(-2)T-T1A mutant (Fig. 2c and Supplementary Fig. 4c,i), exchange of Thr(-2) by Val in β2, a conservative mutation regarding size but drastic with respect to polarity, was found to inhibit maturation of this subunit (Fig. 2d and Supplementary Fig. 4e,j). Notably, the 2FO–FC electron-density map displays a different orientation for the β2 propeptide than has been observed for the β2-T1A proteasome. In particular, Val(-2) is displaced from the S1 site and Gly(-1) is severely shifted (movement of the carbonyl oxygen atom of 3.8 Å), thereby preventing nucleophilic attack of Thr1 (Fig. 2d and Supplementary Fig. 4j,k). These results further confirm that correct positioning of the active-site residues and Gly(-1) is decisive for the maturation of the proteasome. The active site of the proteasome Proton shuttling from the proteasomal active site Thr1OH to Thr1NH2 via a nucleophilic water molecule was suggested to initiate peptide-bond hydrolysis. However, in the immature particle Thr1NH2 is blocked by the propeptide and cannot activate Thr1Oγ. Instead, Lys33NH2, which is in hydrogen-bonding distance to Thr1Oγ (2.7 Å) in all catalytically active β subunits (Fig. 3a,b), was proposed to serve as the proton acceptor. Owing to its likely protonation at neutral pH, however, it was assumed not to act as the general base. A proposed catalytic tetrad model involving Thr1OH, Thr1NH2, Lys33NH2 and Asp17Oδ, as well as a nucleophilic water molecule as the proton shuttle appeared to accommodate all possible views of the proteasomal active site. Twenty years later, with a plethora of yCP X-ray structures in hand, we decided to re-analyse the active site of the proteasome and to resolve the uncertainty regarding the nature of the general base. Mutation of β5-Lys33 to Ala causes a strongly deleterious phenotype, and previous structural and biochemical analyses confirmed that this is caused by failure of propeptide cleavage, and consequently, lack of ChT-L activity (Fig. 4a, Supplementary Fig. 3b and Table 1; for details see Supplementary Note 1). The phenotype of the β5-K33A mutant was however less pronounced than for the β5-T1A-K81R yeast (Fig. 4a). This discrepancy in growth was traced to an additional point mutation L(-49)S in the β5-propeptide of the β5-K33A mutant (see also Supplementary Note 1). Structural comparison of the β5-L(-49)S-K33A and β5-T1A-K81R active sites revealed that mutation of Lys33 to Ala creates a cavity that is filled with Thr1 and the remnant propeptide. This structural alteration destroys active-site integrity and abolishes catalytic activity of the β5 active site (Supplementary Fig. 5a). Additional proof for the key function of Lys33 was obtained from the β5-K33A mutant, with the propeptide expressed separately from the main subunit (pp trans). The Thr1 N terminus of this mutant is not blocked by the propeptide, yet its catalytic activity is reduced by ∼83% (Supplementary Fig. 6b). Consistent with this, the crystal structure of the β5-K33A pp trans mutant in complex with carfilzomib only showed partial occupancy of the ligand at the β5 active sites (Supplementary Fig. 5b and Supplementary Table 1). Since no acetylation of the Thr1 N terminus was observed for the β5-K33A pp trans apo crystal structure, the reduced reactivity towards substrates and inhibitors indicates that Lys33NH2, rather than Thr1NH2, deprotonates and activates Thr1OH. Furthermore, the crystal structure of the β5-K33A pp trans mutant without inhibitor revealed that Thr1Oγ strongly coordinates a well-defined water molecule (∼2 Å; Fig. 3c and Supplementary Fig. 5c,d). This water hydrogen bonds also to Arg19O (∼3.0 Å) and Asp17Oδ (∼3.0 Å), and thereby presumably enables residual activity of the mutant. Remarkably, the solvent molecule occupies the position normally taken by Lys33NH2 in the WT proteasome structure (Fig. 3c), further corroborating the essential role of Lys33 as the general base for autolysis and proteolysis. Conservative substitution of Lys33 by Arg delays autolysis of the β5 precursor and impairs yeast growth (for details see Supplementary Note 1). While Thr1 occupies the same position as in WT yCPs, Arg33 is unable to hydrogen bond to Asp17, thereby inactivating the β5 active site (Supplementary Fig. 5e). The conservative mutation of Asp17 to Asn in subunit β5 of the yCP also provokes a severe growth defect (Supplementary Note 1, Supplementary Fig. 6a and Table 1). Notably, only with the additional point mutation L(-49)S present in the β5 propeptide could we purify a small amount of the β5-D17N mutant yCP. As determined by crystallographic analysis, this mutant β5 subunit was partially processed (Table 1) but displayed impaired reactivity towards the proteasome inhibitor carfilzomib compared with the subunits β1 and β2, and with WT β5 (Supplementary Fig. 7a). In contrast to the cis-construct, expression of the β5 propeptide in trans allowed straightforward isolation and crystallization of the D17N mutant proteasome. The ChT-L activity of the β5-D17N pp in trans CP towards the canonical β5 model substrates N-succinyl-Leu-Leu-Val-Tyr-7-amino-4-methylcoumarin (Suc-LLVY-AMC) and carboxybenzyl-Gly-Gly-Leu-para-nitroanilide (Z-GGL-pNA) was severely reduced (Supplementary Fig. 6b), confirming that Asp17 is of fundamental importance for the catalytic activity of the mature proteasome. Even though the β5-D17N pp trans yCP crystal structure appeared identical to the WT yCP (Supplementary Fig. 7b), the co-crystal structure with the α′, β′ epoxyketone inhibitor carfilzomib visualized only partial occupancy of the ligand in the β5 active site (Supplementary Fig. 7a). This observation is consistent with a strongly reduced reactivity of β5-Thr1 and the crystal structure of the β5-D17N pp cis mutant in complex with carfilzomib. Autolysis and residual catalytic activity of the β5-D17N mutants may originate from the carbonyl group of Asn17, which albeit to a lower degree still can polarize Lys33 for the activation of Thr1. In agreement, an E17A mutant in the proteasomal β-subunit of the archaeon Thermoplasma acidophilum prevents autolysis and catalysis. Strikingly, although the X-ray data on the β5-D17N mutant with the propeptide expressed in cis and in trans looked similar, there was a pronounced difference in their growth phenotypes observed (Supplementary Fig. 6a and Supplementary Fig. 7b). On the basis of these results, we propose that CPs from all domains of life use a catalytic triad consisting of Thr1, Lys33 and Asp/Glu17 for both autocatalytic precursor processing and proteolysis (Fig. 3d). This model is also consistent with the fact that no defined water molecule is observed in the mature WT proteasomal active site that could shuttle the proton from Thr1Oγ to Thr1NH2. To explore this active-site model further, we exchanged the conserved Asp166 residue for Asn in the yeast β5 subunit. Asp166Oδ is hydrogen-bonded to Thr1NH2 via Ser129OH and Ser169OH, and therefore was proposed to be involved in catalysis. The β5-D166N pp cis yeast mutant is significantly impaired in growth and its ChT-L activity is drastically reduced (Supplementary Fig. 6a,b and Table 1). X-ray data on the β5-D166N mutant indicate that the β5 propeptide is hydrolysed, but due to reorientation of Ser129OH, the interaction with Asn166Oδ is disrupted (Supplementary Fig. 8a). Instead, a water molecule is bound to Ser129OH and Thr1NH2 (Supplementary Fig. 8b), which may enable precursor processing. The hydrogen bonds involving Ser169OH are intact and may account for residual substrate turnover. Soaking the β5-D166N crystals with carfilzomib and MG132 resulted in covalent modification of Thr1 at high occupancy (Supplementary Fig. 8c). In the carfilzomib complex structure, Thr1Oγ and Thr1N incorporate into a morpholine ring structure and Ser129 adopts its WT-like orientation. In the MG132-bound state, Thr1N is unmodified, and we again observe that Ser129 is hydrogen-bonded to a water molecule instead of Asn166. Whereas Asn can to some degree replace Asp166 due to its carbonyl group in the side chain, Ala at this position was found to prevent both autolysis and catalysis. These results suggest that Asp166 and Ser129 function as a proton shuttle and affect the protonation state of Thr1N during autolysis and catalysis. Substitution of the active-site Thr1 by Cys Mutation of Thr1 to Cys inactivates the 20S proteasome from the archaeon T. acidophilum. In yeast, this mutation causes a strong growth defect (Fig. 4a and Table 1), although the propeptide is hydrolysed, as shown here by its X-ray structure. In one of the two β5 subunits, however, we found the cleaved propeptide still bound in the substrate-binding channel (Fig. 4c). His(-2) occupies the S2 pocket like observed for the β5-T1A-K81R mutant, but in contrast to the latter, the propeptide in the T1C mutant adopts an antiparallel β-sheet conformation as known from inhibitors like MG132 (Fig. 4c–e and Supplementary Fig. 9b). On the basis of the phenotype of the T1C mutant and the propeptide remnant identified in its active site, we suppose that autolysis is retarded and may not have been completed before crystallization. Owing to the unequal positions of the two β5 subunits within the CP in the crystal lattice, maturation and propeptide displacement may occur at different timescales in the two subunits. Despite propeptide hydrolysis, the β5-T1C active site is catalytically inactive (Fig. 4b and Supplementary Fig. 9a). In agreement, soaking crystals with the CP inhibitors bortezomib or carfilzomib modifies only the β1 and β2 active sites, while leaving the β5-T1C proteolytic centres unmodified even though they are only partially occupied by the cleaved propeptide remnant. Moreover, the structural data reveal that the thiol group of Cys1 is rotated by 74° with respect to the hydroxyl side chain of Thr1 (Fig. 4f and Supplementary Fig. 9b). This presumably results from the larger radius of the sulfur atom compared with oxygen. Consequently, the hydrogen bond bridging the active-site nucleophile and Lys33 in WT CPs is broken with Cys1. Notably, the 2FO–FC electron-density map of the T1C mutant also indicates that Lys33NH2 is disordered. Together, these observations suggest that efficient peptide-bond hydrolysis requires that Lys33NH2 hydrogen bonds to the active site nucleophile. The benefit of Thr over Ser as the active-site nucleophile All proteasomes strictly employ threonine as the active-site residue instead of serine. To investigate the reason for this singularity, we analysed a β5-T1S mutant, which is viable but suffers from growth defects (Fig. 4a and Table 1). Activity assays with the β5-specific substrate Suc-LLVY-AMC demonstrated that the ChT-L activity of the T1S mutant is reduced by 40–45% compared with WT proteasomes depending on the incubation temperature (Fig. 4b and Supplementary Fig. 9c). By contrast, turnover of the substrate Z-GGL-pNA, used to monitor ChT-L activity in situ but in a less quantitative fashion, is not detectably impaired (Supplementary Fig. 9a). Crystal structure analysis of the β5-T1S mutant confirmed precursor processing (Fig. 4g), and ligand-complex structures with bortezomib and carfilzomib unambiguously corroborated the reactivity of Ser1 (Fig. 5). However, the apo crystal structure revealed that Ser1Oγ is turned away from the substrate-binding channel (Fig. 4g). Compared with Thr1Oγ in WT CP structures, Ser1Oγ is rotated by 60°. This renders it unavailable for direct nucleophilic attack onto incoming substrates and first requires its reorientation, which is expected to delay substrate turnover. Because both conformations of Ser1Oγ are hydrogen-bonded to Lys33NH2 (Fig. 4h), the relay system is capable of hydrolysing peptide substrates, albeit at lower rates compared with Thr1. The active-site residue Thr1 is fixed in its position, as its methyl group is engaged in hydrophobic interactions with Thr3 and Ala46 (Fig. 4h). Consequently, the hydroxyl group of Thr1 requires no reorientation before substrate cleavage and is thus more catalytically efficient than Ser1. In agreement, at an elevated growing temperature of 37 °C the T1S mutant is unable to grow (Fig. 4a). In vitro, the mutant proteasome is less susceptible to proteasome inhibition by bortezomib (3.7-fold) and carfilzomib (1.8-fold; Fig. 5). Nevertheless, inhibitor complex structures indicate identical binding modes compared with the WT yCP structures, with the same inhibitors. Notably, the affinity of the tetrapeptide carfilzomib is less impaired, as it is better stabilized in the substrate-binding channel than the dipeptide bortezomib, which lacks a defined P3 site and has only a few interactions with the surrounding protein. Hence, the mean residence time of carfilzomib at the active site is prolonged and the probability to covalently react with Ser1 is increased. Considered together, these results provide a plausible explanation for the invariance of threonine as the active-site nucleophile in proteasomes in all three domains of life, as well as in proteasome-like proteases such as HslV (ref.). Discussion The 20S proteasome CP is the major non-lysosomal protease in eukaryotic cells, and its assembly is highly organized. The β-subunit propeptides, particularly that of β5, are key factors that help drive proper assembly of the CP complex. In addition, they prevent irreversible inactivation of the Thr1 N terminus by N-acetylation. By contrast, the prosegments of β subunits are dispensable for archaeal proteasome assembly, at least when heterologously expressed in Escherichia coli. In eukaryotes, deletion of or failure to cleave the β1 and β2 propeptides is well tolerated. However, removal of the β5 prosegment or any interference with its cleavage causes severe phenotypic defects. These observations highlight the unique function and importance of the β5 propeptide as well as the β5 active site for maturation and function of the eukaryotic CP. Here we have described the atomic structures of various β5-T1A mutants, which allowed for the first time visualization of the residual β5 propeptide. Depending on the (-2) residue we observed various propeptide conformations, but Gly(-1) is in all structures perfectly located for the nucleophilic attack by Thr1Oγ, although it does not adopt the tight turn observed for the prosegment of subunit β1. From these data we conclude that only the positioning of Gly(-1) and Thr1 as well as the integrity of the proteasomal active site are required for autolysis. In this regard, inappropriate N-acetylation of the Thr1 N terminus cannot be removed by Thr1Oγ due to the rotational freedom and flexibility of the acetyl group. The propeptide needs some anchoring in the substrate-binding channel to properly position Gly(-1), but this seems to be independent of the orientation of residue (-2). Autolytic activation of the CP constitutes one of the final steps of proteasome biogenesis, but the trigger for propeptide cleavage had remained enigmatic. On the basis of the numerous CP:ligand complexes solved during the past 18 years and in the current study, we provide a revised interpretation of proteasome active-site architecture. We propose a catalytic triad for the active site of the CP consisting of residues Thr1, Lys33 and Asp/Glu17, which are conserved among all proteolytically active eukaryotic, bacterial and archaeal proteasome subunits. Lys33NH2 is expected to act as the proton acceptor during autocatalytic removal of the propeptides, as well as during substrate proteolysis, while Asp17Oδ orients Lys33NH2 and makes it more prone to protonation by raising its pKa (hydrogen bond distance: Lys33NH3+–Asp17Oδ: 2.9 Å). Analogously to the proteasome, a Thr–Lys–Asp triad is also found in L-asparaginase. Thus, specific protein surroundings can significantly alter the chemical properties of amino acids such as Lys to function as an acid–base catalyst. In this new view of the proteasomal active site, the positively charged Thr1NH3+-terminus hydrogen bonds to the amide nitrogen of incoming peptide substrates and stabilizes as well as activates them for the endoproteolytic cleavage by Thr1Oγ (Fig. 3d). Consistent with this model, the positively charged Thr1 N terminus is engaged in hydrogen bonds with inhibitory compounds like fellutamide B (ref.), α-ketoamides, homobelactosin C (ref.) and salinosporamide A (ref.). Furthermore, opening of the β-lactone compound omuralide by Thr1 creates a C3-hydroxyl group, whose proton originates from Thr1NH3+. The resulting uncharged Thr1NH2 is hydrogen-bridged to the C3-OH group. In agreement, acetylation of the Thr1 N terminus irreversibly blocks hydrolytic activity, and binding of substrates is prevented for steric reasons. By acting as a proton donor during catalysis, the Thr1 N terminus may also favour cleavage of substrate peptide bonds (Fig. 3d). In all proteases, collapse of the tetrahedral transition state results in selective breakage of the substrate amide bond, while the covalent interaction between the substrate and the enzyme persists. Cleavage of the scissile peptide bond requires protonation of the emerging free amine, and in the proteasome, the Thr1 amine group is likely to assume this function. Analogously, Thr1NH3+ might promote the bivalent reaction mode of epoxyketone inhibitors by protonating the epoxide moiety to create a positively charged trivalent oxygen atom that is subsequently nucleophilically attacked by Thr1NH2. During autolysis the Thr1 N terminus is engaged in a hydroxyoxazolidine ring intermediate (Fig. 3d), which is unstable and short-lived. Breakdown of this tetrahedral transition state releases the Thr1 N terminus that is protonated by aspartic acid 166 via Ser129OH to yield Thr1NH3+. The residues Ser129 and Asp166 are expected to increase the pKa value of Thr1N, thereby favouring its charged state. Consistent with playing an essential role in proton shuttling, the mutation D166A prevents autolysis of the archaeal CP and the exchange D166N impairs catalytic activity of the yeast CP about 60%. The mutation D166N lowers the pKa of Thr1N, which is thus more likely to exist in the uncharged deprotonated state (Thr1NH2). This renders the N terminus less suitable to stabilize substrates and to protonate the first cleavage product during catalysis, although it favours its ability to act as a nucleophile. This interpretation agrees with the strongly reduced catalytic activity of the β5-D166N mutant on the one hand, and the ability to react readily with carfilzomib on the other. Hence, the proteasome can be viewed as having a second triad that is essential for efficient proteolysis. While Lys33NH2 and Asp17Oδ are required to deprotonate the Thr1 hydroxyl side chain, Ser129OH and Asp166OH serve to protonate the N-terminal amine group of Thr1. In accord with the proposed Thr1–Lys33–Asp17 catalytic triad, crystallographic data on the proteolytically inactive β5-T1C mutant demonstrate that the interaction of Lys33NH2 and Cys1 is broken. Consequently, efficient substrate turnover or covalent modification by ligands is prevented. However, owing to Cys being a strong nucleophile, the propeptide can still be cleaved off over time. While only one single turnover is necessary for autolysis, continuous enzymatic activity is required for significant and detectable substrate hydrolysis. Notably, in the Ntn hydrolase penicillin acylase, substitution of the catalytic N-terminal Ser residue by Cys also inactivates the enzyme but still enables precursor processing. To investigate why the CP specifically employs threonine as its active-site residue, we used a β5-T1S mutant of the yCP and characterized it biochemically and structurally. Activity assays with the β5-T1S mutant revealed reduced turnover of Suc-LLVY-AMC. We also observed slightly lower affinity of the β5-T1S mutant yCP for the Food and Drug Administration-approved proteasome inhibitors bortezomib and carfilzomib. Structural analyses support these findings with the T1S mutant and provide an explanation for the strict use of Thr residues in proteasomes. Thr1 is well anchored in the active site by hydrophobic interactions of its Cγ methyl group with Ala46 (Cβ), Lys33 (carbon side chain) and Thr3 (Cγ). Notably, proteolytically active proteasome subunits from archaea, yeast and mammals, including constitutive, immuno- and thymoproteasome subunits, either encode Thr or Ile at position 3, indicating the importance of the Cγ for fixing the position of the nucleophilic Thr1. In contrast to Thr1, the hydroxyl group of Ser1 occupies the position of the Thr1 methyl side chain in the WT enzyme, which requires its reorientation relative to the substrate to allow cleavage (Fig. 4g,h). Notably, in the threonine aspartase Taspase1, mutation of the active-site Thr234 to Ser also places the side chain in the position of the methyl group of Thr234 in the WT, thereby reducing catalytic activity. Similarly, although the serine mutant is active, threonine is more efficient in the context of the proteasome active site. The greater suitability of threonine for the proteasome active site, which has been noted in biochemical as well as in kinetic studies, constitutes a likely reason for the conservation of the Thr1 residue in all proteasomes from bacteria to eukaryotes. Methods Yeast mutagenesis Site-directed mutagenesis was performed by standard techniques using oligonucleotides listed in Supplementary Table 2. The pre2/doa3 (β5) mutant alleles in the centromeric, TRP1- or LEU2-marked shuttle vectors YCplac22 and pRS315, respectively, were verified by sequencing and subsequently introduced into the yeast strains MHY784 (ref.) or YWH20 (ref.), which express WT PRE2 from a URA3-marked plasmid. Counter-selection against the URA3 marker with 5-fluoroorotic acid yielded strains expressing only the mutant forms of β5. The strain producing a processed β5-T1A variant and the β5 propeptide in trans is a derivative of YWH212 (ref.). It carries an additional deletion of the NAT1 gene to avoid N-acetylation of Ala1; this strain exhibits extremely slow growth rates and served for crystallographic analysis only. All strains used in this study are listed in Supplementary Table 3. Purification of yeast proteasomes Yeast strains were grown in 18-l cultures at 30 °C in YPD into early stationary phase, and the yCPs were purified according to published procedures. In brief, 120 g yeast cells were solubilized in 150 ml of 50 mM KH2PO4/K2HPO4 buffer (pH 7.5) and disrupted with a French press. Cell debris were removed by centrifugation for 30 min at 21,000 r.p.m. (4 °C). The resulting supernatant was filtered and ammonium sulfate (saturated solution) was added to a final concentration of 30% (v/v). This solution was loaded on a Phenyl Sepharose 6 Fast Flow column (GE Healthcare) pre-equilibrated with 1 M ammonium sulfate in 20 mM KH2PO4/K2HPO4 (pH 7.5). CPs were eluted by applying a linear gradient from 1 to 0 M ammonium sulfate. Proteasome-containing fractions were pooled and loaded onto a hydroxyapatite column (Bio-Rad) equilibrated with 20 mM KH2PO4/K2HPO4 (pH 7.5). Elution of the CPs was achieved by increasing the phosphate buffer concentration from 20 to 500 mM. Anion-exchange chromatogaphy (Resource Q column (GE Healthcare), elution gradient from 0 to 500 mM sodium chloride in 20 mM Tris-HCl (pH 7.5)) and subsequent size-exclusion chromatography (Superose 6 10/300 GL (GE Healthcare), 20 mM Tris-HCl (pH 7.5) and 150 mM NaCl) resulted in pure CPs for crystallization and activity assays. Fluorescence-based activity assay ChT-L (β5) activity of CPs was monitored by fluorescence spectroscopy using the model substrate Suc-LLVY-AMC. Purified yCPs (66 nM in 100 mM Tris-HCl, pH 7.5) were incubated with 300 μM substrate for 1 h at room temperature or 37 °C. The reactions were stopped by diluting samples 1:10 in 20 mM Tris-HCl, pH 7.5. AMC fluorophores released by proteasomal activity were measured in triplicate with a Varian Cary Eclipse Fluorescence Spectrophotometer (Agilent Technologies) at λexc=360 nm and λem=460 nm. Inhibition assays Purified yCPs were mixed with dimethylsulfoxide as a control or serial dilutions of inhibitor and incubated for 45 min at room temperature. A final concentration of yCP of 66 nM was used. After addition of the peptide substrate Suc-LLVY-AMC to a final concentration of 200 μM and incubation for 1 h at room temperature, the reaction was stopped by diluting the samples 1:10 in 20 mM Tris-HCl, pH 7.5. AMC fluorophores released by residual proteasomal activity were measured in triplicate at λexc=360 nm and λem=460 nm. Relative fluorescence units were normalized to the dimethylsulfoxide-treated control. The calculated residual activities were plotted against the logarithm of the applied inhibitor concentration and fitted with GraphPad Prism 5. The IC50 value, the ligand concentration that leads to 50% inhibition of the enzymatic activity, was deduced from the fitted data. Crystallization and structure determination Mutant yCPs were crystallized as previously described for WT 20S proteasomes. Crystals were grown at 20 °C using the hanging drop vapour diffusion method. Drops contained a 1:1 mixture of protein (40 mg ml−1) and reservoir solution (25 mM magnesium acetate, 100 mM 2-(N-morpholino)ethanesulfonic acid (MES; pH 6.8) and 9–13% (v/v) 2-methyl-2,4-pentanediol (MPD)). Crystals were cryoprotected by addition of 5 μl cryobuffer (20 mM magnesium acetate, 100 mM MES, pH 6.8, and 30% (v/v) MPD). Inhibitor complex structures were obtained by incubating crystals in 5 μl cryobuffer supplemented with bortezomib or carfilzomib at a final concentration of 1.5 mM for at least 8 h. Diffraction data were collected at the beamline X06SA at the Paul Scherrer Institute, SLS, Villigen, Switzerland (λ=1.0 Å). Evaluation of reflection intensities and data reduction were performed with the programme package XDS. Molecular replacement was carried out with the coordinates of the yeast 20S proteasome (PDB entry code: 5CZ4) by rigid body refinements (REFMAC5; ref.). MAIN and COOT were used to build models. TLS (Translation/Libration/Screw) refinements finally yielded excellent Rwork and Rfree, as well as root mean squared deviation bond and angle values. The coordinates, proven to have good stereochemistry from the Ramachandran plots, were deposited in the RCSB Protein Data Bank (Supplementary Table 1). The coordinates for the yeast 20S proteasome deposited under the entry code 1RYP do not represent the WT yCP but the double-mutant β5-K33R β1-T1A. At the time of deposition (in 1997), these data were the best available on the yCP. As yCP structure determination has become routine today, and structure refinement procedures have significantly improved, we here provide coordinates for the WT yCP at 2.3 Å resolution (PDB entry code: 5CZ4). Furthermore, the structures of most mutant yCPs described in this work were determined in their apo and ligand-bound states. For mutants with proteolytically inactive β5 subunits, the best crystallographic data obtained are given. For ligands or propeptide segments that were only partially defined in the 2FO–FC electron-density map the occupancy was reduced (for details see Supplementary Table 1). Additional information Accession codes: Coordinates and structure factors have been deposited in the Protein Data Bank, www.pdb.org (for PDB entry codes see Supplementary Table 1). How to cite this article: Huber, E. M. et al. A unified mechanism for proteolysis and autocatalytic activation in the 20S proteasome. Nat. Commun. 7:10900 doi: 10.1038/ncomms10900 (2016). Supplementary Material Autocatalytic subunit processing couples active site formation in the 20S proteasome to completion of assembly Structure of 20S proteasome from yeast at 2.4 Å resolution 20S proteasome biogenesis The catalytic sites of 20S proteasomes and their role in subunit maturation: a mutational and crystallographic study Conformational constraints for protein self-cleavage in the proteasome Immuno- and constitutive proteasome crystal structures reveal differences in substrate and inhibitor specificity Systematic analyses of substrate preferences of 20S proteasomes using peptidic epoxyketone inhibitors Catalytic mechanism and assembly of the proteasome Crystal structure of the 20S proteasome from the archaeon T. acidophilum at 3.4 Å resolution A protein catalytic framework with an N-terminal nucleophile is capable of self-activation Proteasome from Thermoplasma acidophilum: a threonine protease Contribution of proteasomal beta-subunits to the cleavage of peptide substrates analyzed with yeast mutants The active sites of the eukaryotic 20S proteasome and their involvement in subunit precursor processing Identification of the yeast 20S proteasome catalytic centers and subunit interactions required for active-site formation Proteasome beta-type subunits: unequal roles of propeptides in core particle maturation and a hierarchy of active site function Eukaryotic 20S proteasome catalytic subunit propeptides prevent active site inactivation by N-terminal acetylation and promote particle assembly The catalytic triad of serine peptidases Distinct elements in the proteasomal beta5 subunit propeptide required for autocatalytic processing and proteasome assembly Analysis of mammalian 20S proteasome biogenesis: the maturation of beta-subunits is an ordered two-step mechanism involving autocatalysis Proteasome: a complex protease with a new fold and a distinct mechanism Autocatalytic processing of the 20S proteasome Crystal structure of the boronic acid-based proteasome inhibitor bortezomib in complex with the yeast 20S proteasome Bortezomib-resistant mutant proteasomes: structural and biochemical evaluation with carfilzomib and ONX 0914 Crystal structure of heat shock locus V (HslV) from Escherichia coli Critical elements in proteasome assembly Investigations on the maturation and regulation of archaebacterial proteasomes Atomic resolution structure of Erwinia chrysanthemi L-asparaginase Understanding nature's catalytic toolkit Proteasome inhibition by fellutamide B induces nerve growth factor synthesis Systematic comparison of peptidic proteasome inhibitors highlights the alpha-ketoamide electrophile as an auspicious reversible lead motif Inhibitor-binding mode of homobelactosin C to proteasomes: new insights into class I MHC ligand generation Crystal structures of Salinosporamide A (NPI-0052) and B (NPI-0047) in complex with the 20S proteasome reveal important consequences of beta-lactone ring opening and a mechanism for irreversible binding Effects of site-directed mutations on processing and activities of penicillin G acylase from Escherichia coli ATCC 11105 Crystal structure of human Taspase1, a crucial protease regulating the function of MLL Why does threonine, and not serine, function as the active site nucleophile in proteasomes? Analysing properties of proteasome inhibitors using kinetic and X-ray crystallographic studies Purification, crystallization, and X-ray analysis of the yeast 20S proteasome XDS REFMAC5 dictionary: organization of prior chemical knowledge and guidelines for its use MAIN software for density averaging, model building, structure refinement and validation Features and development of Coot Author contributions E.M.H., W.H., X.L., C.S.A. and M.H. created yeast mutants; E.M.H. and W.H. performed activity and growth assays; E.M.H. and M.G. collected and analysed X-ray data; E.M.H., M.H. and M.G. wrote the manuscript. Conformation of proteasomal propeptides. (a) Structural superposition of the β1-T1A propeptide and the matured WT β1 active-site Thr1. Only the residues (-5) to (-1) of the β1-T1A propeptide are displayed. The major determinant of the S1 specificity pocket, residue 45, is depicted. Note the tight conformation of Gly(-1) and Ala1 before propeptide removal (G(-1) turn; cyan double arrow) compared with the relaxed, processed WT active-site Thr1 (red double arrow). The black arrow indicates the attack of Thr1Oγ onto the carbonyl carbon atom of Gly(-1). (b) Structural superposition of the β1-T1A propeptide and the β2-T1A propeptide highlights subtle differences in their conformations, but illustrates that Ala1 and Gly(-1) match well. Thr(-2)OH is hydrogen-bonded to Gly(-1)O (∼2.8 Å; black dashed line). The major determinant of the S1 specificity pocket, residue 45, is depicted. (c) Structural superposition of the β1-T1A, the β2-T1A and the β5-T1A-K81R propeptide remnants depict their differences in conformation. While residue (-2) of the β1 and β2 prosegments fit the S1 pocket, His(-2) of the β5 propeptide occupies the S2 pocket. Nonetheless, in all mutants the carbonyl carbon atom of Gly(-1) is ideally placed for the nucleophilic attack by Thr1Oγ. The hydrogen bond between Thr(-2)OH and Gly(-1)O (∼2.8 Å) is indicated by a black dashed line. Mutations of residue (-2) and their influence on propeptide conformation and autolysis. (a) Structural superposition of the β1-T1A propeptide and the β5-H(-2)L-T1A mutant propeptide. The (-2) residues of both prosegments point into the S1 pocket. (b) Structural superposition of the β5 propeptides in the β5-H(-2)L-T1A, β5-H(-2)T-T1A, β5-(H-2)A-T1A-K81R and β5-T1A-K81R mutant proteasomes. While the residues (-2) to (-4) vary in their conformation, Gly(-1) and Ala1 are located in all structures at the same positions. (c) Structural superposition of the β2-T1A propeptide and the β5-H(-2)T-T1A mutant propeptide. The (-2) residues of both prosegments point into the S1 pocket, but only Thr(-2)OH of β2 forms a hydrogen bridge to Gly(-1)O (black dashed line). (d) Structural superposition of the matured β2 active site, the WT β2-T1A propeptide and the β2-T(-2)V mutant propeptide. Notably, Val(-2) of the latter does not occupy the S1 pocket, thereby changing the orientation of Gly(-1) and preventing nucleophilic attack of Thr1Oγ on the carbonyl carbon atom of Gly(-1). For all panels stereo views are provided in Supplementary Fig. 4g–j. Architecture and proposed reaction mechanism of the proteasomal active site. (a) Hydrogen-bonding network at the mature WT β5 proteasomal active site (dotted lines). Thr1OH is hydrogen-bonded to Lys33NH2 (2.7 Å), which in turn interacts with Asp17Oδ. The Thr1 N terminus is engaged in hydrogen bonds with Ser129Oγ, the carbonyl oxygen of residue 168, Ser169Oγ and Asp166Oδ. (b) The orientations of the active-site residues involved in hydrogen bonding are strictly conserved in each proteolytic centre, as shown by superposition of the β subunits. (c) Structural superposition of the WT β5 and the β5-K33A pp trans mutant active site. In the latter, a water molecule (red sphere) is found at the position where in the WT structure the side chain amine group of Lys33 is located. Similarly to Lys33, the water molecule hydrogen bonds to Arg19O, Asp17Oδ and Thr1OH. Note, the strong interaction with the water molecule causes a minor shift of Thr1, while all other active-site residues remain in place. (d) Proposed chemical reaction mechanism for autocatalytic precursor processing and proteolysis in the proteasome. The active-site Thr1 is depicted in blue, the propeptide segment and the peptide substrate are coloured in green, whereas the scissile peptide bond is highlighted in red. Autolysis (left set of structures) is initiated by deprotonation of Thr1OH via Lys33NH2 and the formation of a tetrahedral transition state. The strictly conserved oxyanion hole Gly47NH stabilizing the negatively charged intermediate is illustrated as a semicircle. Collapse of the transition state frees the Thr1 N terminus (by completing an N-to-O acyl shift of the propeptide), which is subsequently protonated by Asp166OH via Ser129OH. Next, Thr1NH2 polarizes a water molecule for the nucleophilic attack of the acyl-enzyme intermediate. On hydrolysis of the latter, the active-site Thr1 is ready for catalysis (right set of structures). Substrate processing starts with nucleophilic attack of the carbonyl carbon atom of the scissile peptide bond. The charged Thr1 N terminus may engage in the orientation of the amide moiety and donate a proton to the emerging N terminus of the C-terminal cleavage product. The resulting deprotonated Thr1NH2 finally activates a water molecule for hydrolysis of the acyl-enzyme. The proteasome favours threonine as the active-site nucleophile. (a) Growth tests by serial dilution of WT and pre2 (β5) mutant yeast cultures reveal growth defects of the active-site mutants under the indicated conditions after 2 days (2 d) of incubation. (b) Purified WT and mutant proteasomes were tested for their chymotrypsin-like activity (β5) using the substrate Suc-LLVY-AMC. Relative fluorescence units were measured in triplicate after 1 h of incubation at room temperature and are given as mean values. S.d.'s are indicated by error bars. (c) Illustration of the 2FO–FC electron-density map (blue mesh contoured at 1σ) for the β5-T1C propeptide fragment. The prosegment is cleaved but still bound in the substrate-binding channel. Notably, His(-2) does not occupy the S1 pocket formed by Met45, similar to what was observed for the β5-T1A-K81R mutant. (d) Structural superposition of the β5-T1A-K81R and the β5-T1C mutant subunits onto the WT β5 subunit. (e) Structural superposition of the β5-T1C propeptide onto the β1-T1A active site (blue) and the WT β5 active site in complex with the proteasome inhibitor MG132 (ref.). The inhibitor as well as the propeptides adopt similar conformations in the substrate-binding channel. (f) Structural superposition of the WT β5 and β5-T1C mutant active sites illustrates the different orientations of the hydroxyl group of Thr1 and the thiol side chain of Cys1. The SH group is rotated by 74° compared with the OH group. (g) Structural superposition of the WT β5 and β5-T1S mutant active sites reveals different orientations of the hydroxyl groups of Thr1 and Ser1, respectively. The 2FO–FC electron-density map for Ser1 (blue mesh contoured at 1σ) is illustrated. (h) The methyl group of Thr1 is anchored by hydrophobic interactions with Ala46Cβ and Thr3Cγ. Ser1 lacks this stabilization and is therefore rotated by 60°. Inhibition of WT and mutant β5-T1S proteasomes by bortezomib and carfilzomib. Inhibition assays (left panel). Purified yeast proteasomes were tested for the susceptibility of their ChT-L (β5) activity to inhibition by bortezomib and carfilzomib using the substrate Suc-LLVY-AMC. IC50 values were determined in triplicate; s.d.'s are indicated by error bars. Note that IC50 values depend on time and enzyme concentration. Proteasomes (final concentration: 66 nM) were incubated with inhibitor for 45 min before substrate addition (final concentration: 200 μM). Structures of the β5-T1S mutant in complex with both ligands (green) prove the reactivity of Ser1 (right panel). The 2FO–FC electron-density maps (blue mesh) for Ser1 (brown) and the covalently bound ligands (green; only the P1 site (Leu1) is shown) are contoured at 1σ. The WT proteasome:inhibitor complex structures (inhibitor in grey; Thr1 in black) are superimposed and demonstrate that mutation of Thr1 to Ser does not affect the binding mode of bortezomib or carfilzomib.  Growth phenotypes and status of autolysis and catalysis of mutants. Mutant\tViability\tTemperature sensitivity\tAutolysis state of the mutant subunit*\tActivity of the mutant subunit\t \tWT\t++++\t+\t+\t+++\t \tβ1-T1A (ref.)\t++++\t+\t−\t−\t \tβ2-T1A (ref.)\t+++\t++\t−\t−\t \tβ1-T1A β2-T1A (ref.)\t+++\t++\t−\t−\t \tβ5-T1A\t+/−\t++++\t−\t−\t \tβ5-H(-2)A-T1A\t+/−\tND\t−\t−\t \tβ5-H(-2)T-T1A\t+/−\tND\t−\t−\t \tβ5-H(-2)L-T1A\t++\tND\t−\t−\t \tβ5-T1A, pp trans, nat1Δ\t+/−\t++++\tpp trans\t−\t \tβ5-T1A-K81R\t+\t++++\t−\t−\t \tβ5-H(-2)A-T1A-K81R\t+/−\tND\t−\t−\t \tβ5-H(-2)T-T1A-K81R\t+/−\tND\t−\t−\t \tβ5-H(-2)L-T1A-K81R\t++\tND\t−\t−\t \tβ5-H(-2)A\t++++\t++\t+\t+++\t \tβ5-H(-2)K\t++++\t++\t+\t+++\t \tβ5-H(-2)F\t++++\t+++\t+\t++\t \tβ5-H(-2)N\t++++\t+++\t+\t++\t \tβ5pp-β1 (ref. 18)\t++++\t+\t+/−\t+/−\t \tβ2-T(-2)V\t++++\t+\t−\t−\t \tβ5-L(-49S)-K33A (ref.)\t+\t++++\t−\t−\t \tβ5-K33A, pp trans\t+\t++++\tpp trans\t+/−\t \tβ5-F(-45)S-K33R (ref.)\t++\t++++\t+\t−\t \tβ5-D17N\t+/−\t++++\tND†\tND†\t \tβ5-L(-49)S-D17N\t+\t++++\t+/−\t+/−\t \tβ5-D17N, pp trans\t+\t++++\tpp trans\t+/−\t \tβ5-D166N\t++\t++++\t+\t+/−\t \tβ5-D166N, pp trans\t+++\t++++\tpp trans\t+/−\t \tβ5-T1S\t+++\t++++\t+\t++\t \tβ5-T1C\t++\t++++\t+\t−\t \t ND, not determined. *The autolysis state was assessed by purification and crystallization of the mutant proteasomes. †Purification of this mutant proteasome was not 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diff --git a/annotated_BioC_JSON/PMC4795551_ann.json b/annotated_BioC_JSON/PMC4795551_ann.json
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index 0000000000000000000000000000000000000000..dac82d7bc76b1b14b18805b20b4f298e54f7cd80
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+++ b/annotated_BioC_JSON/PMC4795551_ann.json
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+[{"sourceid":"4795551","sourcedb":"","project":"","target":"","text":"Biochemistry and Crystal Structure of Ectoine Synthase: A Metal-Containing Member of the Cupin Superfamily Ectoine is a compatible solute and chemical chaperone widely used by members of the Bacteria and a few Archaea to fend-off the detrimental effects of high external osmolarity on cellular physiology and growth. Ectoine synthase (EctC) catalyzes the last step in ectoine production and mediates the ring closure of the substrate N-gamma-acetyl-L-2,4-diaminobutyric acid through a water elimination reaction. However, the crystal structure of ectoine synthase is not known and a clear understanding of how its fold contributes to enzyme activity is thus lacking. Using the ectoine synthase from the cold-adapted marine bacterium Sphingopyxis alaskensis (Sa), we report here both a detailed biochemical characterization of the EctC enzyme and the high-resolution crystal structure of its apo-form. Structural analysis classified the (Sa)EctC protein as a member of the cupin superfamily. EctC forms a dimer with a head-to-tail arrangement, both in solution and in the crystal structure. The interface of the dimer assembly is shaped through backbone-contacts and weak hydrophobic interactions mediated by two beta-sheets within each monomer. We show for the first time that ectoine synthase harbors a catalytically important metal co-factor; metal depletion and reconstitution experiments suggest that EctC is probably an iron-dependent enzyme. We found that EctC not only effectively converts its natural substrate N-gamma-acetyl-L-2,4-diaminobutyric acid into ectoine through a cyclocondensation reaction, but that it can also use the isomer N-alpha-acetyl-L-2,4-diaminobutyric acid as its substrate, albeit with substantially reduced catalytic efficiency. Structure-guided site-directed mutagenesis experiments targeting amino acid residues that are evolutionarily highly conserved among the extended EctC protein family, including those forming the presumptive iron-binding site, were conducted to functionally analyze the properties of the resulting EctC variants. An assessment of enzyme activity and iron content of these mutants give important clues for understanding the architecture of the active site positioned within the core of the EctC cupin barrel. Introduction Compatible solutes are exploited by members of all three domains of life as versatile cyto-protectants, in particular against cellular stress elicited by high osmolarity environments. They are especially useful for this latter purpose since their benign nature allows their amassing to exceedingly high cellular concentrations. As a result of compatible solute accumulation, dehydration of the cytoplasm of osmotically stressed cells is counteracted, and concomitantly, its solvent properties are optimized for the functioning of vital biochemical and physiological processes. Ectoine [(S)-2-methyl-1,4,5,6-tetrahydropyrimidine-4-carboxylic acid] and its derivative 5-hydroxyectoine [(4S,5S)-5-hydroxy-2-methyl-1,4,5,6-tetrahydropyrimidine-4-carboxylic acid] are such compatible solutes. Both marine and terrestrial microorganisms produce them widely in response to osmotic or temperature stress. Synthesis of ectoine occurs from the intermediate metabolite L-aspartate-ß-semialdehyde and comprises the sequential activities of three enzymes: L-2,4-diaminobutyrate transaminase (EctB; EC 2.6.1.76), 2,4-diaminobutyrate acetyltransferase (EctA; EC 2.3.1.178), and ectoine synthase (EctC; EC 4.2.1.108) (Fig 1). The ectoine derivative 5-hydroxyectoine, a highly effective stress protectant in its own right, is synthesized by a substantial subgroup of the ectoine producers. This stereospecific chemical modification of ectoine (Fig 1) is catalyzed by the ectoine hydroxylase (EctD) (EC 1.14.11), a member of the non-heme containing iron(II) and 2-oxoglutarate-dependent dioxygenase superfamily. The remarkable function preserving effects of ectoines for macromolecules and cells, frequently also addressed as chemical chaperones, led to a substantial interest in exploiting these compounds for biotechnological purposes and medical applications. Biosynthetic routes for ectoine and 5-hydroxyectoine. Scheme of the ectoine and 5-hydroxyectoine biosynthetic pathway. Here we focus on ectoine synthase (EctC), the key enzyme of the ectoine biosynthetic route (Fig 1). Biochemical characterizations of ectoine synthases from the extremophiles Halomonas elongata, Methylomicrobium alcaliphilum, and Acidiphilium cryptum, and from the nitrifying archaeon Nitrosopumilus maritimus have been carried out. Each of these enzymes catalyzes as their main activity the cyclization of N-γ-acetyl-L-2,4-diaminobutyric acid (N-γ-ADABA), the reaction product of the 2,4-diaminobutyrate acetyltransferase (EctA), to ectoine with the concomitant release of a water molecule (Fig 1). In side reactions, EctC can promote the formation of the synthetic compatible solute 5-amino-3,4-dihydro-2H-pyrrole-2-carboxylate (ADPC) through the cyclic condensation of two glutamine molecules and it also possesses a minor hydrolytic activity for ectoine and synthetic ectoine derivatives with either reduced or expanded ring sizes. Although progress has been made with respect to the biochemical characterization of ectoine synthase, a clear understanding of how its structure contributes to its enzyme activity and reaction mechanism is still lacking. With this in mind, we have biochemically characterized the ectoine synthase from the cold-adapted marine bacterium Sphingopyxis alaskensis (Sa). We demonstrate here for the first time that the ectoine synthase is a metal-dependent enzyme, with iron as the most likely physiologically relevant co-factor. The EctC protein forms a dimer in solution and our structural analysis identifies it as a member of the cupin superfamily. The two crystal structures that we report here for the (Sa)EctC protein (with resolutions of 1.2 Å and 2.0 Å, respectively), and data derived from extensive site-directed mutagenesis experiments targeting evolutionarily highly conserved residues within the extended EctC protein family, provide a first view into the architecture of the catalytic core of the ectoine synthase. Materials and Methods Chemicals Ectoine [(S)-2-methyl-1,4,5,6-tetrahydropyrimidine-4-carboxylic acid] was a kind gift from bitop AG (Witten, Germany). Anhydrotetracycline (AHT), desthiobiotine and the strepavidin affinity matrix for the purification of Strep-tag II labeled proteins was purchased from IBA GmbH (Göttingen, Germany). Hydroxylamine and phenanthroline for the photometric determination of the iron-content of the recombinant (Sa)EctC proteins were purchased from Sigma-Aldrich (München, Germany). Synthesis of N-γ-acetyl-L-2,4-diaminobutyric acid and N-α-acetyl-L-2,4-diaminobutyric acid through hydrolysis of ectoine All chemicals used to synthesize the gamma and alpha forms of N-acetyl-l-2,4-diaminobutyric acid (ADABA) for EctC enzyme activity assays were purchased either from Sigma Aldrich (Steinheim, Germany), or Acros (Geel, Belgium). Alkaline hydrolysis of ectoine (284 mg, 2.0 mmol) was accomplished in aqueous KOH (50 mL, 0.1 M) for 20 h at 50°C. The reaction mixture was subsequently neutralized with perchloric acid (60% in water, 4 mL) and the precipitated potassium perchlorate was filtered off. Subsequently, the filtrate was concentrated under reduced pressure. Purification of the residue and separation of the formed compounds was then performed by repeated chromatography on a silica gel column (Merck silica gel 60) using a gradient of ethanol/25% ammonia/water 50:1:2–10:1:2 as eluent to yield pure N-γ-ADABA (192 mg, 1.20 mmol, 60%) and N-α-ADABA (32 mg, 0.20 mmol, 10%). The identity and purity of theses compounds was unequivocally established by thin-layer chromatography (TLC) and nuclear magnetic resonance (1H-NMR and 13C-NMR) spectroscopy (S1a and S1b Fig) as described on a Bruker AVIII-400 or DRX-500 NMR spectrometer. (i) Analytical data for N-γ-ADABA: TLC: Rf = 0.55 (ethanol/25% ammonia/water 7:1:2); 1H-NMR (400 MHz, D2O): δ = 3.71 (dd, 3J(H,H) = 7.6 Hz, 3J(H,H) = 5.6 Hz, 1H, CH), 3.41–3.24 (m, 2H, CH2), 2.15–2.01 (m, 2H, CH2), 1.99 (s, 3H, CH3) ppm; 13C-NMR (100 MHz, D2O): δ = 177.5 (CO), 177.0 (COOH), 55.3 (CH), 38.3 (CH2), 33.0 (CH2), 24.6 (CH3) ppm. (ii) Analytical data for N-α-ADABA: TLC: Rf = 0.38 (ethanol/25% ammonia/water 7:1:2); 1H-NMR (400 MHz, D2O): δ = 4.24 (dd, 3J(H,H) = 8.8 Hz, 3J(H,H) = 5.1 Hz, 1H, CH), 3.07–3.02 (m, 2H, CH2), 2.22–2.11 (m, 2H, CH2), 2.04 (s, 3H, CH3) ppm; 13C-NMR (100 MHz, D2O): δ = 180.1 (CO), 176.7 (COOH), 55.4 (CH), 39.6 (CH2), 32.5 (CH2), 24.7 (CH3) ppm. Bacterial strains, plasmids and media The nucleotide sequence of the ectC gene from S. alaskensis (genome accession number: NC_008048) was used as a template to obtain a codon-optimized ectC DNA sequence (Life Technologies, Darmstadt, Germany) for its expression in E. coli. The nucleotide sequence of the synthetic ectC gene was deposited in the NCBI database under accession number KR002036. The synthetic ectC gene was used to construct an expression plasmid (pNW12) that is based on the pASG-IBA3 vector (IBA GmbH, Göttingen, Germany). In plasmid pNW12, the ectC gene is fused at its 3’ end to a short open reading frame encoding a Strep-tag II affinity peptide (NWSHPQFEK). It is transcribed from the TetR-controlled tet promoter carried by the backbone of the pASG-IBA3 expression vector. De-repression of tet promoter activity can be triggered by adding the synthetic inducer AHT for the TetR repressor to the growth medium. The details of the construction of pNW12 have been reported. Plasmids carrying ectC genes were routinely maintained in the Escherichia coli strain DH5α (Invitrogen, Karlsruhe, Germany) on LB agar plates containing ampicillin (100 μg ml-1). Plasmid DNA was isolated by routine procedures. Minimal medium A (MMA) containing 0.5% (w/v) glucose as the carbon source, 0.5% (w/v) casamino acids, 1 mM MgSO4, and 3 mM thiamine was used to cultivate the E. coli strain BL21 carrying pNW12 for the overproduction of the (Sa)EctC protein and its mutant derivatives. No additional metal solution was added to the components of the original recipe of MMA. Site-directed mutagenesis of the ectC gene Variants of the codon-optimized ectC gene from S. alaskensis present on plasmid pNW12 were prepared by site-directed mutagenesis using the QuikChange Lightning Site-Directed Mutagenesis Kit (Agilent, Waldbronn, Germany) with custom synthesized DNA primers purchased from Microsynth AG (Lindau, Germany). The DNA sequence of the entire coding region of each mutant ectC gene was determined by Eurofins MWG (Ebersberg, Germany) to ensure the presence of the desired mutation and the absence of unwanted alterations. Details on the genetic changes introduced into ectC genes are listed in Table 1. Conversion of N-γ-ADABA into ectoine by (Sa)EctC mutant derivatives and their iron-content. Mutation\tAmino acid-substitution\tEctoine formed [mM]\tActivity (%)\tIron content of the protein preparation (mol %)\t \tWT\t-\t9.33 ± 0.28\t100\t92.1 ± 3.4\t \tTAT/GCT\tTyr-52/Ala\t2.53 ± 0.18\t27\t19.8 ± 1.7\t \tGAA/GCA\tGlu-57/Ala\t0.97 ± 0.16\t10\t4.3 ± 2.9\t \tGAA/GAT\tGlu-57/Asp\t6.19 ± 0.42\t66\t68.7 ± 5.1\t \tTAT/GCT\tTyr-85/Ala\t2.15 ± 0.28\t23\t8.3 ± 2.3\t \tTAT/TTT\tTyr-85/Phe\t2.74± 0.50\t29\t9.4 ± 3.9\t \tTAT/TGG\tTyr-85/Trp\t0.95 ± 0.08\t10\t5.1 ± 1.7\t \tCAT/GCT\tHis-93/Ala\t0.72 ± 0.09\t8\t4.5 ± 0.8\t \tCAT/AAT\tHis-93/Asn\t2.14 ± 0.31\t23\t12.9 ± 2.6\t \tTGG/GCG\tTrp-21/Ala\t2.41 ± 0.39\t26\t89.4 ± 4.5\t \tACG/GCC\tSer-23/Ala\t1.98 ± 0.42\t21\t91.6 ± 2.8\t \tACC/GCC\tThr-40/Ala\t1.12 ± 0.13\t12\t89.6 ± 2.2\t \tTGT/GCT\tCys-105/Ala\t0.96 ± 0.21\t10\t90.1 ± 1.6\t \tTGT/TCT\tCys-105/Ser\t7.81 ± 0.65\t84\t88.7 ± 3.1\t \tTTT/GCT\tPhe-107/Ala\t4.77 ± 0.10\t51\t87.9 ± 2.2\t \tTTT/TAT\tPhe-107/Tyr\t8.87 ± 0.62\t95\t90.9 ± 3.9\t \tTTT/TGG\tPhe-107/Trp\t1.08 ± 0.27\t12\t72.6 ± 5.8\t \tCAT/GCT\tHis-117/Ala\t4.14 ± 0.27\t44\t82.9 ± 1.1\t \tCAT/GCT\tHis-55/Ala\t1.53 ± 0.19\t16\t15.4 ± 4.3\t \tGAA/GCA\tGlu-115/Ala\t1.92 ± 0.44\t21\t87.6 ± 4.4\t \tGAA/GAT\tGlu-115/Asp\t7.15 ± 0.60\t77\t88.0 ± 3.2\t \tCTG/GCG\tLeu-87/Ala\t5.81 ± 0.44\t62\t92.0 ± 1.3\t \tGAT/GCT\tAsp-91/Ala\t7.48 ± 0.81\t80\t89.6 ± 2.2\t \tGAT/GAA\tAsp-91/Glu\t9.13 ± 0.57\t98\t90.0 ± 1.9\t \tACC/GCC\tThr-41/Ala\t8.84 ± 0.63\t95\t91.7 ± 1.9\t \tCAT/GCT\tHis-51/Ala\t8.94 ± 0.47\t96\t90.2 ± 2.6\t \t The conversion of N-γ-ADABA into ectoine by the (Sa)EctC protein and its mutant derivatives was monitored in a reaction that contained 10 mM N-γ-ADABA as the substrate, 1 mM FeSO4 and 5 μg of the EctC protein under study. The amount of ectoine formed was measured after 20 min of incubation of the enzyme-substrate mixture by HPLC analysis. The iron-content of the investigated protein preparations was determined photometrically; note that in comparison with data obtained via ICP-MS, the colorimetric assay overestimates somewhat the iron content of the (Sa)EctC protein preparations. Overproduction and purification of recombinant EctC proteins For the overproduction of the (Sa)EctC-Strep-tag II protein, an overnight culture of strain [BL21 (pNW12)] was prepared in MMA and used to inoculate 1 L of MMA (in a 2 L Erlenmeyer flask) to an OD578 of 0.05. The cells were grown on an aerial shaker (set to 180 rpm) at 37°C until the culture reached an OD578 of 0.5. At this time point, the growth temperature was lowered to 30°C and the speed of the shaker was reduced to 100 rpm. Growth of the culture was continued and when it reached an OD578 of 0.7, AHT was added to the growth medium at a final concentration of 0.2 mg ml-1 to boost expression of the recombinant ectC gene. After 2 h of further incubation of the culture, the E. coli cells were harvested by centrifugation and disrupted by passing them several times through a French Pressure cell; a cleared cell lysate was prepared by ultracentrifugation (100 000 g) at 4°C for 1 h. The supernatant of this cleared lysate was then passed through a column filled with 5 ml of Strep-Tactin Superflow material (IBA GmbH, Göttingen, Germany); the column had been equilibrated with a buffer containing 200 mM NaCl and 20 mM Tris-HCl (pH 8). The (Sa)EctC-Strep-tag II protein was eluted from the affinity matrix with three column volumes of the same buffer containing 2.5 mM desthiobiotin. The recombinant (Sa)EctC-Strep-tag II protein was then concentrated to either 5 mg ml-1 for enzymes assays or 10 mg ml-1 for crystallization trials with Vivaspin 6 columns (Satorius Stedim Biotech GmbH, Göttingen, Germany) in the same buffer as described above. Desthiobiotin was not removed by dialysis from these protein preparations. The purified and concentrated (Sa)EctC-Strep-tag II protein was either used immediately for enzymes assays or kept at 4°C since the flash-freezing of the protein with liquid nitrogen and its subsequent storage at -80°C resulted in a rapid inactivation of ectoine synthase activity. 25 Variants of the (Sa)EctC-Strep-tag II protein carrying singe amino acid substitutions (Table 1) were overproduced and purified using the same procedure employed for the isolation of the wild-type protein. These mutant proteins behaved like the wild-type (Sa)EctC-Strep-tag II protein during the overproduction and purification procedure. Protein concentrations were determined both with a Pierce BCA Protein Assay Kit (Thermo Scientific, Schwerte, Germany) using BSA as the standard protein and spectrophotometrically by using an extinction coefficient of 15 470 M-1 cm-1 for the (Sa)EctC-Strep-tag II protein at a wavelength of 280 nm. The purity and integrity of the isolated (Sa)EctC-Strep-tag II proteins was inspected by SDS-polyacrylamide (15%) gel electrophoresis (SDS-PAGE). Molecular mass marker proteins for SDS-PAGE were purchased from LifeTechnologies (Darmstadt, Germany). Ectoine synthase enzyme activity assays The ectoine synthase activity of the (Sa)EctC protein was determined by HPLC-based enzyme assays. The initial enzyme activity assays were performed in a 30 μl-reaction volume for 20 min at 20°C. The used standard buffer (20 mM Tris, pH 8.0) contained 150 mM NaCl, 1 mM FeCl2, and 10 mM N-γ-ADABA. To determine optimal enzyme assay conditions for the (Sa)EctC-Strep-tag II protein, assay parameters and buffer conditions (e.g., the salt-concentrations, temperature, pH) were individually changed. The finally optimized assay buffer for ectoine synthase activity of the (Sa)EctC protein contained 20 mM Tris (pH 8.5), 200 mM NaCl, 1 mM FeCl2 and 10 mM N-γ-ADABA. Activity assays were run for 20 min at 15°C. Usually, 10 μg of the purified (Sa)EctC protein were added to start the enzyme assay. To assess the kinetic parameters of the ectoine synthase, varying concentrations of the substrates were used in the optimized assay buffer with a constant amount (10 μg) of the (Sa)EctC protein. The concentration of the natural EctC substrate N-γ-ADABA was varied between 0 and 40 mM, whereas that of N-α-ADABA was varied between 0 and 200 mM in the enzyme assays. Enzyme reactions were stopped by adding 30 μl of acetonitrile (100%) to the reaction vessel. The samples were centrifuged (13000 rpm, at room temperature for 5 min) to remove denatured proteins; the supernatant was subsequently analyzed for the formation of ectoine by HPLC analysis. Usually, 5- to 10-μl samples were injected into the HPLC system and the reaction product ectoine was analytically detected on a GROM-SIL Amino-1PR column (125 x 4 mm with a particle size of 3μm; purchased from GROM, Rottenburg-Hailfingen, Germany). Synthesis of ectoine by the purified (Sa)EctC-Strep-tag II protein and its mutant derivatives was monitored using a Infinity 1260 Diode Array Detector (DAD) (Agilent, Waldbronn, Germany) integrated into an Agilent 1260 Infinity LC system (Agilent). The ectoine content of the samples was quantified using the OpenLAB software suite (Agilent). The data shown for each ectC mutant (Table 1) were derived from two independent (Sa)EctC preparations, and each (Sa)EctC protein solution was assayed three times for its enzyme activity. Metal depletion and reconstitution of the (Sa)EctC protein To assess the dependency of the ectoine synthase for its enzyme activity on iron and other metals, purified and concentrated (Sa)EctC protein preparations (10 μM) were treated with different concentrations of EDTA for 10 minutes. They were subsequently dialyzed to remove the EDTA and the remaining (Sa)EctC enzyme activity was analyzed. To determine metal ion specificity of the ectoine synthase, 500 μl of the (Sa)EctC protein (100 μM) were initially treated with 1 mM EDTA for 10 minutes to obtain apo-(Sa)EctC protein preparations and the EDTA was then removed by dialysis. Enzyme activity assays with 10 μM of such protein preparations were then performed in the presence of either stoichiometric (10 μM) or excess amounts (1 mM) of FeCl2, FeCl3, ZnCl2, CoCl2, NiCl2, CuCl2, and MnCl2 to monitor metal ion specificity of the ectoine synthase. Prior to initiation of the enzyme reaction (by addition of the substrate), the (Sa)EctC protein solution was incubated with the different indicated metal ions for 10 minutes. Determination of the oligomeric state of (Sa)EctC protein To determine the oligomeric state of the (Sa)EctC protein in solution, we used high-performance liquid chromatography coupled to multi-angle light scattering detection (HPLC-MALS). A Bio SEC-5 HPLC column (Agilent Technologies Deutschland GmbH, Böblingen, Germany) with a pore size of 300 Å was equilibrated with 20 mM Tris-HCl (pH 7.5), 200 mM NaCl for high-performance liquid chromatography analysis. For these experiments, an Agilent Technologies system connected to a triple-angle light scattering detector (miniDAWN TREOS, Wyatt Technology Europe GmbH, Dernbach, Germany) followed by a differential refractive index detection system (Optilab t-rEX, Wyatt Technology) was used. Typically, 100 μl of purified (Sa)EctC protein (2 mg ml-1) was loaded onto the Bio SEC-5 HPLC column and the obtained data were analyzed with the ASTRA software package (Wyatt Technology). Determination of metal content of recombinant (Sa)EctC protein by ICP-MS The elemental contents of P, Fe, Ni, Cu and Zn of the (Sa)EctC-Strep-Tag-II protein sample were determined by inductive-coupled plasma mass spectrometry (ICP-MS) using an Agilent 7900 ICP-MS system equipped with a HEN nebulizer and cooled scott spray chamber under standard operating conditions. The isotopes 31P, 56Fe, 57Fe, 58Ni, 60Ni, 62Ni, 63Cu, 65Cu, 64Zn, 66Zn, and 67Zn were measured under NoGas, He collision and H2 reaction mode conditions. Some isotopes are strongly interfered from the matrix (mainly 56Fe, 63Cu) in the NoGas mode and are therefore rejected. The (Sa)EctC protein samples and buffer blanks were diluted 100-fold with ultra pure water and spiked with 10 μg kg-1 Y as the internal standard. The calibration of the ICP-MS was performed in the concentration range between 0.1 to 100 μg kg-1 using a homemade P standard solution prepared from titrimetrically analyzed H3PO4 solution and from dilutions of a Merck ICP multi-element standard solution IV (Merck No. 111355, Darmstadt, Germany). Photometric determination of non-heme-iron in (Sa)EctC and its mutant derivatives To determine the iron content in our (Sa)EctC-Strep-Tag-II preparations photometrically, 10 nmol of the purified proteins were heated at 80°C for 10 min in 250 μl of a 1% HCl solution. The reaction assay was cooled down on ice and then centrifuged (13000 rpm, 10 min at room temperature). The supernatant was transferred to a new reaction tube, and 750 μl H2O, 50 μl of 10% hydroxylamine/HCl, and 250 μl of 0.1% phenanthroline were added to the reaction vessel. After 30 min of incubation at room temperature, the absorbance of the solution was measured at 512 nm. 5 to 40 nmol of ammonium iron(II) sulfate were used for calibration of the assay. Crystallization of the (Sa)EctC protein Several conditions under which the (Sa)EctC protein formed crystals were found by using commercial screens (Nextal, Qiagen, Hilden, Germany; Molecular Dimensions, Suffolk, UK) in 96-well sitting drop plates (Corning 3553) at 12°C. Homogeneous (Sa)EctC protein (0.1 μl from a solution of 11 mg protein ml-1) was mixed with 0.1 μl reservoir solution and equilibrated against 50 μl reservoir solution. The most promising condition was found with a solution containing 0.05 M calcium acetate, 0.1 M sodium acetate (pH 4.5), and 40% (v/v) 1,2-propanediol from the Nextal Core IV suite (Qiagen, Hilden, Germany). A second condition under which (Sa)EctC crystallized was identified in microbatch setups (1 μl + 1 μl drops) using 20% (w/v) PEG 6000, 0.9 M lithium chloride, and 0.1 M citric acid (pH 5) from the Nextal Core II suite (Qiagen, Hilden, Germany). These conditions were optimized by grid screens around the initial condition and/or after the addition of tert-butanol as an additive. Large crystals were obtained either without any additive or after the addition of tert-butanol to the (Sa)EctC protein solution 30 minutes before the drops were spotted. Crystals reached their maximum dimensions of about 50 × 50 × 70 μm3 after 3–10 weeks. The crystals were fished after overlaying the drop with 2 μl mineral oil and flash frozen in liquid nitrogen. To obtain heavy atom derivatized crystals, methylmercury(II) chloride was added (final concentration: 0.5 mM) to the crystals in their drop for 30 minutes before they were fished and flash frozen in liquid nitrogen. Data processing and structure determination Native data sets were collected from a single crystal of (Sa)EctC obtained from the various crystallization trials at the ERSF beamline ID23eh2 (Grenoble, France) at 100 K. These data sets were processed using the XDS package and scaled with XSCALE showing a maximum resolution of 1.2 Å. To obtain initial phases of (Sa)EctC, a mercury-derivatized crystal was used to collect a conservative dataset at 2.8 Å resolution. The data were processed and scaled as described above, before the program AUTORICKSHAW using single isomorphous replacement (SIRAS), was used to localize the Hg atom, phase and built an initial model of the (Sa)EctC protein. This initial model was used as a template for molecular replacement on the 2.0 Å dataset revealing four monomers in the asymmetric unit. Once the 2.0 Å structure was refined, a single monomer of this structure was used as a template for molecular replacement to phase the 1.2 Å resolution dataset using the PHENIX software. Model building and refinement were performed using COOT, Refmac5 and Phenix_refine. Data refinement statistics and model content are summarized in Table 1. PDB accession numbers The atomic coordinates and structural factors have been deposited into the Protein Data Bank (PDB) (Brookhaven, USA) under the following accession codes: 5BXX (for the “semi-closed” (Sa)EctC structure) and 5BY5 (for the “open” (Sa)EctC structure). Figure preparation of crystal structures Figures of the crystal structures of SaEctC were prepared using the PyMol software suite (www.pymol.org). Results Overproduction, purification and oligomeric state of the ectoine synthase in solution We focused our biochemical and structural studies on the ectoine synthase from S. alaskensis [(Sa)EctC], a cold-adapted marine ultra-microbacterium, from which we recently also determined the crystal structure of the ectoine hydroxylase (EctD) in complex with either its substrate or its reaction product. We expressed a codon-optimized version of the S. alaskensis ectC gene in E. coli to produce a recombinant protein with a carboxy-terminally attached Strep-tag II affinity peptide to allow purification of the (Sa)EctC-Strep-Tag-II protein by affinity chromatography. The (Sa)EctC protein was overproduced and isolated with good yields (30–40 mg L-1 of culture) and purity (S2a Fig). Conventional size-exclusion chromatography (SEC) has already shown that (Sa)EctC preparations produced in this fashion are homogeneous and that the protein forms dimers in solution. High performance liquid chromatography coupled with multi-angle light-scattering detection (HPLC-MALS) experiments carried out here confirmed that the purified (Sa)EctC protein was mono-disperse and possessed a molecular mass of 33.0 ± 2.3 kDa (S2b Fig). This value corresponds very well with the theoretically calculated molecular mass of an (Sa)EctC dimer (molecular mass of the monomer, including the Strep-tag II affinity peptide: 16.3 kDa). Such a quaternary assembly as dimer has also been reported for the EctC proteins from H. elongata and N. maritimus. Biochemical properties of the ectoine synthase The EctA-produced substrate of the ectoine synthase, N-γ-acetyl-L-2,4-diaminobutyric acid (N-γ-ADABA) (Fig 1), is commercially not available. We used alkaline hydrolysis of ectoine and subsequent chromatography on silica gel columns to obtain N-γ-ADABA in chemically highly purified form (S1a Fig). This procedure also yielded the isomer of N-γ-ADABA, N-α-acetyl-L-2,4-diaminobutyric acid (N-α-ADABA) (S1b Fig). N-α-ADABA has so far not been considered as a substrate for EctC, but microorganisms that use ectoine as a nutrient produce it as an intermediate during catabolism. Using N-γ-ADABA as the substrate, we initially evaluated a set of biochemical parameters of the recombinant (Sa)EctC protein. S. alaskensis, from which the studied ectoine synthase was originally derived, is a microorganism that is well-adapted to a life in permanently cold ocean waters. Consistent with the physicochemical attributes of this habitat, the (Sa)EctC protein was already enzymatically active at 5°C, had a temperature optimum of 15°C and was able to function over a broad range of temperatures (S3a Fig). It possessed an alkaline pH optimum of 8.5 (S3b Fig), a value similar to the ectoine synthases from the halo-tolerant H. elongata (pH optimum of 8.5 to 9.0), the alkaliphile M. alcaliphilum (pH optimum of 9.0), and the acidophile Acidiphilium cryptum (pH optimum of 8.5 to 9.0), whereas the EctC protein from N. maritimus has a neutral pH optimum (pH 7.0). The salinity of the assay buffer had a significant influence on the maximal enzyme activity of the (Sa)EctC protein. An increase in either the NaCl or the KCl concentration led to an approximately 5-fold enhancement of the ectoine synthase activity. The maximum enzyme activity of (Sa)EctC occurred around 250 mM NaCl or KCl, respectively. (Sa)EctC is a highly salt-tolerant enzyme since it exhibited substantial enzyme activity even at NaCl and KCl concentrations of 1 M in the assay buffer (S3c and S3d Fig). The stimulation of EctC enzyme activity by salts has previously also been observed for other ectoine synthases. The ectoine synthase is a metal-containing protein Considerations based on bioinformatics suggests that EctC belongs to the cupin superfamily. Most of these proteins contain catalytically important transition state metals such as iron, copper, zinc, manganese, cobalt, or nickel. Cupins contain two conserved motifs: G(X)5HXH(X)3,4E(X)6G and G(X)5PXG(X)2H(X)3N (the letters in bold represent those residues that often coordinate the metal). Inspection of a previous alignment of the amino acid sequences of 440 EctC-type proteins revealed that the canonical metal-binding motif(s) of cupin-type proteins is not conserved among members of the extended ectoine synthase protein family. An abbreviated alignment of the amino acid sequence of EctC-type proteins is shown in Fig 2. Abbreviated alignment of EctC-type proteins. The amino acid sequences of 20 selected EctC-type proteins are compared. Strictly conserved amino acid residues are shown in yellow. Dots shown above the (Sa)EctC protein sequence indicate residues likely to be involved in iron-binding (red), ligand-binding (green) and stabilization of the loop-architecture (blue). The conserved residue Tyr-52 with so-far undefined functions is indicated by a green dot circled in red. Secondary structural elements (α-helices and β-sheets) found in the (Sa)EctC crystal structure are projected onto the amino acid sequences of EctC-type proteins. Since variations of the above-described metal-binding motif occur frequently, we experimentally investigated the presence and nature of the metal that might be contained in the (Sa)EctC protein by inductive-coupled plasma mass spectrometry (ICP-MS). For this analysis we used recombinant (Sa)EctC preparations from three independent protein overproduction and purification experiments. The ICP-MS analyses yielded an iron content of 0.66 ± 0.06 mol iron per mol of protein and the used (Sa)EctC protein preparations also contained a minor amount of zinc (0.08 mol zinc per mol of protein). All other assayed metals (copper and nickel) were only present in trace amounts (0.01 mol metal per mol of protein, respectively). The presence of iron in these (Sa)EctC protein preparations was further confirmed by a colorimetric method that is based on an iron-complexing reagent; this procedure yielded an iron-content of 0.84 ± 0.05 mol per mol of (Sa)EctC protein. Hence, both ICP-MS and the colorimetric method clearly established that the recombinantly produced ectoine synthase from S. alaskensis is an iron-containing protein. We note in this context, that the values obtained for the iron content of the (Sa)EctC proteins varied by approximately 10 to 20% between the two methods. The reason for this difference is not known, but indicates that the well established colorimetric assay probably overestimates the iron content of (Sa)EctC protein preparations to a certain degree. A metal cofactor is important for the catalytic activity of EctC The iron detected in the (Sa)EctC protein preparations could serve a structural role, or most likely, could be critical for enzyme catalysis as is the case for many members of the cupin superfamily. To address these questions, we incubated the (Sa)EctC enzyme with increasing concentrations of the metal chelator ethylene-diamine-tetraacetic-acid (EDTA) and subsequently assayed ectoine synthase activity. The addition of very low concentrations of EDTA (0.05 mM) to the EctC enzyme already led to a noticeable inhibition of the ectoine synthase activity and the presence of 1 mM EDTA completely inhibited the enzyme (Fig 3a). Dependency of the ectoine synthase activity on metals. (a) Impact of the iron-chelator EDTA on the enzyme activity of the purified (Sa)EctC protein. Metal depletion and reconstitution experiments with (b) stoichiometric and (c) excess amounts of metals. The (Sa)EctC protein was present at a concentration of 10 μM. The level of enzyme activity given in (b) is benchmarked relative to that of ectoine synthase enzyme assays in which 1 mM FeCl2 was added. We then took such an inactivated enzyme preparation, removed the EDTA by dialysis, and added stoichiometric amounts (10 μM) of various metals to the (Sa)EctC enzyme. The addition of FeCl2 to the enzyme assay restored enzyme activity to about 38%, whereas the addition of ZnCl2 or CoCl2 rescued (Sa)EctC enzyme activity only to 5% and 3%, respectively. All other tested metals, including Fe3+, were unable to restore activity (Fig 3b). When the concentration of the various metals in the enzyme assay was increased 100-fold, Fe2+ exhibited again the strongest stimulating effect on enzyme activity, and rescued enzyme activity to a degree similar to that exhibited by (Sa)EctC protein preparations that had not been inactivated through EDTA treatment (Fig 3c). However, a large molar excess of other transition-state metals (zinc, cobalt, nickel, copper, and manganese) typically found in members of the cupin superfamily allowed the partial rescue of ectoine synthase activity as well (Fig 3c). This is in line with literature data showing that cupin-type enzymes are often promiscuous with respect to the use of the catalytically important metal. Kinetic parameters of EctC for N-γ-ADABA and N-α-ADABA Based on the data presented in S3 Fig, we formulated an optimized activity assay for the ectoine synthase of S. alaskensis and used it to determined the kinetic parameters for the (Sa)EctC enzyme for both its natural substrate N-γ-ADABA and the isomer N-α-ADABA. The EctC-catalyzed ring-closure of N-γ-ADABA to form ectoine exhibited Michaelis-Menten-kinetics with an apparent Km of 4.9 ± 0.5 mM, a vmax of 25.0 ± 0.8 U/mg and a kcat of 7.2 s-1 (S4a Fig). Given the chemical relatedness of N-α-ADABA to the natural substrate (N-γ-ADABA) of the ectoine synthase (S1a and S1b Fig), we wondered whether (Sa)EctC could also use N-α-ADABA to produce ectoine. This was indeed the case. (Sa)EctC catalyzed this reaction with Michaelis-Menten-kinetics exhibiting an apparent Km of 25.4 ± 2.9 mM, a vmax of 24.6 ± 1.0 U/mg and a kcat 0.6 s-1 (S4b Fig). Hence, N-α-ADABA is a newly recognized substrate for ectoine synthase. However, both the affinity (Km) of the (Sa)EctC protein and its catalytic efficiency (kcat/Km) were strongly reduced in comparison with N-γ-ADABA. The Km dropped fife-fold from 4.9 ± 0.5 mM to 25.4 ± 2.9 mM, and the catalytic efficiency was reduced from 1.47 mM-1 s-1 to 0.02 mM-1 s-1, a 73-fold decrease. Both N-γ-ADABA and N-α-ADABA are concomitantly formed during the enzymatic hydrolysis of the ectoine ring during catabolism. Our finding that N-α-ADABA is a substrate for ectoine synthase has bearings for an understanding of the physiology of those microorganisms that can both synthesize and catabolize ectoine. However, these types of microorganisms should still be able to largely avoid a futile cycle since the affinity of ectoine synthase for N-γ-ADABA and N-α-ADABA, and its catalytic efficiency for the two compounds, differs substantially (S4a and S4b Fig). Crystallization of the (Sa)EctC protein Since no crystal structure of ectoine synthase has been reported, we set out to crystallize the (Sa)EctC protein. Attempts to obtain crystals of (Sa)EctC in complex either with its substrate N-γ-ADABA or its reaction product ectoine were not successful. However, two crystal forms of the (Sa)EctC protein in the absence of the substrate were obtained. Crystal form A diffracted to 1.2 Å and had a unit cell of a = 72.71 b = 72.71 c = 52.33 Å and α = 90 β = 90 γ = 120° displaying a P3221 symmetry (S1 Table). Crystal form B diffracted to 2.0 Å and had a unit cell of a = 97.52 b = 43.96 c = 138.54 Å and α = 90 β = 101.5 γ = 120° and displayed a C2 symmetry (S1 Table). Attempts to solve the crystal structure of the (Sa)EctC protein by molecular replacement has previously failed. However, we were able to obtain crystals of form B that were derivatized with mercury and these diffracted up to 2.8 Å (S1 Table). This dataset was used to derive an initial structural model of the (Sa)EctC protein, which in turn was employed as a template for molecular replacement to phase the native dataset (2.0 Å) of crystal form B. After several rounds of manual model building and refinement, four monomers of (Sa)EctC were identified and the crystal structure was refined to a final Rcryst of 21.1% and an Rfree of 24.8% (S1 Table). Finally, a monomer of this structure was used as a template for molecular replacement to phase the high-resolution (1.2 Å) dataset of crystal form A, which was subsequently refined to a final Rcryst of 12.4% and an Rfree of 14.9% (S1 Table). Overall fold of the (Sa)EctC protein The two EctC structures that we determined revealed that the ectoine synthase belongs to the cupin superfamily with respect to its overall fold (Fig 4a–4c). However, they represent two different states of the 137 amino acids comprising (Sa)EctC protein (Fig 2). First, the 1.2 Å structure reveals the spatial configuration of the (Sa)EctC protein ranging from amino acid Met-1 to Glu-115; hence, it lacks 22 amino acids at the carboxy-terminus of the authentic (Sa)EctC protein. This structure adopts an open conformation with respect to the typical fold of cupin barrels and is therefore termed in the following the “open” (Sa)EctC structure (Fig 4b). In this structure no metal co-factor was identified. The second crystal structure of the (Sa)EctC protein was solved at a resolution of 2.0 Å and contained four molecules of the protein in the asymmetric unit of which protomer A comprised amino acid Met-1 to Gly-121 and adopts a closed conformation. Hence, it still lacks 16 amino acid residues of the carboxy-terminus of the authentic 137 amino acids comprising (Sa)EctC protein (Fig 2). We therefore cannot exclude that this crystal structure does not represent the fully closed state of the ectoine synthase; consequently, we tentatively termed it the “semi-closed” (Sa)EctC structure. Interestingly, the three other monomers present in the asymmetric unit all range from Met-1 to Glu-115 and adopt a conformation similar to the “open” EctC structure. Overall structure of the “open” and “semi-closed” crystal structures of (Sa)EctC. (a) The overall structure of the “semi-closed” (Sa)EctC resolved at 2.0 Å is depicted in green in a cartoon (upper panel) and surface (lower panel) representation. The β-strands are numbered β1-β11 and the helices α-I to α-II. (b) The overall structure of the “open” (Sa)EctC was resolved at 1.2 Å and is depicted in yellow in a cartoon (upper panel) and surface (lower panel) representation. The entrance to the active site of the ectoine synthase is marked. (c) Overlay of the “semi-closed” and “open” (Sa)EctC structures. The overall structure of (Sa)EctC is basically the same in both crystals except for the carboxy-terminus, which covers the entry of one side of the cupin barrel from the surroundings in monomer A in the “semi-closed” structure. This is reflected by the calculated root mean square deviation (RMSD) of the Cα atoms that was about 0.56 Å (over 117 residues) when the four “open” monomers were compared with each other. However, the “semi-closed” monomer has a slightly higher RMSD of 1.4 Å (over 117 residues) when compared with the “open” 2.0 Å structure. Therefore, we describe in the following the overall structure for the “semi-closed” form of the (Sa)EctC protein and subsequently highlight the structural differences between the “open” and “semi-closed” forms in more detail. The structure of the “semi-closed” (Sa)EctC protein consists of 11 β-strands (β1-β11) and two α-helices (α-I and α-II) (Fig 4a). The β-strands form two anti-parallel β-sheets: β2 β3, β4, β11, β6, and β9, and a smaller three-stranded β-sheet (β7, β8, and β10), respectively. These two β-sheets pack against each other, forming a cup-shaped β-sandwich with a topology characteristic for the cupin-fold. Hence, (Sa)EctC adopts an overall bowl shape in which one side is opened towards the solvent (Fig 4a to 4c). In the “semi-closed” structure, a longer carboxy-terminal tail is visible in the electron density, folding into a small helix (α-II) that closes the active site of the (Sa)EctC protein (Fig 4a). The formation of this α-II helix induces a reorientation and shift of a long unstructured loop (as observed in the “open” structure) connecting β4 and β6, resulting in the formation of the stable β-strand β5 as observed in the “semi-closed”state of the (Sa)EctC protein (Fig 4a). Structural comparison analyses using the DALI server revealed that (Sa)EctC adopts a fold similar to other members of the cupin superfamily. The highest structural similarities are observed for the Cupin 2 conserved barrel domain protein (YP_751781.1) from Shewanella frigidimarina (PDB accession code: 2PFW) with a Z-score of 13.1 and an RMSD of 2.2 Å over 104 Cα-atoms (structural data for this protein have been deposited in the PDB but no publication connected to this structure is currently available), a manganese-containing cupin (TM1459) from Thermotoga maritima (PDB accession code: 1VJ2) with a Z-score of 12.8 and an RMSD of 2.0 Å over 103 Cα-atoms, the cyclase RemF from Streptomyces resistomycificus (PDB accession code: 3HT1 with a Z-score of 11.9 and an RMSD of 1.9 Å over 102 Cα-atoms), and an auxin-binding protein 1 from Zea mays (PDB accession code: 1LR5) with an Z-score of 11.8 and an RMSD of 2.8 Å over 104 Cα-atoms). Our data classify EctC, in addition to the polyketide cyclase RemF, as the second known cupin-related enzyme that catalyze a cyclocondensation reaction. Next to RemF and the aldos-2-ulose dehydratase/isomerase, the ectoine synthase is only the third characterized dehydratase within the cupin superfamily. Analysis of the EctC dimer interface as observed in the (Sa)EctC crystal structure Both the SEC analysis and the HPLC-MALS experiments (S2b Fig) have shown that the ectoine synthase from S. alaskensis is a dimer in solution. The crystal structure of this protein reflects this quaternary arrangement. In the “semi-closed” crystal structure, (Sa)EctC has crystallized as a dimer of dimers within the asymmetric unit. This dimer (Fig 5a and 5b) is composed of two monomers arranged in a head-to-tail orientation and is stabilized via strong interactions mediated by two antiparallel β-strands, β-strand β1 (sequence 1MIVRN5) from monomer A and β-strand β8 from monomer B (sequence 82GVMYAL87) (Fig 5c). The strong interactions between these β-strands rely primarily on backbone contacts. In addition to these interactions, some weaker hydrophobic interactions are also observed between the two monomers in some loops connecting the β-strands. As calculated with PDBePISA, the surface area buried upon dimer formation is 1462 Å2, which is 20.5% of the total accessible surface of a monomer of this protein. Both values fall within the range for known functional dimers. Crystal structure of (Sa)EctC. (a) Top-view of the dimer of the (Sa)EctC protein. The position of the water molecule, described in detail in the text, is shown in one of the monomers as an orange sphere. (b) Side-view of a (Sa)EctC dimer allowing an assessment of the dimer interface formed by two β-strands of each monomer. (c) Close-up representation of the dimer interface mediated by beta-strand β1 and β6. In the “open” (Sa)EctC structure, one monomer is present in the asymmetric unit. We therefore inspected the crystal packing and analyzed the monomer-monomer interactions with symmetry related molecules to elucidate whether a physiologically relevant dimer could be deduced from this crystal form as well. Indeed, a similar dimer configuration to the one described for the “semi-closed” (Sa)EctC structure is observed with the same monomer-monomer interactions mediated by the two β-sheets. The crystallographic two-fold axis present within the crystal symmetry is located exactly in between the two monomers, resulting in a monomer within the asymmetric unit. Hence, the same dimer observed in the “semi-closed” structure of (Sa)EctC can also be observed in the “open” structure. Interestingly, the proteins identified by the above-described DALI search not only have folds similar to EctC, but are also functional dimers that adopt similar monomer-monomer interactions within the dimer assembly as deduced from the inspection of the corresponding PDB files (2PFW, 3HT1, 1VJ2, 1LR5). Structural rearrangements of the flexible (Sa)EctC carboxy-terminus The cupin core represents the structural framework of ectoine synthase (Figs 4 and 5). The major difference in the two crystal structures of the (Sa)EctC protein reported here is the orientation of the carboxy-terminus. Some amino acids located in the carboxy-terminal region of the 137 amino acids comprising (Sa)EctC protein are highly conserved (Fig 2) within the extended EctC protein family. At the end of β-strand β11, two consecutive conserved proline residues (Pro-109 and Pro-110) are present that are responsible for a turn in the main chain of the (Sa)EctC protein. In the “semi-closed” (Sa)EctC structure, the visible electron density of the carboxy-terminus is extended by 7 amino acid residues and ends at position Gly-121. These additional amino acids fold into a small helix, which seals the open cavity of the cupin-fold of the (Sa)EctC protein (Fig 4a). Furthermore, this helix is stabilized via interactions with the loop region between β-strands β4 and β6, thereby inducing a structural rearrangement. This induces the formation of β-strand β5, which is not present when the small C-terminal helix is absent as observed in the “open” (Sa)EctC structure. As a result, the newly formed β-strand β5 is reoriented and moved by 2.4 Å within the “semi-closed” (Sa)EctC structure (Fig 4a to 4c). It is worth mentioning that β-strand β5 is located next to His-93, which in all likelihood involved in metal binding (see below). The position of this His residue is slightly shifted in both (Sa)EctC structures, likely the result of the formation of β-strand β5. Therefore the sealing of the cupin fold, as described above, seem to have an indirect influence on the architecture of the postulated iron-binding site. The consecutive Pro-109 and Pro-110 residues found at the end of β-strand β11are highly conserved in EctC-type proteins (Fig 2). They are responsible for redirecting the main chain of the remaining carboxy-terminus (27 amino acid residues) of (Sa)EctC to close the cupin fold. In the “semi-closed” structure this results in a complete closure of the entry of the cupin barrel (Fig 4a to 4c). In the “open” (Sa)EctC structure, both proline residues are visible in the electron density; however, almost directly after Pro-110, the electron density is drastically diminished caused by the flexibility of the carboxy-terminus. A search for partners interacting with Pro-109 revealed that it interacts via its backbone oxygen with the side chain of His-55 as visible in both the “open” and “semi-closed” (Sa)EctC structures. The Pro-109/His-55 interaction ensures the stable orientation of both proline residues at the end of β-strand β11. Since these proline residues are followed by the carboxy-terminal region of the (Sa)EctC protein, the interaction of His-55 with Pro-109 will likely play a substantial role in spatially orienting this very flexible part of the protein. In addition to the interactions between Pro-109 and His-55, the carboxy-terminal region of (Sa)EctC is held in position via an interaction of Glu-115 with His-55, which stabilizes the conformation of the small helix in the carboxy-terminus further. The interaction between Glu-115 and His-55 is only visible in the “semi-closed” structure where the partially extended carboxy-terminus is resolved in the electron density. In the “open” structure of the (Sa)EctC protein, this interaction does not occur since Glu-115 is rotated outwards (Fig 6a and 6b). Hence, one might speculate that this missing interaction might be responsible for the flexibility of the carboxy-terminus in the “open” (Sa)EctC structure and consequently results in less well defined electron density in this region. Architecture of the presumed metal-binding site of the (Sa)EctC protein and its flexible carboxy-terminus. (a) The described water molecule (depicted as orange sphere) is bound via interactions with the side chains of Glu-57, Tyr-85, and His-93. The position occupied by this water molecule represents probably the position of the Fe2+ cofactor in the active side of the ectoine synthase. His-55 interacts with the double proline motif (Pro-109 and Pro-110). It is further stabilized via an interaction with the side chain of Glu-115 which is localized in the flexible carboxy-terminus (colored in orange) of (Sa)EctC that is visible in the “semi-closed” (Sa)EctC structure. (b) An overlay of the “open” (colored in light blue) and the “semi-closed” (colored in green) structure of the (Sa)EctC protein. The putative iron binding site of (Sa)EctC In the “semi-closed” structure of (Sa)EctC, each of the four monomers in the asymmetric unit contains a relative strong electron density positioned within the cupin barrel. Since (Sa)EctC is a metal containing protein (Fig 3), we tried to fit either Fe2+, or Zn2+ ions into this density and also refined occupancy. Only the refinement of Fe2+ resulted in a visibly improved electron density, however with a low degree of occupancy. This possible iron molecule is bound via interactions with Glu-57, Tyr-85 and His-93 (Fig 6a and 6b). The distance between the side chains of these residues and the (putative) iron co-factor is 3.1 Å for Glu-57, 2.9 Å for Tyr-85, and 2.9 Å for His-93, respectively. These distances are to long when compared to other iron binding sites, a fact that might be caused by the absence of the proper substrate in the (Sa)EctC crystal structure. Since both the refinement and the distance did not clearly identify an iron molecule, we decided to conservatively place a water molecule at this position. The position of this water molecule is described in more detail below and is highlighted in Figs 5a and 5b and 6a and 6b as a sphere. Interestingly, all three amino acids coordinating this water molecule are strictly conserved within an alignment of 440 members of the EctC protein family (for an abbreviated alignment of EctC-type proteins see Fig 2). In the “open” structure of the (Sa)EctC protein, electron density is visible where the presumptive iron is positioned in the “semi-closed” structure. However, this electron density fits perfectly to a water molecule and not to an iron, and the water molecule was clearly visible after the refinement at this high resolution (1.2 Å) of the “open” (Sa)EctC structure. In a superimposition of both (Sa)EctC crystal structures, the spatial arrangements of the side chains of the three amino acids (Glu-57, Tyr-85, and His-93) likely to contact the iron in the “semi-closed” structure match nicely with those of the corresponding residues of the “iron-free” “open” structure (Fig 6b). Only His-93 is slightly rotated inwards in the “semi-closed” structure, most likely due to formation of β-strand β5 as described above. Taken together, this observations indicate, that the architecture of the presumptive iron-binding site is pre-set for the binding of the catalytically important metal by the ectoine synthase. Of note is the different spatial arrangement of the side-chain of Tyr-52 (located in a loop after the end of β-strand β5) in the “open” and “semi-closed” (Sa)EctC structures. In the “semi-closed” structure, the hydroxyl-group of the side-chain of Tyr-52 points towards the iron (Fig 6a and 6b), but the corresponding distance (3.9 Å) makes it highly unlikely that Tyr-52 is directly involved in metal binding. Nevertheless, its substitution by an Ala residue causes a strong decrease in iron-content and enzyme activity of the mutant protein (Table 1). It becomes apparent from an overlay of the “open” and “semi-closed” (Sa)EctC crystal structures that the side-chain of Tyr-52 rotates away from the position of the presumptive iron, whereas the side-chains of those residues that probably contacting the metal directly [Glu-57, Tyr-85, and His-93], remain in place (Fig 6a and 6b). Since Tyr-52 is strictly conserved in an alignment of 440 EctC-type proteins (Fig 2), we speculate that it might be involved in contacting the substrate of the ectoine synthase and that the absence of N-γ-ADABA in our (Sa)EctC crystal structures might endow the side chain of Tyr-52 with extra spatial flexibility. To further analyze the putative iron binding site (Fig 6a), we performed structure-guided site-directed mutagenesis and assessed the resulting (Sa)EctC variants for their iron content and studied their enzyme activity. When those three residues (Glu-57, Tyr-85, His-93) that likely form the mono-nuclear iron center in the (Sa)EctC crystal structure were individually replaced by an Ala residue, both the catalytic activity and the iron content of the mutant proteins was strongly reduced (Table 1). For some of the presumptive iron-coordinating residues, additional site-directed mutagenesis experiments were carried out. To verify the importance of the negative charge in the position of Glu-57, we created an Asp variant. This mutant protein rescued the enzyme activity and iron content of the Ala substitution substantially (Table 1). We also replaced Tyr-85 with either a Phe or a Trp residue and both mutant proteins largely lost their catalytic activity and iron content (Table 1) despite the fact that these substitutions were conservative. Collectively, these data suggest that the hydroxyl group of the Tyr-85 side chain is needed for the binding of the iron (Fig 6a). We also replaced the presumptive iron-binding residue His-93 by an Asn residue, yielding a (Sa)EctC protein variant that possessed an enzyme activity of 23% and iron content of only 14% relative to that of the wild-type protein (Table 1). Collectively, the data addressing the functionality of the putative iron-coordinating residues (Glu-57, Tyr-85, His-93) buttress our notion that the Fe2+ present in the (Sa)EctC protein is of catalytic importance. A chemically undefined ligand in the (Sa)EctC structure provides clues for the binding of the N-γ-ADABA substrate Despite considerable efforts, either by trying co-crystallization or soaking experiments, we were not able to obtain a (Sa)EctC crystal structures that contained either the substrate N-γ-ADABA, or ectoine, the reaction product of ectoine synthase (Fig 1). However, in the “semi-closed” (Sa)EctC structure where the carboxy-terminal loop is largely resolved, a long stretched electron density feature was detected in the predicted active site of the enzyme; it remained visible after crystallographic refinement. This is in contrast to the high-resolution “open” structure of the (Sa)EctC protein where no additional electron density was observed after refinement. We tried to fit all compounds used in the buffers during purification and crystallization into the observed electron density, but none matched. This observation indicates that the chemically undefined ligand was either trapped by the (Sa)EctC protein during its heterologous production in E. coli or during crystallization. Since we used PEG molecules in the crystallization conditions, the observed density might stem from an ordered part of a PEG molecule, or low molecular weight PEG species that might have been present in the PEG preparation used in our experiments. We therefore stress that we cannot identify neither the true chemically nature of this compound, nor its precise origin. Estimating from the dimensions of the electron density feature, we modeled the chemically undefined compound trapped by the (Sa)EctC protein as a hexane-1,6-diol molecule (PDB identifier: HEZ) to best fit the observed electron density. However, to the best of our knowledge, hexane-1,6-diol is not part of the E. coli metabolome. Despite these notable limitations, we considered the serendipitously trapped compound as a mock ligand that might provide useful insights into the spatial positioning of the true EctC substrate and those residues that coordinate it within the ectoine synthase active site. We note that both N-γ-ADABA and hexane-1,6-diol are both C6-compounds and display similar length (Fig 7a). A chemically undefined ligand is captured in the active site of the “semi-closed” (Sa)EctC crystal structure. (a) The observed electron density in the active site of the “semi-closed” structure of (Sa)EctC is modeled as a hexane-1,6-diol molecule and compared with the electron density of the N-γ-ADABA substrate of the ectoine synthase to emphasize the similarity in size of these compounds. (b) The presumable binding site of the iron co-factor and of the modeled hexane-1,6-diol molecule is depicted. The amino acid side chains involved in iron-ligand binding are colored in blue and those involved in the binding of the chemically undefined ligand are colored in green using a ball and stick representation. The flexible carboxy-terminal loop of (Sa)EctC is highlighted in orange. The electron density was calculated as an omit map and contoured at 1.0 σ. We refined the (Sa)EctC structure with the trapped compound, and by doing so, the refinement parameters (especially R- and Rfree-factor) dropped by 1.5%. We also calculated an omit map and the electron density reappeared (Fig 7b). When analyzing the interactions of this compound within the (Sa)EctC protein, we found that it is bound via interactions with Trp-21 and Ser-23 of β-sheet β3, Thr-40 located in β-sheet β4, and Cys-105 and Phe-107, which are both part of β-sheet β11. Remarkably, all of these residues are highly conserved throughout the extended EctC protein family (Fig 2). Structure-guided site-directed mutagenesis of the catalytic core of the ectoine synthase In a previous alignment of the amino acid sequences of 440 EctC-type proteins, 13 amino acids were identified as strictly conserved residues. These correspond to amino acids Thr-40, Tyr-52, His-55, Glu-57, Gly-64, Tyr-85- Leu-87, His-93, Phe-107, Pro-109, Gly-113, Glu-115, and His-117 in the (Sa)EctC protein (Fig 2). Amino acid residues Gly-64, Pro-109, and Gly-113 likely fulfill structural roles since they are positioned either at the end or at the beginning of β-strands and α-helices. We considered the remaining ten residues as important either for ligand binding, for catalysis, or for the structurally correct orientation of the flexible carboxy-terminus of the (Sa)EctC protein. As described above, the side chains of Glu-57, Tyr-85, and His-93 are probably involved in iron binding (Table 1 and Fig 6a). In view of the (Sa)EctC structure with the serendipitously trapped compound (Fig 7b), we probed the functional importance of the seven residues that contact this ligand by structure-guided site-directed mutagenesis (Table 1). Each of these mutant (Sa)EctC proteins was overproduced in E. coli and purified by affinity chromatography; they all yielded pure and stable protein preparations. We benchmarked the activity of the (Sa)EctC variants in a single time-point enzyme assay under conditions where 10 μM of the wild-type (Sa)EctC protein converted almost completely the supplied 10 mM N-γ-ADABA substrate to 9.33 mM ectoine within a time frame of 20 min. In addition, we determined the iron content of each of the mutant (Sa)EctC protein by a colorimetric assay (Table 1). The side chains of the evolutionarily conserved Trp-21, Ser-23, Thr-40, Cys-105, and Phe-107 residues (Fig 2) make contacts with the chemically undefined ligand that we observed in the “semi-closed” (Sa)EctC structure (Fig 7b). We replaced each of these residues with an Ala residue and found that none of them had an influence on the iron content of the mutant proteins. However, their catalytic activity was substantially impaired (Table 1). Thr-40 is positioned on β-strand β5 and its side chain protrudes into the lumen of the cupin barrel formed by the (Sa)EctC protein (Fig 7b). We also replaced Phe-107 with either an Tyr or an Trp residue: the Phe-107/Tyr substitution possessed near wild-type enzyme activity (about 95%) and the full iron content, but the Phe-107/Trp substitution possessed only 12% enzyme activity and 72% iron content compared to the wild-type protein. The properties of these mutant proteins indicate that the aromatic side chain at position 107 of (Sa)EctC is of importance but that a substitution with a bulky aromatic side chain is strongly detrimental to enzyme activity and concomitantly moderately impairs iron binding. Replacement of the only Cys residue in (Sa)EctC (Cys-105; Fig 2) by a Ser residue, a configuration that is naturally found in two EctC proteins among 440 inspected amino acid sequences, yielded a (Sa)EctC variant with 84% wild-type activity and an iron content similar to that of the wild-type protein. However, the Cys-105/Ala variant was practically catalytically inactive while largely maintaining its iron content (Table 1). Since the side-chains of Cys residues are chemically reactive and often participate in enzyme catalysis, Cys-105 (or Ser-105) might serve such a role for ectoine synthase. We observed two amino acid substitutions that simultaneously strongly affected enzyme activity and iron content; these were the Tyr-52/Ala and the His-55/Ala (Sa)EctC protein variants (Table 1). Based on the (Sa)EctC crystal structures that we present here, we can currently not firmly understand why the replacement of Tyr-52 by Ala impairs enzyme function and iron content so drastically (Table 1). This is different for the His-55/Ala substitution. The carboxy-terminal region of the (Sa)EctC protein is held in its position via an interaction of Glu-115 with His-55, where His-55 in turn interacts with Pro-110 (Fig 6a and 6b). Each of these residues is evolutionarily highly conserved. The individual substitution of either Glu-115 or His-55 by an Ala residue is predicted to disrupt this interactive network and therefore should affect enzyme activity. Indeed, the Glu-115/Ala and the His-55/Ala substitutions possessed only 21% and 16% activity of the wild-type protein, respectively (Table 1). The Glu-115/Ala mutant possessed wild-type levels of iron, whereas the iron content of the His-55/Ala substitutions dropped to 15% of the wild-type level (Table 1). We also replaced Glu-115 with a negatively charged residue (Asp); this (Sa)EctC variant possessed wild-type levels of iron and still exhibited 77% of wild-type enzyme activity. Collectively, these data suggest that the correct positioning of the carboxy-terminus of the (Sa)EctC protein is of structural and functional importance for the activity of the ectoine synthase. Residues Leu-87 and Asp-91 are highly conserved in the ectoine synthase protein family. The replacement of Leu-87 by Ala led to a substantial drop in enzyme activity (Table 1). Conversely, the replacement of Asp-91 by Ala and Glu, resulted in (Sa)EctC protein variants with 80% and 98% enzyme activity, respectively (Table 1). We currently cannot comment on possible functional role Asp-91. However, Leu-87 is positioned at the end of one of the β-sheets that form the dimer interface (Fig 5c) and it might therefore possess a structural role. It is also located near Tyr-85, one of the residues that probably coordinate the iron molecule with in the (Sa)EctC active site (Fig 6a) and therefore might exert indirect effects. His-117 is a strictly conserved residue and its substitution by an Ala residue results in a drop of enzyme activity (down to 44%) and an iron content of 83% (Table 1). We note that His-117 is located close to the chemically undefined ligand in the (Sa)EctC structure (Fig 7b) and might thus play a role in contacting the natural substrate of the ectoine synthase. As an internal control for our mutagenesis experiments, we also substituted Thr-41 and His-51, two residues that are not evolutionarily conserved in EctC-type proteins with Ala residues. Both (Sa)EctC protein variants exhibited wild-type level enzyme activities and possessed a iron content matching that of the wild-type (Table 1). This illustrates that not every amino acid substitution in the (Sa)EctC protein leads to an indiscriminate impairment of enzyme function and iron content. Discussion The crystallographic data presented here firmly identify ectoine synthase (EctC), an enzyme critical for the production of the microbial cytoprotectant and chemical chaperone ectoine, as a new member of the cupin superfamily. The overall fold and bowl shape of the (Sa)EctC protein (Figs 4 and 5) with its 11 β-strands (β1-β11) and two α-helices (α-I and α-II) closely adheres to the design principles typically found in crystal structures of cupins. In addition to the ectoine synthase, the polyketide cyclase RemF is the only other currently known cupin-related enzyme that catalyze a cyclocondensation reaction although the substrates of EctC and RemF are rather different. As a consequence of the structural relatedness of EctC and RemF and the type of chemical reaction these two enzymes catalyze, is now understandable why bona fide EctC-type proteins are frequently (mis)-annotated in microbial genome sequences as “RemF-like” proteins. The pro- and eukaryotic members of the cupin superfamily perform a variety of both enzymatic and non-enzymatic functions that are built upon a common structural scaffold. Most cupins contain transition state metals that can promote different types of chemical reactions. Except for some cupin-related proteins that seem to function as metallo-chaperones, the bound metal is typically an essential part of the active sites. We report here for the first time that the ectoine synthase is a metal-dependent enzyme. ICP-MS, metal-depletion and reconstitution experiments (Fig 3) consistently identify iron as the biologically most relevant metal for the EctC-catalyzed cyclocondensation reaction. However, as observed with other cupins, EctC is a somewhat promiscuous enzyme as far as the catalytically important metal is concerned when they are provided in large molar excess (Fig 3c). Although some uncertainty remains with respect to the precise identity of amino acid residues that participate in metal binding by (Sa)EctC, our structure-guided site-directed mutagenesis experiments targeting the presumptive iron-binding residues (Fig 6a and 6b) demonstrate that none of them can be spared (Table 1). The architecture of the metal center of ectoine synthase seems to be subjected to considerable evolutionary constraints. The three residues (Glu-57, Tyr-85, His-93) that we deem to form it (Figs 6 and 7b) are strictly conserved in a large collection of EctC-type proteins originating from 16 bacterial and three archaeal phyla (Fig 2). We also show here for the first time that, in addition to its natural substrate N-γ-ADABA, EctC also converts the isomer N-α-ADABA into ectoine, albeit with a 73-fold reduced catalytic efficiency (S3a and S3b Fig). Hence, the active site of ectoine synthase must possess a certain degree of structural plasticity, a notion that is supported by the report on the EctC-catalyzed formation of the synthetic compatible solute ADPC through the cyclic condensation of two glutamine molecules. Our finding that N-α-ADABA serves as a substrate for ectoine synthase has physiologically relevant ramifications for those microorganisms that can both synthesize and catabolize ectoine, since they need to prevent a futile cycle of synthesis and degradation when N-α-ADABA is produced as an intermediate in the catabolic route. Although we cannot identify the true chemical nature of the C6 compound that was trapped in the (Sa)EctC structure nor its precise origin, we treated this compound as a proxy for the natural substrate of ectoine synthase, which is a C6 compound as well (Fig 7a). We assumed that its location and mode of binding gives, in all likelihood, clues as to the position of the true substrate N-γ-ADABA within the EctC active site. Indeed, site-directed mutagenesis of those five residues that contact the unknown C6 compound (Fig 7b) yielded (Sa)EctC variants with strongly impaired enzyme function but near wild-type levels of iron (Table 1). This set of data and the fact that the targeted residues are strongly conserved among EctC-type proteins (Fig 2) is consistent with their potential role in N-γ-ADABA binding or enzyme catalysis. We therefore surmise that our crystallographic data and the site-directed mutagenesis study reported here provide a structural and functional view into the architecture of the EctC active site (Fig 7b). The ectoine synthase from the cold-adapted marine bacterium S. alaskensis can be considered as a psychrophilic enzyme (S3a Fig), types of proteins with a considerable structural flexibility. This probably worked to the detriment of our efforts in solving crystal structures of the full-length (Sa)EctC protein in complex with either N-γ-ADABA or ectoine. Because microbial ectoine producers can colonize ecological niches with rather different physicochemical attributes, it seems promising to exploit this considerable biodiversity to identify EctC proteins with enhanced protein stability. It is hoped that these can be further employed to obtain EctC crystal structures with either the substrate or the reaction product. Together with our finding that ectoine synthase is metal dependent, these crystal structures should allow a more detailed understanding of the chemistry underlying the EctC-catalyzed cyclocondensation reaction. Supporting Information References Compatible and counteracting solutes: protecting cells from the Dead Sea to the deep sea Uptake and synthesis of compatible solutes as microbial stress responses to high osmolality environments Physiological and genetic responses of bacteria to osmotic stress Intracellular organic osmolytes: function and regulation Osmoadaptation in bacteria and archaea: common principles and differences A molecular mechanism for osmolyte-induced protein stability Responses of E. coli to osmotic stress: large changes in amounts of cytoplasmic solutes and water Bacterial osmoregulation: a paradigm for the study of cellular homeostasis Roles of cytoplasmic osmolytes, water, and crowding in the response of Escherichia coli to osmotic stress: biophysical basis of osmoprotection by glycine betaine 1,4,5,6-Tetrahydro-2-methyl-4-pyrimidinecarboxylic acid. A novel cyclic amino acid from halophilic phototrophic bacteria of the genus Ectothiorhodospira The structure and biosynthesis of new tetrahydropyrimidine derivatives in actinomycin D producer Streptomyces parvulus. Use of 13C- and 15N-labeled L-glutamate and 13C and 15N NMR spectroscopy Ectoines in cell stress protection: uses and biotechnological production Biochemical properties of ectoine hydroxylases from extremophiles and their wider taxonomic distribution among microorganisms Strangers in the archaeal world: osmostress-responsive biosynthesis of ectoine and hydroxyectoine by the marine thaumarchaeon Nitrosopumilus maritimus Osmotically regulated synthesis of the compatible solute ectoine in Bacillus pasteurii and related Bacillus spp Synthesis of the compatible solute ectoine in Virgibacillus pantothenticus is triggered by high salinity and low growth temperature Unravelling the adaptation responses to osmotic and temperature stress in Chromohalobacter salexigens, a bacterium with broad salinity tolerance Cohesion group approach for evolutionary analysis of aspartokinase, an enzyme that feeds a branched network of many biochemical pathways A specialized aspartokinase enhances the biosynthesis of the osmoprotectants ectoine and hydroxyectoine in Pseudomonas stutzeri A1501 Characterization of biosynthetic enzymes for ectoine as a compatible solute in a moderately halophilic eubacterium, Halomonas elongata  Glass-forming property of hydroxyectoine is the cause of its superior function as a dessication protectant Synthesis and uptake of the compatible solutes ectoine and 5-hydroxyectoine by Streptomyces coelicolor A3(2) in response to salt and heat stresses The ectD gene, which is involved in the synthesis of the compatible solute hydroxyectoine, is essential for thermoprotection of the halophilic bacterium Chromohalobacter salexigens Osmotically induced synthesis of the compatible solute hydroxyectoine is mediated by an evolutionarily conserved ectoine hydroxylase Molecular dynamics simulations and structure-guided mutagenesis provide insight into the architecture of the catalytic core of the ectoine hydroxylase Crystal structure of the ectoine hydroxylase, a snapshot of the active site Enzyme stabilization by ectoine-type compatible solutes: protection against heating, freezing and drying Compatible solute influence on nucleic acids: many questions but few answers Extrinsic protein stabilization by the naturally occurring osmolytes beta-hydroxyectoine and betaine The effect of compatible solute ectoines on the structural organization of lipid monolayer and bilayer membranes The multifunctional role of ectoine as a natural cell protectant Industrial production of the cell protectant ectoine: protection, mechanisms, processes, and products Characterization of genes for the biosynthesis of the compatible solute ectoine from Marinococcus halophilus and osmoregulated expression in Escherichia coli Characterization of biosynthetic enzymes for ectoine as a compatible solute in a moderately halophilic eubacterium, Halomonas elongata Unexpected property of ectoine synthase and its application for synthesis of the engineered compatible solute ADPC The hydroxyectoine gene cluster of the non-halophilic acidophile Acidiphilium cryptum Characterization of the ectoine biosynthesis genes of haloalkalotolerant obligate methanotroph \"Methylomicrobium alcaliphilum 20Z\" A modified FMOC-method for the detection of amino acid-type osmolytes and tetrahydropyrimidines (ectoines) Cold adaptation in the marine bacterium, Sphingopyxis alaskensis, assessed using quantitative proteomics Overproduction, crystallization and X-ray diffraction data analysis of ectoine synthase from the cold-adapted marine bacterium Sphingopyxis alaskensis  Studies on the chemical nature of Clostridial ferredoxin XDS Integration, scaling, space-group assignment and post-refinement On the combination of molecular replacement and single-wavelength anomalous diffraction phasing for automated structure determination Towards automated crystallographic structure refinement with phenix.refine Features and development of Coot REFMAC5 for the refinement of macromolecular crystal structures  A blueprint of ectoine metabolism from the genome of the industrial producer Halomonas elongata DSM 2581T Evolution of functional diversity in the cupin superfamily Protein similarity networks reveal relationships among sequence, structure, and function within the Cupin superfamily Cupins: the most functionally diverse protein superfamily? Structure-based phylogeny as a diagnostic for functional characterization of proteins with a cupin fold The polyketide cyclase RemF from Streptomyces resistomycificus contains an unusual octahedral zinc binding site Biochemical and structural characterization of a novel bacterial manganese-dependent hydroxynitrile lyase The crystal structure of a quercetin 2,3-dioxygenase from Bacillus subtilis suggests modulation of enzyme activity by a change in the metal ion at the active site(s) Dali server: conservation mapping in 3D Crystal structure of a novel manganese-containing cupin (TM1459) from Thermotoga maritima at 1.65 A resolution Crystal structure of auxin-binding protein 1 in complex with auxin Crystal structure of bifunctional aldos-2-ulose dehydratase/isomerase from Phanerochaete chrysosporium with the reaction intermediate ascopyrone M Inference of macromolecular assemblies from crystalline state Real-time metabolome profiling of the metabolic switch between starvation and growth Protein-folding location can regulate manganese-binding versus copper- or zinc-binding Ring-cleaving dioxygenases with a cupin fold X-ray structure of a novel L-ribose isomerase acting on a non-natural sugar L-ribose as its ideal substrate The universality of enzymatic rate-temperature dependency Some like it cold: biocatalysis at low temperatures Accumulation of compatible solutes, ectoine and hydroxyectoine, in a moderate halophile, Halomonas elongata KS3, isolated from dry salty land in 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diff --git a/annotated_BioC_JSON/PMC4802042_ann.json b/annotated_BioC_JSON/PMC4802042_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..ec8e7fac77adb41a4042495819ee8dfca8ca45e3
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4802042_ann.json
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+[{"sourceid":"4802042","sourcedb":"","project":"","target":"","text":"A conserved motif in JNK/p38-specific MAPK phosphatases as a determinant for JNK1 recognition and inactivation Mitogen-activated protein kinases (MAPKs), important in a large array of signalling pathways, are tightly controlled by a cascade of protein kinases and by MAPK phosphatases (MKPs). MAPK signalling efficiency and specificity is modulated by protein–protein interactions between individual MAPKs and the docking motifs in cognate binding partners. Two types of docking interactions have been identified: D-motif-mediated interaction and FXF-docking interaction. Here we report the crystal structure of JNK1 bound to the catalytic domain of MKP7 at 2.4-Å resolution, providing high-resolution structural insight into the FXF-docking interaction. The 285FNFL288 segment in MKP7 directly binds to a hydrophobic site on JNK1 that is near the MAPK insertion and helix αG. Biochemical studies further reveal that this highly conserved structural motif is present in all members of the MKP family, and the interaction mode is universal and critical for the MKP-MAPK recognition and biological function.  The important MAPK family of signalling proteins is controlled by MAPK phosphatases (MKPs). Here, the authors report the structure of MKP7 bound to JNK1 and characterise the conserved MKP-MAPK interaction. The mitogen-activated protein kinases (MAPKs) are central components of the signal-transduction pathways, which mediate the cellular response to a variety of extracellular stimuli, ranging from growth factors to environmental stresses. The MAPK signalling pathways are evolutionally highly conserved. The basic assembly of MAPK pathways is a three-tier kinase module that establishes a sequential activation cascade: a MAPK kinase kinase activates a MAPK kinase, which in turn activates a MAPK. The three best-characterized MAPK signalling pathways are mediated by the kinases extracellular signal-regulated kinase (ERK), c-Jun N-terminal kinase (JNK) and p38. The ERK pathway is activated by various mitogens and phorbol esters, whereas the JNK and p38 pathways are stimulated mainly by environmental stress and inflammatory cytokines. The MAPKs are activated by MAPK kinases that phosphorylate the MAPKs at conserved threonine and tyrosine residues within their activation loop. After activation, each MAPK phosphorylates a distinct set of protein substrates, which act as the critical effectors that enable cells to mount the appropriate responses to varied stimuli. MAPKs lie at the bottom of conserved three-component phosphorylation cascades and utilize docking interactions to link module components and bind substrates. Two types of docking motifs have been identified in MAPK substrates and cognate proteins: kinase-interacting motif (D-motif) and FXF-motif (also called DEF motif, docking site for ERK FXF). The best-studied docking interactions are those between MAP kinases and ‘D-motifs', which consists of two or more basic residues followed by a short linker and a cluster of hydrophobic residues. The D-motif-docking site (D-site) in MAPKs is situated in a noncatalytic region opposite of the kinase catalytic pocket and is comprised of a highly acidic patch and a hydrophobic groove. D-motifs are found in many MAPK-interacting proteins, including substrates, activating kinases and inactivating phosphatases, as well as scaffolding proteins. A second docking motif for MAPKs consists of two Phe residues separated by one residue (FXF-motif). This motif has been observed in several MAPK substrates. The FXF-motif-binding site of ERK2 has been mapped to a hydrophobic pocket formed between the P+1 site, αG helix and the MAPK insert. However, the generality and mechanism of the FXF-mediated interaction is unclear. The physiological outcome of MAPK signalling depends on both the magnitude and the duration of kinase activation. Downregulation of MAPK activity can be achieved through direct dephosphorylation of the phospho-threonine and/or tyrosine residues by various serine/threonine phosphatases, tyrosine phosphatases and dual-specificity phosphatases (DUSPs) termed MKPs. MKPs constitute a group of DUSPs that are characterized by their ability to dephosphorylate both phosphotyrosine and phosphoserine/phospho-threonine residues within a substrate. Dysregulated expression of MKPs has been associated with pathogenesis of various diseases, and understanding their precise recognition mechanism presents an important challenge and opportunity for drug development. Here, we present the crystal structure of JNK1 in complex with the catalytic domain of MKP7. This structure reveals the molecular mechanism underlying the docking interaction between MKP7 and JNK1. In the JNK1–MKP7 complex, a hydrophobic motif (285FNFL288) that initiates the helix α5 in the MKP7 catalytic domain directly binds to the FXF-motif-binding site on JNK1, providing the structural insight into the classic FXF-type docking interaction. Biochemical and modelling studies further demonstrate that the molecular interactions mediate this key element for substrate recognition are highly conserved among all MKP-family members. Thus, our study reveals a hitherto unrecognized interaction mode for encoding complex target specificity among MAPK isoforms. Results Interaction of JNK1 with the MKP7 catalytic domain DUSPs belong to the protein-tyrosine phosphatases (PTPase) superfamily, which is defined by the PTPase-signature motif CXXGXXR. MKPs represent a distinct subfamily within a larger group of DUSPs. In mammalian cells, the MKP subfamily includes 10 distinct catalytically active MKPs. All MKPs contain a highly conserved C-terminal catalytic domain (CD) and an N-terminal kinase-binding domain (KBD). The KBD is homologous to the rhodanese family and contains an intervening cluster of basic amino acids, which has been suggested to be important for interacting with the target MAPKs. On the basis of sequence similarity, substrate specificity and predominant subcellular localization, the MKP family can be further divided into three groups (Fig. 1). Biochemical and structural studies have revealed that the KBD of MKPs is critical for MKP3 docking to ERK2, and MKP5 binding to p38α, although their binding mechanisms are completely different. However, it is unknown if other MAPKs can interact with the KBD of their cognate phosphatases in the same manner as observed for recognition of ERK2 and p38α by their MKPs, or whether they recognize distinct docking motifs of MKPs. MKP7, the biggest molecule in the MKP family, selectively inactivates JNK and p38 following stress activation. In addition to the CD and KBD, MKP7 has a long C-terminal region that contains both nuclear localization and export sequences by which MKP7 shuttles between the nucleus and the cytoplasm (Fig. 2a). To quantitatively assess the contribution of the N-terminal domain to the MKP7-catalysed JNK1 dephosphorylation, we first measured the kinetic parameters of the C-terminal truncation of MKP7 (MKP7ΔC304, residues 5–303) and MKP7-CD (residues 156–301) towards phosphorylated JNK1 (pJNK1). Figure 2b shows the variation of initial rates of the MKP7ΔC304 and MKP7-CD-catalysed reaction with the concentration of phospho-JNK1. Because the concentrations of MKP7 and pJNK1 were comparable in the reaction, the assumption that the free-substrate concentration is equal to the total substrate concentration is not valid. Thus, the kinetic data were analysed using the general initial velocity equation, taking substrate depletion into account: The kcat and Km of the MKP7-CD (0.028 s−1 and 0.26 μM) so determined were nearly identical to those of MKP7ΔC304 (0.029 s−1 and 0.27 μM), indicating that the MKP7-KBD has no effect on enzyme catalysis. We next examined the interaction of JNK1 with the CD and KBD of MKP7 by gel filtration analysis. When 3 molar equivalents of CD were mixed with 1 molar equivalent of JNK1, a significant amount of CD co-migrated with JNK1 to earlier fractions, and the excess amount of CD was eluted from the size exclusion column as a monomer, indicating stable complex formation (Fig. 2c). In contrast, no KBD–JNK1 complex was detected when 3 molar equivalents of KBD were mixed with 1 molar equivalent of JNK1. To further confirm the JNK1–MKP7-CD interaction, we performed a pull-down assay using the purified proteins. As shown in Fig. 2d, the CD of MKP7 can be pulled down by JNK1, while the KBD failed to bind to the counterpart protein. Taken together, our data indicate that the CD of MKP7, but not the KBD domain, is responsible for JNK substrate-binding and enzymatic specificity. Crystal structure of JNK1 in complex with the MKP7-CD To understand the molecular basis of JNK1 recognition by MKP7, we determined the crystal structure of unphosphorylated JNK1 in complex with the MKP7-CD (Fig. 3a, Supplementary Fig. 1a and Table 1). In the complex, JNK1 has its characteristic bilobal structure comprising an N-terminal lobe rich in β-sheet and a C-terminal lobe that is mostly α-helical. The overall folding of MKP7-CD is typical of DUSPs, with a central twisted five-stranded β-sheet surrounded by six α-helices. One side of the β-sheet is covered with two α-helices and the other is covered with four α-helices (Fig. 3b). The catalytic domain of MKP7 interacts with JNK1 through a contiguous surface area that is remote from the active site. MKP7-CD is positioned onto the JNK1 molecule so that the active site of the phosphatase faces towards the activation segment. In an alignment of the structure of MKP7-CD with that of VHR, an atypical ‘MKP' consisting of only a catalytic domain, 119 of 147 MKP7-CD residues could be superimposed with a r.m.s.d. (root mean squared deviation) of 1.05 Å (Fig. 3c). The most striking difference is that helix α0 and loop α0–β1 of VHR are absent in MKP7-CD. Another region that cannot be aligned with VHR is found in loop β3–β4. This loop is shortened by nine residues in MKP7-CD compared with that in VHR. Since helix α0 and the following loop α0–β1 are known for a substrate-recognition motif of VHR and other phosphatases, the absence of these moieties implicates a different substrate-binding mode of MKP7. The active site of MKP7 consists of the phosphate-binding loop (P-loop, Cys244-Leu245-Ala246-Gly247-Ile248-Ser249-Arg250), and Asp213 in the general acid loop (Fig. 3b and Supplementary Fig. 1b). The MKP7-CD structure near the active site exhibits a typical active conformation as found in VHR and other PTPs. The catalytic residue, Cys244, is located just after strand β5 and optimally positioned for nucleophilic attack. Asp213 in MKP7 also adopts a position similar to that of Asp92 in VHR (Supplementary Fig. 1c), indicating that Asp213 is likely to function as the general acid in MKP7. We also observed the binding of a chloride ion in the active site of MKP7-CD. It is located 3.36 Å from the Cys244 side chain and makes electrostatic interactions with the dipole moment of helix α3 and with several main-chain amide groups. The side chain of strictly conserved Arg250 is oriented towards the negatively charged chloride, similar to the canonical phosphate-coordinating conformation. Thus this chloride ion is a mimic for the phosphate group of the substrate, as revealed by a comparison with the structure of PTP1B in complex with phosphotyrosine (Supplementary Fig. 1d). Although the catalytically important residues in MKP7-CD are well aligned with those in VHR, the residues in the P-loop of MKP7 are smaller and have a more hydrophobic character than those of VHR (Cys124-Arg125-Glu126-Gly127-Tyr128-Gly129-Arg130; Fig. 3b,c). The difference in the polarity/hydrophobicity of the surface may also point to the origin of the differences in the substrate-recognition mechanism for these two phosphatases (Supplementary Fig. 1e,f). In the complex, MKP7-CD and JNK1 form extensive protein–protein interactions involving the C-terminal helices of MKP7-CD and C-lobe of JNK1 (Fig. 3d,e). As a result, the buried solvent-accessible surface area is ∼1,315 Å. In the C-terminal domain, JNK1 has an insertion after the helix αG. This insertion consists of two helices (α1L14 and α2L14) that are common to all members of the MAPK family. The interactive surface in JNK1, formed by the helices αG and α2L14, displays a hydrophobic region, centred at Trp234 (Fig. 3d). The MKP7-docking region includes two helices, α4 and α5, and the general acid loop. The aromatic ring of Phe285 on MKP7 α5-helix is nestled in a hydrophobic pocket on JNK1, formed by side chains of Ile197, Leu198, Ile231, Trp234, Val256, Tyr259, Val260 and the aliphatic portion of His230 (Fig. 3d,f and Supplementary Fig. 1g). In addition, there are hydrogen bonds between Ser282 and Asn286 of MKP7 and His230 and Thr255 of JNK1, and the main chain of Phe215 in the general acid loop of MKP7 is hydrogen-bonded to the side chain of Gln253 in JNK1. The second interactive area involves the α4 helix of MKP7 and charged/polar residues of JNK1 (Fig. 3e). The carboxylate of Asp268 in MKP7 forms a salt bridge with side chain of Arg263 in JNK1, and Lys275 of MKP7 forms a hydrogen bond and a salt bridge with Thr228 and Asp229 of JNK1, respectively. Mutational analysis of the JNK1–MKP7 docking interface To assess the importance of the aforementioned interactions, we generated a series of point mutations on the MKP7-CD and examined their effect on the MKP7-catalysed JNK1 dephosphorylation (Fig. 4a). When the hydrophobic residues Phe285 and Phe287 on the α5 of MKP7-CD were replaced by Asp or Ala, their phosphatase activities for JNK1 dephosphorylation decreased ∼10-fold. In comparison, replacement of the other residues (Phe215, Asp268, Lys275, Ser282, Asn286 and Leu292) with an Ala or Asp individually led to a modest decrease in catalytic efficiencies, suggesting that this position may only affect some selectivity of MKP. Mutation of Leu288 markedly reduced its solubility when expressed in Escherichia coli, resulting in the insoluble aggregation of the mutant protein. Gel filtration analysis further confirmed the key role of Phe285 in the MKP7–JNK1 interaction: no F285D–JNK1 complex was detected when 3 molar equivalents of MKP7-CD (F285D) were mixed with 1 molar equivalent of JNK1 (Fig. 4b). Interestingly, mutation of Phe287 results in a considerable loss of activity against pJNK1 without altering the affinity of MKP7-CD for JNK1 (Supplementary Fig. 2a). We also generated a series of point mutations in the JNK1 and assessed the effect on JNK1 binding using the GST pull-down assay (Fig. 4c). Substitution at Asp229, Trp234, Thr255, Val256, Tyr259 and Val260 significantly reduced the binding affinity of MKP7-CD for JNK. To determine whether the deficiencies in their abilities to bind partner proteins or carry out catalytic function are owing to misfolding of the purified mutant proteins, we also examined the folding properties of the JNK1 and MKP7 mutants with circular dichroism. The spectra of these mutants are similar to the wild-type proteins, indicating that these mutants fold as well as the wild-type proteins (Fig. 4d,e). Taken together, these results are consistent with the present crystallographic model, which reveal the hydrophobic contacts between the MKP7 catalytic domain and JNK1 have a predominant role in the enzyme–substrate interaction, and hydrophobic residue Phe285 in the MKP7-CD is a key residue for its high-affinity binding to JNK1. It has previously been reported that several cytosolic and inducible nuclear MKPs undergo catalytic activation upon interaction with the MAPK substrates. This allosteric activation of MKP3 has been well-documented in vitro using pNPP, a small-molecule phosphotyrosine analogue of its normal substrate. We then assayed pNPPase activities of MKP7ΔC304 and MKP7-CD in the presence of JNK1. Incubation of MKP7 with JNK1 did not markedly stimulate the phosphatase activity, which is consistent with previous results that MKP7 solely possesses the intrinsic activity (Supplementary Fig. 2b). The small pNPP molecule binds directly at the enzyme active site and can be used to probe the reaction mechanism of protein phosphatases. We therefore examined the effects of the MKP7-CD mutants on their pNPPase activities. As shown in Fig. 4f, all the mutants, except F287D/A, showed little or no activity change compared with the wild-type MKP7-CD. In the JNK1/MKP7-CD complex structure, Phe287 of MKP7 does not make contacts with JNK1 substrate. It penetrates into a pocket formed by residues from the P-loop and general acid loop and forms hydrophobic contacts with the aliphatic portions of side chains of Arg250, Glu217 and Ile219, suggesting that Phe287 in MKP7 would play a similar role to that of its structural counterpart in the PTPs (Gln266 in PTP1B) and VHR (Phe166 in VHR) in the precise alignment of active-site residues in MKP7 with respect to the substrate for efficient catalysis (Supplementary Fig. 2c). Kinase-associated phosphatase (KAP), a member of the DUSP family, plays a crucial role in cell cycle regulation by dephosphorylating the pThr160 residue of CDK2 (cyclin-dependent kinase 2). The crystal structure of the CDK2/KAP complex has been determined at 3.0 Å (Fig. 5a). The interface between these two proteins consists of three discontinuous contact regions. Biochemical results suggested that the affinity and specificity between KAP and CDK2 results from the recognition site comprising CDK2 residues from the αG helix and L14 loop and the N-terminal helical region of KAP (Fig. 5b). There is a hydrogen bond between the main-chain nitrogen of Ile183 (KAP) and side chain oxygen of Glu208 (CDK2), and salt bridges between Lys184 of KAP and Asp235 of CDK2. Structural analysis and sequence alignment reveal that one of the few differences between MKP7-CD and KAP in the substrate-binding region is the presence of the motif FNFL in MKP7-CD, which corresponds to IKQY in KAP (Fig. 5c). The substitution of the two hydrophobic residues with charged/polar residues (F285I/N286K) seriously disrupts the hydrophobic interaction required for MKP7 binding on JNK1 (Fig. 4a). In addition, His230 and Val256 in JNK1 are replaced by the negatively charged residues Glu208 and Asp235 in CDK2 (Fig. 5d), and the charge distribution on the CDK2 interactive surface is quite different from that of JNK. These data indicated that a unique hydrophobic pocket formed between the MAPK insert and αG helix plays a major role in the substrate recognition by MKPs. F-site interaction is crucial for JNK1 inactivation in vivo JNK is activated following cellular exposure to a number of acute stimuli such as anisomycin, H2O2, ultraviolet light, sorbitol, DNA-damaging agents and several strong apoptosis inducers (etoposide, cisplatin and taxol). To assess the effects of MKP7 and its mutants on the activation of endogenous JNK in vivo, HEK293T cells were transfected with blank vector or with HA-tagged constructs for full-length MKP7, MKP7ΔC304 and MKP7-CD or MKP7 mutants, and stimulated with ultraviolet or etoposide treatment. As shown in Fig. 6a–c, immunobloting showed similar expression levels for the different MKP7 constructs in all the cells. Overexpressed full-length MKP7, MKP7ΔC304 and MKP7-CD significantly reduced the endogenous level of phosphorylated JNK compared with vector-transfected cells. Parallel experiments showed clearly that the D-motif mutants (R56A/R57A and V63A/I65A) dephosphorylated JNK as did the wild type under the same conditions, further confirming that the MKP7-KBD is not required for the JNK inactivation in vivo. Consistent with the in vitro data, the level of phosphorylated JNK was not or little altered in MKP7 FXF-motif mutants (F285D, F287D and L288D)-transfected cells, and the MKP7 D268A and N286A mutants retained the ability to reduce the phosphorylation levels of JNK. We next tested in vivo interactions between JNK1 mutants and full-length MKP7 by coimmunoprecipitation experiments under unstimulated conditions. When co-expressed in HEK293T cells, wild-type (HA)-JNK1 was readily precipitated with (Myc)-MKP7 (Fig. 6d), indicating that MKP7 binds dephosphorylated JNK1 protein in vivo. In agreement with the in vitro pull-down results, the mutants D229A, W234D and Y259D were not co-precipitated with MKP7, and the I231D mutant had only little effect on the JNK1–MKP7 interaction (Fig. 6d and Supplementary Fig. 3a). Activation of the JNK signalling pathway is frequently associated with apoptotic cell death, and inhibition of JNK can prevent apoptotic death of multiple cells. To examine whether the inhibition of JNK activity by MKP7 would provide protections against the apoptosis, we analysed the rate of apoptosis in ultraviolet-irradiated cells transfected with MKP7 (wild type or mutants) by flow cytometry. The results showed similar apoptotic rates between cells transfected with blank vector or with MKP7 (wild type or mutants) under unstimulated conditions (Supplementary Fig. 3b), while ultraviolet-irradiation significantly increased apoptotic rate in cells transfected with blank vector (Fig. 6e). Expressions of wild-type MKP7, MKP7ΔC304 and MKP7-CD significantly decreased the proportion of apoptotic cells after ultraviolet treatment. Moreover, treatment of cells expressing MKP7-KBD mutants (R56A/R57A and V63A/I65A) decreased the apoptosis rates to a similar extent as MKP7 wild type did. In contrast, cells transfected with the MKP7 FXF-motif mutants (F285D, F287D and L288D) showed little protective effect after ultraviolet treatment and similar levels of apoptosis rates were detected to cells transfected with control vectors (Fig. 6e,f). Taken together, our results suggested that FXF-motif-mediated, rather than KBD-mediated, interaction is essential for MKP7 to block ultraviolet-induced apoptosis. A similar docking mechanism for JNK1 recognition by MKP5 MKP5 belongs to the same subfamily as MKP7. MKP5 is unique among the MKPs in possessing an additional domain of unknown function at the N-terminus (Fig. 7a). The KBD of MKP5 interacts with the D-site of p38α to mediate the enzyme–substrate interaction. Deletion of the KBD in MKP5 leads to a 280-fold increase in Km for p38α substrate. In contrast to p38α substrate, deletion of the MKP5-KBD had little effects on the kinetic parameters for the JNK1 dephosphorylation, indicating that the KBD of MKP5 is not required for the JNK1 dephosphorylation (Fig. 7b). The substrate specificity constant kcat /Km value for MKP5-CD was calculated as 1.0 × 105 M−1 s−1, which is very close to that of MKP7-CD (1.07 × 105 M−1 s−1). The crystal structure of human MKP5-CD has been determined. Comparisons between catalytic domains structures of MKP5 and MKP7 reveal that the overall folds of the two proteins are highly similar, with only a few regions exhibiting small deviations (r.m.s.d. of 0.79 Å; Fig. 7c). Given the distinct interaction mode revealed by the crystal structure of JNK1–MKP7-CD, one obvious question is whether this is a general mechanism used by all members of the JNK-specific MKPs. To address this issue, we first examined the docking ability of JNK1 to the KBD and CD of MKP5 using gel filtration analysis and pull-down assays. It can be seen from gel filtration experiments that JNK1 can forms a stable heterodimer with MKP5-CD in solution, but no detectable interaction was found with the KBD domain (Fig. 7d). Pull-down assays also confirmed the protein–protein interactions observed above. The catalytic domain of MKP5, but not its KBD, was able to pull-down a detectable amount of JNK1 (Fig. 7e), implicating a different substrate-recognition mechanisms for p38 and JNK MAPKs. To further test our hypothesis, we generated forms of MKP5-CD bearing mutations corresponding to the changes we made on MKP7-CD on the basis of sequence and structural alignment and examined their effects on the phosphatase activity. As shown in Fig. 7f, the T432A and L449F MKP5 mutant showed little or no difference in phosphatase activity, whereas the other mutants showed reduced specific activities of MKP5. As in the case of MKP7, all the mutants, except F451D/A, showed no pNPPase activity changes compared with the wild-type MKP5-CD (Fig. 7g), and the point mutations in JNK1 also reduced the binding affinity of MKP5-CD for JNK1 (Fig. 7h). In addition, there were no significant differences in the CD spectra between wild-type and mutant proteins, indicating that the overall structures of these mutants did not change significantly from that of wild-type MKP5 protein (Supplementary Fig. 4a). Taken together, our results suggest that MKP5 binds JNK1 in a docking mode similar to that in the JNK1–MKP7 complex, and the detailed interaction model can be generated using molecular dynamics simulation based on the structure of JNK1–MKP7-CD complex (Supplementary Fig. 4b,c). In this model, the MKP5-CD adopts a conformation nearly identical to that in its unbound form, suggesting that the conformation of the catalytic domain undergoes little change, if any at all, upon JNK1 binding. In particular, Leu449 of MKP5, which is equivalent to the key residue Phe285 of MKP7, buried deeply within the hydrophobic pocket of JNK1 in the same way as Phe285 in the JNK1–MKP7-CD complex (Supplementary Fig. 4d). Despite the strong similarities between JNK1–MKP5-CD and JNK1–MKP7-CD, however, there are differences. The JNK1–MKP7-CD interaction is better and more extensive. Asp268 of MKP7-CD forms salt bridge with JNK1 Arg263, whereas the corresponding residue Thr432 in MKP5-CD may not interact with JNK1. In addition, the key interacting residues of MKP7-CD, Phe215, Leu267 and Leu288, are replaced by less hydrophobic residues, Asn379, Met431 and Met452 in MKP5-CD (Fig. 5c), respectively, which may result in weaker hydrophobic interactions between MKP5-CD and JNK1. This is consistent with the experimental observation showing that JNK1 binds to MKP7-CD much more tightly than MKP5-CD (Km value of MKP5-CD for pJNK1 substrate is ∼20-fold higher than that of MKP7-CD). Discussion The MAPKs p38, ERK and JNK, are central to evolutionarily conserved signalling pathways that are present in all eukaryotic cells. Each MAPK cascade is activated in response to a diverse array of extracellular signals and culminates in the dual-phosphorylation of a threonine and a tyrosine residue in the MAPK-activation loop. The propagation of MAPK signals is attenuated through the actions of the MKPs. Most studies have focused on the dephosphorylation of MAPKs by phosphatases containing the ‘kinase-interaction motif ' (D-motif), including a group of DUSPs (MKPs) and a distinct subfamily of tyrosine phosphatases (HePTP, STEP and PTP-SL). Crystal structures of ERK2 bound with the D-motif sequences derived from MKP3 and HePTP have been reported. These structures revealed that linear docking motifs in interacting proteins bind to a common docking site on MAPKs outside the kinase active site. The particular amino acids and their spacing within D-motif sequences and amino acid composition of the docking sites on MAPKs appear to determine the specificity of D-motifs for individual MAPKs. Recently, the crystal structure of a complex between the KBD of MKP5 and p38α has been obtained. This complex has revealed a distinct interaction mode for MKP5. The KBD of MKP5 binds to p38α in the opposite polypeptide direction compared with how the D-motif of MKP3 binds to ERK2. In contrast to the canonical D-motif-binding mode, separate helices, α2 and α3′, in the KBD of MKP5 engage the p38α-docking site. Further structural and biochemical studies indicate that KBD of MKP7 may interact with p38α in a similar manner to that of MKP5. In contrast to MKP5, removal of the KBD domain from MKP7 does not drastically affect enzyme catalysis, and the kinetic parameters of MKP7-CD for p38α substrate are very similar to those for JNK1 substrate. Taken together, these results suggest that MKP7 utilizes a bipartite recognition mechanism to achieve the efficiency and fidelity of p38α signalling. The MKP7-KBD docks to the D-site located on the back side of the p38α catalytic pocket for high-affinity association, whereas the interaction of the MKP7-CD with another p38α structural region, which is close to the activation loop, may not only stabilize binding but also provide contacts crucial for organizing the MKP7 active site with respect to the phosphoreceptor in the p38α substrate for efficient dephosphorylation. In addition to the canonical D-site, the MAPK ERK2 contains a second binding site utilized by transcription factor substrates and phosphatases, the FXF-motif-binding site (also called F-site), that is exposed in active ERK2 and the D-motif peptide-induced conformation of MAPKs. This hydrophobic site was first identified by changes in deuterium exchange profiles, and is near the MAPK insertion and helix αG. Interestingly, many of the equivalent residues in JNK1, important for MKP7-CD recognition, are also used for substrate binding by ERK2 (ref.), indicating that this site is overlapped with the DEF-site previously identified in ERK2 (Fig. 5d). MKP3 is highly specific in dephosphorylating and inactivating ERK2, and the phosphatase activity of the MKP3-catalysed pNPP reaction can be markedly increased in the presence of ERK2 (refs). Sequence alignment of all MKPs reveals a high degree of conservation of residues surrounding the interacting region observed in JNK1–MKP7-CD complex (Supplementary Fig. 5). Therefore, it is tempting to speculate that the catalytic domain of MKP3 may bind to ERK2 in a manner analogous to the way by which MKP7-CD binds to JNK1. A comprehensive examination of the molecular basis of the specific ERK2 recognition by MKP3 is underway. The ongoing work demonstrates that although the overall interaction modes are similar between the JNK1–MKP7-CD and ERK2–MKP3-CD complexes, the ERK2–MKP3-CD interaction is less extensive and helix α4 from MKP3-CD does not interact directly with ERK2. The FXF-motif-mediated interaction is critical for both pERK2 inactivation and ERK2-induced MKP3 activation (manuscript in preparation). In summary, we have resolved the structure of JNK1 in complex with the catalytic domain of MKP7. This structure reveals an FXF-docking interaction mode between MAPK and MKP. Results from biochemical characterization of the Phe285 and Phe287 MKP7 mutants combined with structural information support the conclusion that the two Phe residues serve different roles in the catalytic reaction. Phe285 is essential for JNK1 substrate binding, whereas Phe287 plays a role for the precise alignment of active-site residues, which are important for transition-state stabilization. This newly identified FXF-type motif is present in all MKPs, except that the residue at the first position in MKP5 is an equivalent hydrophobic leucine residue (see also Fig. 7f,g), suggesting that these two Phe residues would play a similar role in facilitating substrate recognition and catalysis, respectively. An important feature of MKP–JNK1 interactions is that MKP7 or MKP5 only interact with the F-site of JNK1. One possible explanation is that JNK1 needs to use the D-site to interact with JIP-1, a scaffold protein for JNK signalling. The N-terminal JNK-binding domain of JIP-1 interacts with the D-site on JNK and this interaction is required for JIP-1-mediated enhancement of JNK activation. In addition, JIP-1 can also associate with MKP7 via the C-terminal region of MKP7 (ref.). When MKP7 is bound to JIP-1, it reduces JNK activation, leading to reduced phosphorylation of the JNK target c-Jun. Thus, our biochemical and structural data allow us to present a model for the JNK1–JIP-1–MKP7 ternary complex and provide an important insight into the assembly and function of JNK signalling modules (Supplementary Fig. 6). Methods Protein preparation The cDNAs of human MKP7 and MKP5 were kindly provided by Dr Mathijs Baens (University of Leuven) and Dr Eisuke Nishida (Kyoto University), respectively. The cDNAs of human ASK1, MKK4, MKK7 and JNK1 were kindly provided by Dr Zhenguo Wu (Hong Kong University of Science and Technology). The catalytic domains of MKP7 (MKP7-CD, residues 156–301) and MKP5 (MKP5-CD, 320–467) and the full-length MKP5 were cloned into the pET15b vector, resulting in the N-terminal His-fusion proteins. The KBD domains of MKP7 (MKP7-KBD, 5–138) and MKP5 (MKP5-KBD, 139–287), and the C-terminal truncation of MKP7 (MKP7ΔC304, 5–303) were cloned into pET21b vector for generation of C-terminal His-tagged proteins. The human full-length JNK1, MKK4, MKK7 and the kinase domain of ASK1 (659–951) were cloned into pGEX4T-2, pET15b and/or pET21b vectors to produce a GST- or His-tagged protein. Mutations of MKP7-CD, MKP5-CD and JNK1 were generated by overlap PCR procedure. All constructs were verified by DNA sequencing. All proteins, overexpressed in BL21(DE3) cells at 20 °C, were first purified over Ni-NTA (Qiagen) or GS4B (GE Healthcare) columns, and then by ion exchange and gel filtration chromatography (Source-15Q/15S and Superdex-200, GE Healthcare) at 4 °C. The double-phosphorylated JNK1 (phospho-JNK1) was generated by mixing JNK1 with upstream kinases MKK4, MKK7 and ASK1 in buffer containing 10 mM MgCl2 and 2 mM ATP, and further purified by gel filtration chromatography (Superdex-200, GE Healthcare) at 4 °C (ref.). Proteins were stored at −80 °C, and stocks for phosphatase assays were supplemented with glycerol to a final concentration of 20% (v/v). Protein concentrations were determined spectrophotometrically using theoretical molar extinction coefficients at 280 nm (ref.). Crystallography The mixture of unphosphorylated JNK1 and MKP7-CD at 1:1 molar ratio was subjected to crystallization trials. Crystals were grown by the vapor-diffusion technique in hanging drops, and the drops were prepared by mixing equal volumes of protein with reservoir solution containing 0.1 M HEPES, pH 7.0, 14% PEG3350, 0.2 M MgCl2, 6% 1,6-Hexanediol and 0.005 M EDTA at 21 °C. Crystals were cryo-protected in reservoir solutions supplemented with 10% glycerol and then flash frozen in liquid nitrogen. The diffraction data sets were collected at beamline 17U at Shanghai Synchrotron Radiation Facility and processed with the HKL2000 package. The crystals belong to space group P1 and comprise eight molecules per asymmetric unit (four complexes). Structure was solved by molecular replacement using Phaser with JNK1 (PDB 1UKH) and MKP5-CD (PDB 1ZZW) as the search models. Standard refinement was performed with programs PHENIX and Coot. The crystal structure of unphosphorylated JNK1 in complex with the catalytic domain of MKP7 was refined to 2.4 Å resolution. Initial structural refinement was performed with NCS restraints, and after several rounds the restraints were removed from the calculations. The final Rwork and Rfree were 21.7 and 23.9%, respectively. The crystallographic asymmetric unit contains four JNK1–MKP7-CD complexes. The four complexes are nearly identical with an r.m.s.d.\u003c1 Å for any complex pair in the asymmetric unit. Ramachandran analysis was carried out using PROCHECK. Additional density at the active site of MKP7-CD was attributed to a chloride ion incorporated as a crystallizing agent, similar to those observed in the structures of MKP3-CD and MKP5-CD (refs). The data collection and refinement statistics are summarized in Table 1. All structural representations in this paper were prepared with PYMOL (http://www.pymol.org). Phosphatase assays The activities of MKP7 and MKP5 was assayed using phospho-JNK1 as substrate in the coupled enzyme system containing 50 mM MOPS, pH 7.0, 100 mM NaCl, 0.1 mM EDTA, 50 μM MESG and 0.1 mg ml−1 PNPase. This coupled assay uses PNPase and its chromogenic substrate MESG to monitor the production of inorganic phosphate. The reactions were initiated by addition of 0.4 μM MKP7-CD and MKP7ΔC304 (or 0.1 μM MKP5 full-length and 0.15 μM MKP5-CD) for substrate phospho-JNK1. All experiments were carried out at 25 °C in 1.8 ml reaction mixtures, and the continuous absorbance changes were recorded with a PerkinElmer LAMBDA 45 spectrophotometer equipped with a magnetic stirrer in the cuvette holder. The quantification of inorganic phosphate produced was monitored at 360 nm with the extinction coefficient of 11,200 M−1 cm−1 (ref.). The initial rates were determined from the linear slope of the progress curves obtained. The activity of MKP7-CD or MKP5-CD mutants were assayed using pNPP or phospho-JNK1 as substrate. The phospho-JNK1 assay was performed as the same procedure mentioned above, and in the presence of wild type (as a control) or indicated mutants, and equal concentrations of phospho-JNK1. The pNPP assay was performed in the reaction mixture containg 50 mM MOPS, pH 7.0, 100 mM NaCl, 0.1 mM EDTA and 20 mM pNPP. The amount of the product p-nitrophenol was determined from the absorbance at 405 nm using a molar extinction coefficient of 18,000 M−1 cm−1 (ref.). Assays for protein–protein interaction The interactions of JNK1 with the CD and KBD domains of MKP7 and MKP5 were examined by gel filtration analyses using a Superdex-200 10/300 column on an ÄKTA FPLC (GE Healthcare). The column was equilibrated with a buffer containing 10 mM HEPES, pH 7.5, 150 mM NaCl and 2 mM dithiothreitol, and calibrated with molecular mass standards. Samples of individual proteins and indicated mixtures (500 μl each) were loaded to the Superdex-200 column and then eluted at a flow rate of 0.5 ml min−1. Fractions of 0.5 ml each were collected, and aliquots of relevant fractions were subjected to SDS-polyacrylamide gel electrophoresis (PAGE) followed Coomassie Blue staining. The interactions between various JNK1 mutants and MKP7-CD or MKP5-CD were assessed by GST-mediated pull-down assays at 4 °C. First, 0.5 ml GST-JNK1 proteins (6 μM) were loaded to 0.2 ml GS4B resin. The excess unbound JNK1 or other contaminants were removed by washing the column 5 times, each with 1.0 ml buffer containing 25 mM Tris-HCl, pH 8.0, 150 mM NaCl and 2 mM dithiothreitol. Then, 0.5 ml MKP7-CD or MKP5-CD (20 μM) was allowed to flow through the JNK1-bound column. After extensive washing, the bound proteins were eluted with 0.5 ml reduced glutathione (10 mM). The interactions of JNK1 with the CD and KBD domains of MKP7 and MKP5 were also examined by GST-mediated pull-down assays. The GST protein alone was used as a control. Aliquots of all eluates were subjected to SDS-PAGE, and proteins were visualized by Coomassie Blue staining. The uncropped gels are shown in Supplementary Fig. 7. Building JNK and MKP5 interaction model The model of catalytic domain of MKP5 bound to JNK was constructed by superimposition of previous deposited structures of MKP5-CD (PDB 1ZZW) to the corresponding domains in the crystal structure of JNK1–MKP7-CD. The fractured loops in deposited structures were computationally generated using Modeller. The program CHARMM22 (ref.) was then used to add hydrogen atoms, N- and C-terminal patches to the model. The model was then subjected to restrained energy minimization to optimize bonds and remove any nonbonded steric clashes. Refinement of the modelled complex was performed using NAMD2.9 package at 1 atm pressure and 300 K. The generated complex structure was solvated and neutralized in a box with TIP3P water at a minimum of 13 Å between the model and the wall of the box. The simulation was first set up with 1 fs time step under periodic boundary conditions. The particle mesh Ewald method was applied to model the electrostatics and the van der Waals interactions cutoff was set at 12 Å. The system was restrained for 5 ps minimization and 5 ps simulation, and followed by removing all the restraints and performing a minimization of 10 ps and an equilibration of 10 ns. Simulations were viewed using VMD. Circular dichroism spectra The experiments were performed on a Chirascan-plus circular dichroism Spectrometer (Applied Photophysics, Surrey, UK) using 0.1 mm quartz cuvette. The protein sample were analysed at a concentration of 0.5 mg ml−1. Data were collected over a wavelength range from 260 to 190 nm with 1 nm intervals at room temperature, three scans were averaged, and the baseline spetrum of solution buffer containing 10 mM HEPES (pH 7.5) and 150 mM NaCl was subtracted. Cell culture and transfections pcDNA3.3-Myc-MKP7, pCMV5-3HA-JNK1 and pBOBI-HA-MKP7 were generated with standard molecular techniques. Mutants with amino acid substitution and truncation constructs were generated through PCR-based site-directed mutagenesis method using Pfu polymerase (Stratagene). The authenticities of all constructs were confirmed by sequencing (Invitrogen, China). HEK293T and HeLa cells (ATCC) were maintained in DMEM supplemented with 10% fetal bovine serum, 100 IU penicillin, 100 mg ml−1 streptomycin at 37 °C in a humidified incubator containing 5% CO2. Polyethylenimine (Polysciences, #23966) at a final concentration of 10 μM was used to transfect HEK293T cells. Total DNA for each plate was adjusted to the same amount by adding relevant blank vector. Lentiviruses for infection were packaged in HEK293T cells after transfection using Lipofectamine 2000 (Invitrogen, 11668-027). At 30 h post transfection, medium was collected for further infection. Coimmunoprecipitation and immunoblotting Cells were lysed in a lysis buffer containing 20 mM Tris-HCl (pH 7.4), 150 mM NaCl, 0.5% NP-40, 1 mM EDTA, 2 mM Na3VO4, 25 mM NaF, 1 mM phenylmethanesulfonyl fluoride, 1 μg ml−1 leupeptin and 1 μg ml−1 aprotinin. Cell lysates were incubated with respective antibodies overnight at 4 °C. Protein aggregates resulting from the overnight incubation were removed by centrifugation, and protein A/G beads (Santa-Cruz Biotechnology, Dallas, TX, USA) were then added into the lysates and incubated for another 3 h. After spinning and washing for three times with the lysis buffer, the beads were mixed with 2 × SDS sample buffer, boiled and subjected to 15% SDS/PAGE. The samples were transferred to PVDF membranes (Millipore), and immunoblotted with indicated antibodies. Levels of total proteins and the levels of phosphorylation of proteins were analysed on separate gels. The uncropped blots are shown in Supplementary Fig. 7. Antibodies and drugs Antibodies used in this study: mouse anti-HA (1:100 for immunoprecipitation; F-7) and anti-JNK1 (1:1,000 for immunoblotting; F-3) antibodies were purchased from Santa-Cruz Biotechnology. Anti-c-Myc Agarose Affinity Gel antibody produced in rabbit (1:200 for immunoprecipitation; A7470) was purchased from Sigma. Rabbit anti-HA-tag (1:1,000 for immunoblotting; #3724), anti-phospho-JNK-T183/Y185 (1:1,000 for immunoblotting; #4668) and mouse anti-Myc-tag (1:1,000 for immunoblotting; #2276) antibodies were purchased from Cell Signaling Technology. Etoposide (E1383) was purchased from Sigma. Apoptosis assay HeLa cells were infected with lentiviruses expressing MKP7 or its mutants. At 36 h post infection, cells were irradiated with 25 J m−2 ultraviolet light and collected at 6 h after irradiation. Cells were then stained with the Annexin-V-APC/PI double-staining solution (BD Biosciences) and analysed with a flow cytometer (BD LSRFortessa). The percentages of apoptotic cells were quantified with FlowJo 7.6.1 software. Additional information Accession codes: The coordinates and structure factors have been deposited in the Protein Data Bank with accession codes 4YR8 for the JNK1–MKP7-CD structure. How to cite this article: Liu, X. et al. A conserved motif in JNK/p38-specific MAPK phosphatases as a determinant for JNK1 recognition and inactivation. Nat. Commun. 7:10879 doi: 10.1038/ncomms10879 (2016). Supplementary Material MAP kinase pathways: the first twenty years Activation and function of the MAPKs and their substrates, the MAPK-activated protein kinases Differential regulation and properties of MAPKs Coordinating ERK/MAPK signalling through scaffolds and inhibitors The JNK signal transduction pathway Signal integration by JNK and p38 MAPK pathways in cancer development Mechanisms of MAPK signalling specificity Unique MAP kinase binding sites Multiple docking sites on substrate proteins form a modular system that mediates recognition by ERK MAP kinase Docking sites on substrate proteins direct extracellular signal-regulated kinase to phosphorylate specific residues Selective targeting of MAPKs to the ETS domain transcription factor SAP-1 ERK2 mitogen-activated protein kinase binding, phosphorylation, and regulation of the PDE4D cAMP-specific phosphodiesterases. The involvement of COOH-terminal docking sites and NH2-terminal UCR regions Molecular interpretation of ERK signal duration by immediate early gene products Docking motif interactions in MAP kinases revealed by hydrogen exchange mass spectrometry Regulation of MAP kinases by MAP kinase phosphatases Differential regulation of MAP kinase signalling by dual-specificity protein phosphatases Dual-specificity phosphatases: critical regulators with diverse cellular targets Targeting dual-specificity phosphatases: manipulating MAP kinase signalling and immune responses Dual-specificity MAP kinase phosphatases (MKPs) and cancer Protein tyrosine phosphatases: from genes, to function, to disease Diverse physiological functions for dual-specificity MAP kinase phosphatases Structural basis of docking interactions between ERK2 and MAP kinase phosphatase 3 A distinct interaction mode revealed by the crystal structure of the kinase p38alpha with the MAPK binding domain of the phosphatase MKP5 A Novel MAPK phosphatase MKP-7 acts preferentially on JNK/SAPK and p38 alpha and beta MAPKs Crystal structure of the dual specificity protein phosphatase VHR Protein-tyrosine phosphatases: biological function, structural characteristics, and mechanism of catalysis Structural basis for the recognition of a bisphosphorylated MAP kinase peptide by human VHR protein Phosphatase Structure and regulation of MAPK phosphatases Identification of a second aryl phosphate-binding site in protein-tyrosine phosphatase 1B: a paradigm for inhibitor design Catalytic activation of the phosphatase MKP-3 by ERK2 mitogen-activated protein kinase Mechanism of mitogen-activated protein kinase phosphatase-3 activation by ERK2 Altering the nucleophile specificity of a protein-tyrosine phosphatase-catalyzed reaction. Probing the function of the invariant glutamine residues Large-scale structural analysis of the classical human protein tyrosine phosphatome Structure and catalytic mechanism of human protein tyrosine phosphatome Insights into the reaction of protein-tyrosine phosphatase 1B: crystal structures for transition state analogs of both catalytic steps Phosphoprotein-protein interactions revealed by the crystal structure of kinase-associated phosphatase in complex with phosphoCDK2 From JNK to pay dirt: jun kinases, their biochemistry, physiology and clinical importance c-Jun NH2-terminal kinase-mediated activation of interleukin-1beta converting enzyme/CED-3-like protease during anticancer drug-induced apoptosis CL100/MKP-1 modulates JNK activation and apoptosis in response to cisplatin JNK signaling in apoptosis Conditional expression of mitogen-activated protein kinase phosphatase-1, MKP-1, is cytoprotective against UV-induced apoptosis Conditional expression of the mitogen-activated protein kinase (MAPK) phosphatase MKP-1 preferentially inhibits p38 MAPK and stress-activated protein kinase in U937 cells Regulation of JNK activity in the apoptotic response of intestinal epithelial cells Molecular cloning and characterization of a novel dual specificity phosphatase, MKP-5 Crystal structure of the catalytic domain of human MAP kinase phosphatase 5: structural insight into constitutively active phosphatase Differential interaction of the tyrosine phosphatases PTP-SL, STEP and HePTP with the mitogen-activated protein kinases ERK1/2 and p38alpha is determined by a kinase specificity sequence and influenced by reducing agents PTP-SL and STEP protein tyrosine phosphatases regulate the activation of the extracellular signal-regulated kinases ERK1 and ERK2 by association through a kinase interaction motif Docking interactions induce exposure of activation loop in the MAP kinase ERK2 Activation mechanism of the MAP kinase ERK2 by dual phosphorylation A cytoplasmic inhibitor of the JNK signal transduction pathway Docking interactions in the c-Jun N-terminal kinase pathway Structural basis for the selective inhibition of JNK1 by the scaffolding protein JIP1 and SP600125 A mammalian scaffold complex that selectively mediates MAP kinase activation The JNK-interacting protein-1 scaffold protein targets MAPK phosphatase-7 to dephosphorylate JNK High yield purification of JNK1beta1 and activation by in vitro reconstitution of the MEKK1--\u003eMKK4--\u003eJNK MAPK phosphorylation cascade Calculation of protein extinction coefficients from amino acid sequence data Processing of X-ray diffraction data collected in oscillation mode Phaser crystallographic software PHENIX: building new software for automated crystallographic structure determination Coot: model-building tools for molecular graphics Procheck—a program to check the stereochemical quality of protein structures Crystal structure of the MAPK phosphatase Pyst1 catalytic domain and implications for regulated activation Crystal structure of the MAP kinase binding domain and the catalytic domain of human MKP5 A continuous spectrophotometric assay for inorganic phosphate and for measuring phosphate release kinetics in biological systems A continuous spectrophotometric method for the determination of glycogen phosphorylase-catalyzed reaction in the direction of glycogen synthesis Comparative protein structure modeling using Modeller All-atom empirical potential for molecular modeling and dynamics studies of proteins Scalable molecular dynamics with NAMD VMD: visual molecular dynamics Author contributions X.L. and C.-S.Z. performed the experiments and analysed the data. C.L. performed the MD simulation. Z.-X.W., J.-W.W. and S.-C.L. conceived and designed the experiments, and wrote the manuscript. Domain structures of ten human MKPs and the atypical VHR. On the basis of sequence similarity, protein structure, substrate specificity and subcellular localization, the ten members of MKP family can be divided into three groups. The first subfamily comprises MKP1, MKP2, PAC1 and hVH3, which are inducible nuclear phosphatases and can dephosphorylate ERK (and JNK, p38) MAPKs. The second subfamily contains MKP3, MKP4 and MKPX, which are cytoplasmic ERK-specific MKPs. The third subfamily comprises MKP5, MKP7 and hVH5, which were located in both nucleus and cytoplasm, and selectively inactivate JNK and p38. All MKPs contain both the CD and KBD domains, whereas VHR, an atypical MKP, only contains a highly conserved catalytic domain. In addition to the CD and KBD, MKP7 contains a unique long C-terminal region that contains NES, NLS and PEST motifs, which has no effect on the binding ability and phosphatase activity of MKP7 toward MAPKs. NES, nuclear export signal; NLS, nuclear localization signal; PEST, C-terminal sequence rich in prolines, glutamates, serines and threonines. MKP7-CD is crucial for JNK1 binding and enzyme catalysis. (a) Domain organization of human MKP7 and JNK1. The KBD and CD of MKP7 are shown in green and blue, and the N-lobe and C-lobe of JNK1 are coloured in lemon and yellow, respectively. The key structural elements are indicated. The colour scheme is the same in the following figures unless indicated otherwise. (b) Plots of initial velocity of the MKP7-catalysed reaction versus phospho-JNK1 concentration. The solid lines are best-fitting results according to equation (1). Each experiment was performed in replicate for at least three times. The error bars represent s.e.m. (c) Gel filtration analysis for interaction of JNK1 with MKP7-CD and MKP7-KBD. (d) GST-mediated pull-down assay for interaction of JNK1 with MKP7-CD and MKP7-KBD. The top panel shows the relative affinities of MKP7-CD and MKP7-KBD to JNK1, with the affinity of MKP7-CD defined as 100%; the middle panel is the electrophoretic pattern of MKP7 and JNK1 after GST pull-down assays. The protein amounts of MKP7 used are shown at the bottom. Structure of JNK1 in complex with MKP7-CD. (a) Ribbon diagram of JNK1–MKP7-CD complex in two views related by a 45° rotation around a vertical axis. (b) Structure of MKP7-CD with its active site highlight in cyan. The 2Fo−Fc omit map (contoured at 1.5σ) for the P-loop of MKP7-CD is shown at inset of b. (c) Structure of VHR with its active site highlighted in marine blue. (d) Close-up view of the JNK1–MKP7 interface showing interacting amino acids of JNK1 (orange) and MKP7-CD (cyan). The JNK1 is shown in surface representation coloured according to electrostatic potential (positive, blue; negative, red). (e) Interaction networks mainly involving helices α4 and α5 from MKP7-CD, and αG and α2L14 of JNK1. MKP7-CD is shown in surface representation coloured according to electrostatic potential (positive, blue; negative, red). Blue dashed lines represent polar interactions. (f) The 2Fo−Fc omit map (contoured at 1.5σ) clearly shows electron density for the 285FNFL288 segment of MKP7-CD. Mutational analysis on interactions between MKP7-CD and JNK1. (a) Effects of mutations in MKP7-CD on the JNK1 dephosphorylation (mean±s.e.m., n=3). Residues involved in hydrophobic and hydrophilic contacts are coloured in red and blue, respectively. (b) Gel filtration analysis for interaction of JNK1 with MKP7-CD mutant F285D. Mutant F285D and JNK1 were eluted as monomers, with the molecular masses of ∼17 and 44 kDa, respectively. However, in contrast to the wild-type MKP7-CD, mutant F285D did not co-migrate with JNK1. (c) Pull-down assays of MKP7-CD by GST-tagged JNK1 mutants. The top panel shows the relative affinities of MKP7-CD to JNK1 mutants, with the affinity of wild-type JNK1 defined as 100%, the middle panel is the electrophoretic pattern of MKP7-CD and JNK1 mutants after GST pull-down assays. The protein amounts of MKP7-CD used are shown at the bottom. (d) Circular dichroism spectra for MKP7-CD wild type and mutants. Measurements were averaged for three scans. (e) Circular dichroism spectra for JNK1 wild type and mutants. Measurements were averaged for three scans. (f) Effects of mutations in MKP7-CD on the pNPP hydrolysis reaction (mean±s.e.m., n=3). Comparison of CDK2-KAP and JNK1–MKP7-CD. (a) Superposition of the complex structures of CDK2-KAP (PDB 1FQ1) and JNK1–MKP7-CD. The N-lobe and C-lobe of CDK2 are coloured in grey and pink, respectively, and KAP is coloured in green. The interactions between these two proteins consist of three discontinuous contact regions, centred at the multiple hydrogen bonds between the pThr160 of CDK2 and the active site of KAP (region I). Interestingly, the recognition of CDK2 by KAP is augmented by a similar interface as that observed in the complex of JNK1 and MKP7-CD (region II). (b) Interactions networks at the auxiliary region II mainly involving helix α7 from KAP and the αG helix and following L14 loop of CDK2. The orientation of the panel is almost identical to those of Fig. 3d. The CDK2 is shown in surface representation coloured according to the electrostatic potential (positive, blue; negative, red). Residues of KAP and CDK2 are highlighted as green and red sticks, respectively. Blue dashed lines represent polar interactions. One remarkable difference between these two kinase-phosphatase complexes is that helix α6 of KAP (corresponding to helix α4 of MKP7-CD) plays little, if any, role in the formation of a stable heterodimer of CDK2 and KAP. (c) Sequence alignment of the JNK-interacting regions on MKPs. Residues of MKP7-CD involved in JNK1 recognition are indicated by cyan asterisks, and the conserved FXF-motif is highlighted in cyan. The secondary structure assignments of MKP7-CD and KAP are shown above and below each sequence. (d) Sequence alignment of the F-site regions on MAPKs. Residues of JNK1 involved in recognition of MKP7 are indicated by orange asterisks, and those forming the F-site are highlighted in yellow. FXF-motif is critical for controlling the phosphorylation of JNK and ultraviolet-induced apoptosis. (a–c) FXF-motif is essential for the dephosphorylation of JNK by MKP7. HEK293T cells were infected with lentiviruses expressing MKP7 and its mutants (1.0 μg). After 36 h infection, cells were untreated in a, stimulated with 30 μM etoposide for 3 h in b or irradiated with 25 J m−2 ultraviolet light at 30 min before lysis in c. Whole-cell extracts were then immunoblotted with antibody indicated. Shown is a typical immunoblot for phosphorylated JNK from three independent experiments. (d) F-site is required for JNK1 to interact with MKP7. HEK293T cells were co-transfected with MKP7 full-length (1.0 μg) and JNK1 (wild type or mutants as indicated, 1.0 μg). At 16 h post transfection, cells were lysed. Whole-cell extracts were then immunoprecipitated with antibody against Myc for MKP7; immunobloting was carried out with antibodies indicated. IP, immunoprecipitation; TCL, total cell lysate. Shown is a typical result from three independent experiments. (e) Effect of MKP7 (wild type or mutants) expression on ultraviolet-induced apoptosis. HeLa cells were infected with lentiviruses expressing MKP7 full-length and its mutants. At 36 h post infection, cells were irradiated with 25 J m−2 ultraviolet light and collected at 6 h after irradiation. Cells were then subjected to flow cytometry analysis. Apoptotic cells were determined by Annexin-V-APC/PI staining. The results using Annexin-V stain for membrane phosphatidylserine eversion, combined with propidium iodide (PI) uptake to evaluate cells whose membranes had been compromised. Staining with both Annexin-V and PI indicate apoptosis (upper right quadrant). The values shown in the lower left, and upper right quadrants of each panel represent the percentage of viable, and apoptotic cells, respectively. All results are representative of three independent experiments. (f) Statistical analysis of apoptotic cells (mean±s.e.m., n=3), *P\u003c0.05, ***P\u003c0.001 (ANOVA followed by Tukey's test). NS, not significant. MKP5-CD is crucial for JNK1 binding and enzyme catalysis. (a) Domain organization of human MKP5. The KBD and CD of MKP5 are shown in brown and grey, respectively. (b) Plots of initial velocity of the MKP5-catalysed reaction versus phospho-JNK1 concentration. The solid lines are best-fitting results according to the Michaelis–Menten equation with Km and kcat values indicated. Each experiment was performed in replicate for at least three times. The error bars represent s.e.m. (c) Structural comparison of the JNK-interacting residues on MKP5-CD (PDB 1ZZW) and MKP7-CD. The corresponding residues on MKP5 are depicted as orange sticks, and MKP5 residues numbers are in parentheses. (d) Gel filtration analysis for interaction of JNK1 with MKP5-CD and MKP5-KBD. (e) GST-mediated pull-down assays for interaction of JNK1 with MKP5-CD and MKP5-KBD. The panels are arranged the same as in Fig. 2d. (f) Effects of mutations in MKP5-CD on the JNK1 dephosphorylation (mean±s.e.m., n=3). (g) Effects of mutations in MKP5-CD on the pNPP hydrolysis reaction (mean±s.e.m., n=3). (h) Pull-down assays of MKP5-CD by GST-tagged JNK1 mutants. The panels are arranged the same as in Fig. 4c. Data collection and refinement statistics.  \tJNK1–MKP7-CD\t \tData collection*\t \t Space group\tP1\t \t Cell dimensions\t \t  a, b, c (Å)\t58.1, 74.8, 134.8\t \t  α, β, γ (°)\t76.9, 84.3, 67.4\t \t Resolution (Å)\t40.00–2.40 (2.49–2.40)†\t \t Rmerge\t6.4 (60.9)\t \t I/σI\t14.1 (1.9)\t \t Completeness (%)\t99.6 (99.9)\t \t Redundancy\t3.5 (3.4)\t \t \t \t \tRefinement\t \t Resolution (Å)\t38.02–2.40 (2.43–2.40)\t \t No. of reflections\t66889\t \t Rwork/Rfree\t21.7/23.9\t \t No. atoms\t \t  Protein\t14637\t \t  Ligand/ion\t4\t \t  Water\t457\t \t B-factors\t \t  Protein\t47.04\t \t  Ligand/ion\t30\t \t  Water\t37.28\t \t R.m.s.d.\t \t  Bond lengths (Å)\t0.015\t \t  Bond angles (°)\t1.564\t \t *The data set was collected from a single crystal. †Values in parentheses are for the highest resolution 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diff --git a/annotated_BioC_JSON/PMC4802085_ann.json b/annotated_BioC_JSON/PMC4802085_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..12a1ef294e8cab8046d50dbeb6b9932dc61be2bf
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4802085_ann.json
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+[{"sourceid":"4802085","sourcedb":"","project":"","target":"","text":"Haem-dependent dimerization of PGRMC1/Sigma-2 receptor facilitates cancer proliferation and chemoresistance Progesterone-receptor membrane component 1 (PGRMC1/Sigma-2 receptor) is a haem-containing protein that interacts with epidermal growth factor receptor (EGFR) and cytochromes P450 to regulate cancer proliferation and chemoresistance; its structural basis remains unknown. Here crystallographic analyses of the PGRMC1 cytosolic domain at 1.95 Å resolution reveal that it forms a stable dimer through stacking interactions of two protruding haem molecules. The haem iron is five-coordinated by Tyr113, and the open surface of the haem mediates dimerization. Carbon monoxide (CO) interferes with PGRMC1 dimerization by binding to the sixth coordination site of the haem. Haem-mediated PGRMC1 dimerization is required for interactions with EGFR and cytochromes P450, cancer proliferation and chemoresistance against anti-cancer drugs; these events are attenuated by either CO or haem deprivation in cancer cells. This study demonstrates protein dimerization via haem–haem stacking, which has not been seen in eukaryotes, and provides insights into its functional significance in cancer.  PGRMC1 binds to EGFR and cytochromes P450, and is known to be involved in cancer proliferation and in drug resistance. Here, the authors determine the structure of the cytosolic domain of PGRMC1, which forms a dimer via haem–haem stacking, and propose how this interaction could be involved in its function. Much attention has been paid to the roles of haem-iron in cancer development. Increased dietary intake of haem is a risk factor for several types of cancer. Previous studies showed that deprivation of iron or haem suppresses tumourigenesis. On the other hand, carbon monoxide (CO), the gaseous mediator generated by oxidative degradation of haem via haem oxygenase (HO), inhibits tumour growth. Thus, a tenuous balance between free haem and CO plays key roles in cancer development and chemoresistance, although the underlying mechanisms are not fully understood. To gain insight into the underlying mechanisms, we took chemical biological approaches using affinity nanobeads carrying haem and identified progesterone-receptor membrane component 1 (PGRMC1) as a haem-binding protein from mouse liver extracts (Supplementary Fig. 1). PGRMC1 is a member of the membrane-associated progesterone receptor (MAPR) family with a cytochrome b5-like haem-binding region, and is known to be highly expressed in various types of cancers. PGRMC1 is anchored to the cell membrane through the N-terminal transmembrane helix and interacts with epidermal growth factor receptor (EGFR) and cytochromes P450 (ref). While PGRMC1 is implicated in cell proliferation and cholesterol biosynthesis, the structural basis on which PGRMC1 exerts its function remains largely unknown. Here we show that PGRMC1 exhibits a unique haem-dependent dimerization. The dimer binds to EGFR and cytochromes P450 to enhance tumour cell proliferation and chemoresistance. The dimer is dissociated to monomers by physiological levels of CO, suggesting that PGRMC1 serves as a CO-sensitive molecular switch regulating cancer cell proliferation. Results X-ray crystal structure of PGRMC1 We solved the crystal structure of the haem-bound PGRMC1 cytosolic domain (a.a.72–195) at 1.95 Å resolution (Supplementary Fig. 2). In the presence of haem, PGRMC1 forms a dimeric structure largely through hydrophobic interactions between the haem moieties of two monomers (Fig. 1a, Table 1 and Supplementary Fig. 3; a stereo-structural image is shown in Supplementary Fig 4). While the overall fold of PGRMC1 is similar to that of canonical cytochrome b5, their haem irons are coordinated differently. In cytochrome b5, the haem iron is six-coordinated by two axial histidine residues. These histidines are missing in PGRMC1, and the haem iron is five-coordinated by Tyr113 (Y113) alone (Fig. 1b and Supplementary Fig. 3). A homologous helix that holds haem in cytochrome b5 is longer, shifts away from haem, and does not form a coordinate bond in PGRMC1 (Fig. 1c). Consequently, the five-coordinated haem of PGRMC1 has an open surface that allows its dimerization through hydrophobic haem–haem stacking. Contrary to our finding, Kaluka et al. recently reported that Tyr164 of PGRMC1 is the axial ligand of haem because mutation of this residue impairs haem binding. Our structural data revealed that Tyr164 and a few other residues such as Tyr107 and Lys163 are in fact hydrogen-bonded to haem propionates. This is consistent with observations by Min et al. that Tyr 107 and Tyr113 of PGRMC1 are involved in binding with haem. These amino acid residues are conserved among MAPR family members (Supplementary Fig. 5a), suggesting that these proteins share the ability to exhibit haem-dependent dimerization. PGRMC1 exhibits haem-dependent dimerization in solution In the PGRMC1 crystal, two different types of crystal contacts (chain A–A″ and A–B) were observed in addition to the haem-mediated dimer (chain A–A′) (Supplementary Figs 3 and 6a). To confirm that haem-assisted dimerization of PGRMC1 occurs in solution, we analysed the structure of apo- and haem-bound PGMRC1 by two-dimensional nuclear magnetic resonance (NMR) using heteronuclear single-quantum coherence and transverse relaxation-optimized spectroscopy (Supplementary Figs 6b and 7). NMR signals from some amino acid residues of PGRMC1 disappeared due to the paramagnetic relaxation effect of haem (Supplementary Figs 6b); these residues were located in the haem-binding region. When chemical shifts of apo- and haem-bound forms of PGMRC1 were compared, some amino acid residues close to those which disappeared because of the paramagnetic relaxation effect of haem exhibit notable chemical shifts (Supplementary Fig. 6a,b; dark yellow). However, at the interfaces of the other possible dimeric structures (Supplementary Fig. 6a, chain A–A″; cyan and chain A–B; violet), no significant difference was observed. Furthermore, free energy of dissociation predicted by PISA suggested that the haem-mediated dimer is stable in solution while the other potential interactions are not. We also attempted to predict the secondary structure of PGRMC1 through NMR data by calculating with TALOS+ program (Supplementary Fig. 8); the prediction suggested that the overall secondary structure is comparable between apo- and haem-bound forms of PGRMC1 in solution. We analysed the haem-dependent dimerization of the PGRMC1 cytosolic domain (a.a.44–195) in solution (Fig. 2 and Table 2). Mass spectrometry (MS) analyses under non-denaturing condition demonstrated that the apo-monomer PGRMC1 resulted in dimerization by binding with haem (Fig. 2a). It should be noted that a disulfide bond between two Cys129 residues is observed in the crystal of PGRMC1 (Fig. 1a), while Cys129 is not conserved among the MAPR family proteins (Supplementary Fig. 5a). This observation led us to examine whether or not the disulfide bond contributes to PGRMC1 dimerization. MS analyses under non-denaturing conditions clearly showed that the Cys129Ser (C129S) mutant is dimerized in the presence of haem, indicating that the haem-mediated dimerization of PGRMC1 occurs independently of the disulfide bond formation via Cys129 (Fig. 2a). Supporting this, MS analyses under denaturing conditions showed that haem-mediated PGRMC1 dimer is completely dissociated into monomer, indicating that dimerization of this kind is not mediated by any covalent bond such as disulfide bond (Supplementary Fig. 9). We also analysed the haem-dependent dimerization of PGRMC1 by diffusion-ordered NMR spectroscopy (DOSY) analyses (Table 2, Supplementary Fig. 10). The results suggested that the hydrodynamic radius of haem-bound PGRMC1 is larger than that of apo-PGRMC1. To further evaluate changes in molecular weights in dimerization of PGRMC1, sedimentation velocity analytical ultracentrifugation (SV-AUC) analysis was carried out. Whereas the wild-type (wt) apo-PGRMC1 appeared at a 1.9 S peak as monomer, the haem-binding PGRMC1 was converted into dimer at a 3.1 S peak (Fig. 2b). Similarly, the C129S mutant of PGRMC1 converted from monomer to dimer by binding to haem (Fig. 2b). SV-AUC analyses also allowed us to examine the stability of haem/PGRMC1 dimer. To this end, we used different concentrations (3.5–147 μmol l−1) of haem-bound PGRMC1 protein (a.a. 72–195), which were identical to that used in the crystallographic analysis. The sedimentation coefficients calculated on the basis of the crystal structure were 1.71 S for monomer and 2.56 S for dimer (Supplementary Fig. 11, upper panel). The results showed that the PGRMC1 dimer is not dissociated into monomer at all concentrations examined (Supplementary Fig. 11, lower panel), suggesting that the Kd value of haem-mediated dimer of PGRMC1 is under 3.5 μmol l−1. A value of this kind implies that the PGRMC1 dimer is more stable than other dimers of extracellular domain of membrane proteins such as Toll like receptor 9 (dimerization Kd of 20 μmol l−1) (ref.) and plexin A2 receptor (dimerization Kd higher than 300 μmol l−1) (ref.). The current analytical data confirmed that apo-PGRMC1 monomer converts into dimer by binding to haem in solution (Table 2). We also showed by haem titration experiments that haem binding to PGRMC1 was of low affinity with a Kd value of 50 nmol l−1; this is comparable with that of iron regulatory protein 2, which is known to be regulated by intracellular levels of haem (Fig. 2c and Supplementary Table 1). These results raised the possibility that the function of PGRMC1 is regulated by intracellular haem concentrations. CO inhibits haem-dependent dimerization of PGRMC1 Crystallographic analyses revealed that Tyr113 of PGRMC1 is an axial ligand for haem and contributes to haem-dependent dimerization (Fig. 1a). Analysis of UV-visible spectra revealed that the heme of PGRMC1 is reducible from ferric to ferrous state, thus allowing CO binding (Fig. 3a). Furthermore, the UV-visible spectrum of the wild type PGRMC1 was the same as that of the C129S mutant of PGRMC1, and the R/Z ratio determined by the intensities between the Soret band (394 nm) peak and the 274-nm peak showed that these proteins were fully loaded with haem (Supplementary Fig. 12). Analysis of the ferric form of PGRMC1 using resonance Raman spectroscopy (Supplementary Fig. 13) showed that the relative intensity of oxidation and spin state marker bands (ν4 and ν3) is close to 1.0, which is consistent with it being a haem protein with a proximal Tyr coordination. A specific Raman shift peaking at vFe–CO=500 cm−1 demonstrated that the CO-bound haem of PGRMC1 is six-coordinated (Supplementary Fig. 13). Since PGRMC1 dimerization involves the open surface of haem on the opposite side of the axial Tyr113, no space for CO binding is available in the dimeric structure (Fig. 3b). This prompted us to ask if CO binding to haem causes dissociation of the PGRMC1 dimer. Analysis by gel filtration chromatography revealed that the relative molecular sizes of the wild-type and the C129S mutant of PGRMC1 are increased by adding haem to apo-PGRMC1 regardless of the oxidation state of the iron (Fig. 3c), which is in agreement with the results in Table 1. CO application to ferrous PGRMC1 abolished the haem-dependent increase in its molecular size. Under this reducing condition in the presence of dithionite, analyses of UV-visible spectra indicated that CO-binding with haem-PGRMC1 is stable, showing only 20% reduction of the absorbance at 412 nm within 2 h (Supplementary Fig. 14). Furthermore, the Tyr113Phe (Y113F) mutant of PGRMC1 was not responsive to haem. These results suggest that CO favours the six-coordinate form of haem and interferes with the haem-mediated dimerization of PGRMC1. To examine the inhibitory effects of CO on haem-mediated PGRMC1 dimerization, SV-AUC analysis was carried out. The peak corresponding to the haem/PGRMC1 dimer was detected under reducing conditions in the presence of dithionite (Supplementary Fig. 15, middle panel). Under these circumstances, CO application induced dissociation of the haem-mediated dimers of PGRMC1 to generate a peak of monomers (Supplementary Fig. 15, lower panel). These observations raised the transition model for structural regulation of PGRMC1 in response to haem (Fig. 3d). As mentioned above, apo-PGRMC1 exists as monomer. By binding with haem (binding Kd=50 nmol l−1), PGRMC1 forms a stable dimer (dimerization Kd\u003c\u003c3.5 μmol l−1) through stacking of the two open surfaces of the five-coordinated haem molecules in each monomer. CO induces the dissociation of the haem-mediated dimer of PGRMC1 by interfering with the haem-stacking interface via formation of the six-coordinated CO-haem-PGRMC1 complex. Such a dynamic structural regulation led us to further examine the regulation of PGRMC1 functions in cancer cells. PGRMC1 dimerization is required for binding to EGFR Because PGRMC1 is known to interact with EGFR and to accelerate tumour progression, we examined the effect of haem-dependent dimerization of PGRMC1 on its interaction with EGFR by using purified proteins. As shown in Fig. 4a, the cytosolic domain of wild-type PGRMC1, but not the Y113F mutant, interacted with purified EGFR in a haem-dependent manner. This interaction was disrupted by the ruthenium-based CO-releasing molecule, CORM3, but not by RuCl3 as a control reagent (Fig. 4b). We further analysed the intracellular interaction between PGRMC1 and EGFR. FLAG-tagged PGRMC1 ectopically expressed in human colon cancer HCT116 cells was immunoprecipitated with anti-FLAG antibody, and co-immunoprecipitated EGFR and endogenous PGRMC1 binding to FLAG-PGRMC1 were detected by Western blotting (Fig. 4c). The C129S mutant of PGRMC1 also interacted with endogenous PGRMC1 and EGFR (Supplementary Fig. 16). Whereas FLAG-tagged wild-type PGRMC1 interacted with endogenous PGRMC1 and EGFR, the Y113F mutant did not. We also examined the effect of succinylacetone (SA), an inhibitor of haem biosynthesis (Fig. 4d). As expected, SA significantly reduced PGRMC1 dimerization and its interaction with EGFR (Fig. 4e), indicating that haem-mediated dimerization of PGMRC1 is critical for its binding to EGFR. PGRMC1 dimer facilitates EGFR-mediated cancer growth Next, we investigated the functional significance of PGRMC1 dimerization in EGFR signaling. EGF-induced phosphorylations of EGFR and its downstream targets AKT and ERK were decreased by PGRMC1 knockdown (PGRMC1-KD) (Fig. 4f). Similarly, EGFR signaling was suppressed by treatment of HCT116 cells with SA (Fig. 4g) or CORM3 (Fig. 4h). These results suggested that haem-mediated dimerization of PGRMC1 is critical for EGFR signaling. To further investigate the role of the dimerized form of PGRMC1 in cancer proliferation, we performed PGRMC1 knockdown-rescue experiments using FLAG-tagged wild-type and Y113F PGRMC1 expression vectors, in which silent mutations were introduced into the nucleotide sequence targeted by shRNA (Fig. 5a). While proliferation of HCT116 cells was not affected by knocking down PGRMC1, PGRMC1-KD cells were more sensitive to the EGFR inhibitor erlotinib than control HCT116 cells, and the knockdown effect was reversed by co-expression of shRNA-resistant wild-type PGRMC1 but not of the Y113F mutant (Fig. 5b). Chemosensitivity enhancement by two different shRNAs to PGRMC1 was seen also in HCT116 cells and human hepatoma HuH7 cells (Supplementary Fig. 17). Furthermore, PGRMC1-KD inhibited spheroid formation of HCT116 cells in culture, and this inhibition was reversed by co-expression of wild-type PGRMC1 but not of the Y113F mutant (Fig. 5c and Supplementary Fig. 18). Thus, PGRMC1 dimerization is important for cancer cell proliferation and chemoresistance. We examined the role of PGRMC1 in metastatic progression by xenograft transplantation assays using super-immunodeficient NOD/scid/γnull (NOG) mice. Ten days after intra-splenic implantation of HCT116 cells that were genetically tagged with a fluorescent protein Venus, the group implanted with PGRMC1-KD cells showed a significant decrease of liver metastasis in comparison with the control group (Fig. 5d). Interaction of PGRMC1 dimer with cytochromes P450 Since PGRMC1 has been shown to interact with cytochromes P450 (ref), we investigated whether the haem-mediated dimerization of PGRMC1 is necessary for their interactions. Recombinant CYP1A2 and CYP3A4 including a microsomal formulation containing cytochrome b5 and cytochrome P450 reductase, drug-metabolizing cytochromes P450, interacted with wild-type PGRMC1, but not with the Y113F mutant, in a haem-dependent manner (Fig. 6a,b). Moreover, the interaction of PGRMC1 with CYP1A2 was blocked by CORM3 under reducing conditions (Fig. 6c), indicating that PGRMC1 dimerization is necessary for its interaction with cytochromes P450. Doxorubicin is an anti-cancer reagent that is metabolized into inactive doxorubicinol by CYP2D6 and CYP3A4 (Fig. 6d). PGRMC1-KD significantly suppressed the conversion of doxorubicin to doxorubicinol (Fig. 6d) and increased sensitivity to doxorubicin (Fig. 6e). Enhanced doxorubicin sensitivity was modestly but significantly induced by PGRMC1-KD. This effect was reversed by co-expression of the wild-type PGRMC1 but not of the Y113F mutant, suggesting that PGRMC1 enhances doxorubicin resistance of cancer cells by facilitating its degradation via cytochromes P450. To gain further insight into the interaction between PGRMC1 and cytochromes P450, surface plasmon resonance analyses were conducted using recombinant CYP51 and PGRMC1. This was based on a previous study showing that PGRMC1 binds to CYP51 and enhances cholesterol biosynthesis by CYP51 (refs). CYP51 interacted with PGRMC1 in a concentration-dependent manner in the presence of haem, but not in its absence (Supplementary Fig. 19), suggesting the requirement for the haem-dependent dimerization of PGRMC1. The Kd value of PGRMC1 binding to CYP51 was in a micromolar range and comparable with those of other haem proteins, such as cytochrome P450 reductase and neuroglobin/Gαi1 (ref.), suggesting that haem-dependent PGRMC1 interaction with CYP51 is biologically relevant. Discussion In this study, we showed that PGRMC1 dimerizes by stacking interactions of haem molecules from each monomer. Recently, Lucas et al. reported that translationally-controlled tumour protein was dimerized by binding with haem, but its structural basis remains unclear. This is the report showing crystallographic evidence that indicates roles of the direct haem–haem stacking in haem-mediated dimerization in eukaryotes, although a few examples are known in bacteria. Sequence alignments show that haem-binding residues (Tyr113, Tyr107, Lys163 and Tyr164) in PGRMC1 are conserved among MAPR proteins (Supplementary Fig. 5). In the current study, the Y113 residue plays a crucial role for the haem-dependent dimerization of PGRMC1 and resultant regulation of cancer proliferation and chemoresistance (Figs 5c and 6e). Since the Y113 residue is involved in the putative consensus motif of phosphorylation by tyrosine kinases such as Abl and Lck, we investigated whether phosphorylated Y113 is present in HCT116 cells by ESI-MS analysis. It was, however, undetectable under current experimental conditions (Supplementary Fig. 20). Recently, Peluso et al. reported that PGRMC1 binds to PGRMC2, suggesting that MAPR family members may also undergo haem-mediated heterodimerization. We showed that the haem-mediated dimer of PGRMC1 enables interaction with different subclasses of cytochromes P450 (CYP) (Fig. 6). While the effects of PGRMC1 on cholesterol synthesis mediated by CYP51 have been well documented in yeast and human cells, it has not been clear whether drug-metabolizing CYP activities are regulated by PGRMC1. Szczesna-Skorupa and Kemper reported that PGRMC1 exhibited an inhibitory effect on CYP3A4 drug metabolizing activity by competitively binding with cytochrome P450 reductase (CPR) in HEK293 or HepG2 cells. On the other hand, Oda et al. reported that PGRMC1 had no effect to CYP2E1 and CYP3A4 activities in HepG2 cell. Several other groups showed that PGRMC1 enhanced chemoresistance in several cancer cells such as uterine sarcoma, breast cancer, endometrial tumour and ovarian cancer; however, no evidence of PGRMC1-dependent regulation of CYP activity was provided. Our results showed that PGRMC1 contributes to enhancement of the doxorubicin metabolism, which is mediated by CYP2D6 or CYP3A4 in human colon cancer HCT116 cells (Fig. 6d). While the effects of structural diversity of CYP family proteins and interactions with different xenobiotic substrates should further be examined, the current results suggest that the interaction of drug-metabolizing CYPs with the haem-mediated dimer of PGRMC1 plays a crucial role in regulating their activities. We showed that haem-mediated dimerization of PGRMC1 enhances proliferation and chemoresistance of cancer cells through binding to and regulating EGFR and cytochromes P450 (illustrated in Fig. 7). Since the haem-binding affinity of PGRMC1 is lower than those of constitutive haem-binding proteins such as myoglobin, PGMRC1 is probably interconverted between apo-monomer and haem-bound dimer forms in response to changes in the intracellular haem concentration. Considering microenvironments in and around malignant tumours, the haem concentration in cancer cells is likely to be elevated through multiple mechanisms, such as (i) an increased intake of haem, (ii) mutation of enzymes in TCA cycle (for example, fumarate hydratase) that increases the level of succinyl CoA, a substrate for haem biosynthesis and (iii) metastasis to haem-rich organs such as liver, brain and bone marrow. Moreover, exposure of cancer cells to stimuli such as hypoxia, radiation and chemotherapy causes cell damages and leads to protein degradation, resulting in increased levels of TCA cycle intermediates and in an enhanced haem biosynthesis. On the other hand, excessive haem induces HO-1, the enzyme that oxidatively degrades haem and generates CO. Thus, HO-1 induction in cancer cells may inhibit the haem-mediated dimerization of PGRMC1 through the production of CO and thereby suppress tumour progression. This idea is consistent with the observation that HO-1 induction or CO inhibits tumour growth. Besides the regulatory roles of PGRMC1/Sigma-2 receptor in proliferation and chemoresistance in cancer cells (ref.), recent reports show that PGRMC1 is able to bind to amyloid beta oligomer to enhance its neurotoxicity. Furthermore, Sigma-2 ligand-binding is decreased in transgenic amyloid beta deposition model APP/PS1 female mice. These results suggest a possible involvement of PGRMC1 in Alzheimer's disease. The roles of haem-dependent dimerization of PGRMC1 in the functional regulation of its target proteins deserve further studies to find evidence that therapeutic interventions to interfere with the function of the dimer may control varied disease conditions. Methods Materials Recombinant EGF, CYP1A2 and CYP3A4 proteins were purchased from Sigma. Erlotinib was purchased from Cayman. Doxorubicin was purchased from Wako. Anti-FLAG (M2) antibody, FLAG peptide and anti-FLAG antibody-conjugated agarose were purchased from Sigma. Haemin and protoporphyrin-IX (PP-IX) were purchased from Porphyrin Science. Plasmid constructions Human PGRMC1 cDNA was cloned from the cDNA library of HuH7 cells. The PGMRC1 (a.a.44–195 for in vitro studies and a.a.72–195 for crystallographic analyses and SV-AUC) cDNA fragment was amplified with PCR, digested with Bam HI and Sal I and then ligated into pGEX6P-1 (GE Healthcare). For NMR analysis, the PGRMC1 (a.a.44–195) cDNA fragment was amplified with PCR (with primers containing the factor Xa site) and ligated into the Bam HI and Sal I sites of pGEX6P-1. The full-length PGRMC1 cDNA fragment containing resistant sequences for shRNA was generated by using the primers (Supplementary Methods), and ligated into the Eco RI and Bam HI sites of the C-terminus of the 3xFLAG-tagged expression vector p3xFLAG CMV14 (Sigma). Preparations of recombinant proteins pGEX-PGRMC1 wt, Y113F or C129S mutant expression vectors were transformed into BL21 (DE3), and the bacteria were incubated in LB with ampicillin at 37 °C until OD600 reached at 0.8. Protein expression was induced by 1 mmol l−1 isopropyl-β-thiogalactopyranoside for 4 h at 37 °C. Cell pellets were resuspended in the buffer containing 20 mmol l−1 Tris-HCl (pH 7.5), 100 mmol l−1 NaCl and 0.1% Tween 20, sonicated twice for 5 min at 4 °C and centrifuged at 20,000 × g for 30 min. The supernatant was incubated with glutathione Sepharose 4B (GE Healthcare) for 1 h at 4 °C. The resin was then washed five times with the same buffer, and the GST tag was cleared by addition of Precision Protease (GE Healthcare) and further incubation for 16 h at 4 °C. The apo-PGRMC1 was prepared by eliminating the bacterial holo-PGRMC1 with size-exclusion chromatography (Superdex 200; GE Healthcare). Haem-bound PGRMC1 were prepared by treatment with 100 μmol l−1 haemin and purified by size-exclusion chromatography. The PGRMC1 protein treated with Precision Protease to cleave the GST-tag contained additional amino acid residues (GPLGSEF) derived from the restriction site and the protease site for Precision Protease at the N-terminal region of PGRMC1. Isotope-labelled PGRMC1 proteins for NMR analyses were prepared by growing cells (BL21 (DE3)) in minimal M9 media in H2O or 99.9% 2H2O, including ampicillin, metals, vitamins, 15N-ammonium chloride and 13C or 12C glucose as sources of nitrogen and carbon, respectively. These procedures were followed by addition of 1 mmol l−1 isopropyl-β-thiogalactopyranoside for 40 h at 20 °C. Protein purification was performed as mentioned above. The GST tag was cleaved with Factor Xa (GE Healthcare). The proteins were treated with Factor Xa to cleave the GST tag at the direct site of N-terminal region of PGRCM1 (a.a.44–195). X-ray crystallography PGRMC1 (a.a.72–195) crystals were grown at 20 °C using hanging-drop vapour diffusion by mixing equal volumes of protein solution and reservoir solution containing 100 mmol l−1 sodium cacodylate (pH 6.5) and 1.26–1.45 mol l−1 ammonium sulphate. Brown crystals reached maximum size in three weeks. The crystals were soaked in reservoir solution containing 30% trehalose and then flash-frozen in liquid nitrogen. The X-ray diffraction data for PGRMC1 crystals were collected at SPring-8 BL41XU and processed with XDS. The initial phase was obtained by single-wavelength anomalous dispersion, using a dataset collected at 1.73 Å with PHENIX AutoSol. Manual modeling and refinement were performed with COOT and phenix.refine. The deposited model was refined to a resolution of 1.95 Å. In this model, 95.4% of the residues were in favoured regions of Ramachandran plot, and all the others were in allowed regions. Data collection and refinement statistics are shown in Table 1. Molecular figures were created by PyMOL (Schrodinger, LLC. The PyMOL Molecular Graphics System, Version 1.5.0.3). Mass spectrometry analyses The purified PGRMC1 (a.a.44–195) proteins, the wild-type (apo and haem) and the C129S mutant (apo and haem), which included additional amino acid residues (GPLGSEF), were buffer-exchanged into 100 mmol l−1 ammonium acetate, pH 7.5, by passing the proteins through a Bio-Spin 6 column (Bio-Rad). The buffer-exchanged PGRMC1 wild-type (apo and haem) and PGMRC1 C129S mutants were immediately analysed by nanoflow electrospray ionization MS using gold-coated glass capillaries made in house. In the case of ESI-MS analyses under denaturing conditions, buffer-exchanged proteins were denatured before ESI-MS analyses by adding aliquots of formic acid at final concentration of 30%). Spectra were recorded on a SYNAPT G2 HDMS mass spectrometer (Waters, Manchester, UK) in positive ionization mode at 1.20 kV with a 120 V sampling cone voltage. The spectra were calibrated using 1 mg ml−1 caesium iodide and analysed with Mass Lynx software (Waters). SV-AUC analyses SV-AUC experiments were performed in a ProteomeLab XL-I analytical ultracentrifuge (Beckman Coulter) equipped with 4-hole An60Ti rotors at 20 °C using Beckman Coulter 12-mm double-sector aluminium centerpieces and sapphire windows. Recombinant PGRMC1 proteins were diluted with the buffer (20 mmol l−1 Tris-HCl (pH 7.5) and 100 mmol l−1 NaCl) at the indicated concentration. Scanning was performed as quickly as possible at 262,080 g at 6.5 cm (60,000 rpm), between 6.0 and 7.2 cm from the axis of rotation with a radial increment of 30 μm using an absorbance optical system. The sedimentation coefficient distributions were obtained using the c(s) method of SEDFIT. The partial specific volume, buffer density and viscosity were calculated using the program SEDNTERP 1.09 and were 0.7216, cm3 g−1, 1.00293, g ml−1, and 1.017 cP, respectively. The sedimentation coefficient of PGRMC1 (a.a.72–195) was calculated with the UltraScan Solution Modeler (US-SOMO) suite using the crystal structure determined in this study. To analyse the effect of CO, protein samples were prepared in a deaerated solution and treated with dithionite at 5 mmol l−1 and/or CO gas. DOSY analysis Diffusion-ordered 2D NMR spectroscopy (DOSY) was used to investigate the oligomerization state of PGRMC1 (a.a.44–195) induced by haem binding. Apo- or haem-bound PGRMC1 and reference proteins (including hen egg lysozyme, ovalbumin and bovine serum albumin (BSA)) dissolved in 50 mmol l−1 phosphate buffer (pH 7.0) containing 5% D2O were measured at 25 °C. The protein concentrations were 0.15–0.2 mmol l−1. DOSY spectra were measured using the stimulated echo sequence with a longitudinal-eddy-current delay, and diffusion coefficients were calculated from signals in the aliphatic regions using the software TOPSPIN (Bruker). The signal intensities fit the Stejskal-Tanner equation: where I represents the signal intensity when gradient pulses of length δ are applied at strength g, varying from 2 to 95% of the full gradient strength (55 G cm−1). The diffusion coefficient D25 is estimated by curve fitting with the term I(0), which corresponds to the signal intensity at a gradient strength of 0. The term γ represents the gyromagnetic ratio, and Δ represents the delay between two sets of gradients responsible for the stimulated echo. In this study, δ was set to 8 or 10 ms, and Δ was set to 40 ms. The hydrodynamic radii of the proteins were estimated on the basis of the Stokes–Einstein equation as follows: where T is the absolute temperature, r is the hydrodynamic radius of the spherical molecule, η is the viscosity of the solvent, and κB is the Boltzmann constant. The molecular weights (MWs) of apo- and haem-bound PGRMC1 proteins were estimated from a relationship between r and MWs. MWs of apo and haem-bound PGRMC1 proteins were obtained from the linear-fitting of measured r values for the reference proteins with known MWs according to the following equation: MW=1.2864 (r3)+8.0411. UV-visible absorption spectrometry and haem titration analysis UV-visible absorption spectra of the protein were recorded with a V-660 (Jasco) spectrophotometer at room temperature. Haem binding was tracked by difference spectroscopy in the Soret region of the UV-visible spectrum. Successive aliquots of 0.5 mmol l−1 haemin in N,N-dimethylformamide were added to both the sample cuvette, which contained 10 μmol l−1 apo-PGRMC1 (a.a.44–195), and the reference cuvette. Spectra were recorded 3 min after the addition of each haem aliquot. The absorbance difference at 400 nm was plotted as a function of haem concentration, and the dissociation constant (Kd) was calculated using a quadratic binding equation. Gel filtration chromatography Recombinant PGRMC1 (a.a.44–195) (10 μg) wt, Y113F or C129S mutant, treated with 5 mmol l−1 sodium dithionite and/or CO gas or left untreated, was separated on a Superdex 200 column equilibrated in buffer containing 20 mmol l−1 Tris-HCl (pH 7.5) and 100 mmol l−1 NaCl using a SMART system (GE Healthcare). To prepare the reducing conditions for ferrous haem proteins, the running buffer was deaerated by boiling and saturating it with argon gas according to modified versions of previously reported methods. Namely, immediately after adding dithionite to give a final concentration of 5 mmol l−1, the buffer was equilibrated into the column. The SMART system was sealed with gas-tight taping to maintain anaerobic conditions. Separations of proteins were completed within 1 h. Protein samples were also prepared in the deaerated solution and treated with dithionite at 5 mmol l−1 and/or CO gas, right before being injected into the column. Fractions were then subjected to SDS-PAGE under ambient conditions and visualized by silver staining. The size of proteins was estimated using molecular mass markers (thyroglobin, 669 kDa; catalase, 232 kDa; aldolase, 150 kDa; bovine albumin, 66 kDa and β-amylase, 20 kDa). Results showing that the molecular size of PGRMC1 became smaller in CO-treated conditions (Fig. 3) were collected ∼60 min after the start of experiments. The stability of CO-binding to PGRMC1 was examined with UV-visible absorption spectra to chase temporal alterations for 2 h, as shown in Supplementary Fig. 14. In vitro binding assays For in vitro binding assays, EGFR protein was obtained from ENZO (BML-SE116) as full length protein isolated from human A431 cells. Human CYP1A2, CYP3A4 proteins purified as a microsomal formulation containing cytochrome b5 and cytochrome P450 reductase were obtained from Sigma (C1561 and C4982, respectively). Proteins for human CYP1A2, CYP3A4 or EGFR (1 μg) were incubated with 10 μg of FLAG-PGRMC1 (a.a.44–195) treated with or without 50 μmol l−1 haemin in 500 μl of binding buffer containing 20 mmol l−1 HEPES-NaOH (pH 7.9), 100 mmol l−1 NaCl, 0.2 mmol l−1 EDTA, 10% glycerol and 0.1% NP40 for 60 min at room temperature. 5 mmol l−1 sodium dithionite was added to produce the reducing conditions specified in the aforementioned methods, and the effects of CORM3 or RuCl3 at 10 μmol l−1 were examined. Then, 10 μl of equilibrated anti-FLAG (M2) agarose was added to the mixture, which was then incubated for 60 min at room temperature. Bound proteins were washed three times with 200 μl of binding buffer and eluted with 10 μl of 2 μg ml−1 FLAG peptide. The eluates were subjected to SDS-PAGE and visualized by Western blotting using antibodies against CYP1A2, CYP3A4 (Santa Cruz: sc-30085 and sc-53850, respectively), FLAG and EGFR (Cell signaling: #2232S). Cell culture analyses The human colon cancer cell line HCT116 and human hepatoma cell line HuH7 were maintained in DMEM medium containing 10% FCS. To generate a stable PGRMC1 knockdown cell line, lentivirus vectors encoding a control or PGRMC1 targeting shRNA sequence were transfected into 293FT cells. The lentivirus was prepared according to the manufacturer's instructions (Invitrogen). HCT116 and HuH7 cells were infected with the lentivirus, and a stable cell line was selected by maintaining the cells in medium containing 10 μg ml−1 blasticidin (Invitrogen) for 1 week. For co-immunoprecipitation assay, the expression vector of FLAG-PGRMC1 or an empty vector into HCT116 by using a transfection reagent Lipofectamine 2000 (Invitrogen). Cells were incubated with or without 250 μ mol l−1 succinylacetone (SA) for 48 h, and the cells were then lysed with NP40 lysis buffer (20 mmol l−1 Tris-HCl (pH 7.5), 150 mmol l−1 NaCl, 1% NP40). The lysates were incubated with 10 μl of equilibrated anti-FLAG (M2) agarose for 60 min at room temperature. Bound proteins were washed three times, and were subjected to SDS-PAGE and visualized by Western blotting using antibodies against PGRMC1 (NOVUS: NBP1–83220) and EGFR. For analysis of EGFR signaling, cells were incubated overnight with serum-deprived medium, and then 100 ng ml−1 EGF was added for 5 min. Cells were lysed with RIPA buffer, and the lysates were subjected to SDS-PAGE and visualized by Western blotting using antibodies against PGRMC1, EGFR, phospho-Y1068 EGFR (Cell signaling: #2234S), AKT (Cell signaling: #9272S), phospho-S473AKT (Cell signaling: #4060S), ERK (Cell signaling: #4695S) and phospho-T185 Y187 ERK (Invitrogen: 44680 G). To analyse proliferation of HCT116 cells, Lipofectamine 2000 (Invitrogen) was used to transfect the shRNA-resistant expression vector of FLAG-PGRMC1 or an empty vector into HCT116 control or PGRMC1-knockdown cells. After 24 h, the cells were seeded and incubated for 12 h on a 96-well plate, after which erlotinib or doxorubicin was added for 24 h. Cell viability was determined by using an MTT assay kit (Millipore) according to the manufacturer's instructions. For analysis of spheroid formation of HCT116 cells, the shRNA-resistant expression vector of FLAG-PGRMC1 or an empty vector was transfected as described above. After 24 h, cells were seeded at 1 × 104 cells per well onto a 96-well spheroid culture plate (NanoCulture plate with a microsquare pattern, SCIVAX Corp.) and incubated for three days. The size of individual spheroid was determined by measuring their optical areas using Image-J and by calculating the apparent radius (r) to estimate their apparent volume (v) according to the following formula: v=4/3 × πr3. Measurements of intracellular haem concentrations To measure protohaem (haem b) concentrations, LC-UV and LC-MS analyses were performed for quantification and molecular identification, respectively. Briefly, HCT116 cells (1 × 107 cells) were treated with vehicle or 250 μmol l−1 SA for 48 h. After centrifugation, haem b was extracted from cell pellets twice by adding acetone containing 30% formate, followed by a 5 min sonication and centrifugation. The supernatant was collected, and the solvent was evaporated. The dried residues were re-dissolved in acetonitrile containing 0.2% formate and subjected to a LCMS-8030 system equipped with photodiode array (PDA) detector (SPD-20A) (Shimadzu Corporation, Kyoto, Japan). Haem b was detected by monitoring the absorption at 400 nm. Its identity was confirmed by simultaneous mass spectrometric analysis at m/z 616. Analyses of doxorubicin metabolism HCT116 cells (5 × 106 cells/10 cm dish) were cultured for 2 days, after which the cells were cultured in the presence of 0.3 μmol l−1 doxorubicin overnight. The cells were lysed with methanol containing internal standard compounds, and then water-soluble fraction was separated by liquid-liquid extraction (chloroform: methanol: water=1: 2: 1). The amounts of doxorubicin and its metabolites were quantified using LC-MS/MS. Briefly, a triple-quadrupole mass spectrometer equipped with an electrospray ionization (ESI) ion source (LCMS-8030; Shimadzu Corporation) was used in the positive-ESI and multiple reaction monitoring modes. The samples were resolved on an ACQUITY UPLC BEH C18 column (100 × 2.1 mm i.d., 1.7 μm particle) using water and acetonitrile as mobile phases A and B, respectively, at a flow rate of 0.15 ml min−1 and a column temperature of 40 °C. Ion transitions from m/z 544 to m/z 130 and from m/z 546 to m/z 399 for doxorubicin and doxorubicinol, respectively, were monitored for their quantification. Xenograft implantation of HCT116 cells All the protocols for animal experiments in this study were approved by the Experimental Animal Committee of Keio University School of Medicine (the approved number; 08037-(7)). A model of liver metastases of human colon cancer was prepared according to our previous methods with minor modifications. Briefly, HCT116 cells transfected with the cDNA of Venus (1 × 106 cells/mice), a highly sensitive fluorescent protein on tissue slice sections, were transplanted into the spleens of 10-week-old male NOG mice. Ten days after transplantation, the animals were anesthetized with sevoflurane (Maruishi Pharmaceutical), and their livers were removed, embedded in super-cryoembedding medium and frozen quickly in liquid nitrogen. The frozen tissues were sliced with a cryostat (Leica CM1900) at a thickness of 5 μm, and haematoxylin-eosin staining was performed. Fluorescent microscopy (Keyence: BIOREVO, BZ-9000) was used to observe Venus fluorescence. Percentages of cross-sectional areas showing metastatic tumours were calculated by Image-J as previously described. Additional information Accession code: Structural information on PGRMC1 is available from the Protein Data Bank under accession code 4X8Y. How to cite this article: Kabe, Y. et al. Haem-dependent dimerization of PGRMC1/sigma-2 receptor facilitates cancer proliferation and chemoresistance. Nat. Commun. 7:11030 doi: 10.1038/ncomms11030 (2016). Supplementary Material Iron, hemochromatosis, and hepatocellular carcinoma Meat and heme iron intake and esophageal adenocarcinoma in the European prospective investigation into cancer and nutrition study Heme iron from meat and risk of colorectal cancer: a meta-analysis and a review of the mechanisms involved Iron metabolism regulates p53 signaling through direct heme-p53 interaction and modulation of p53 localization, stability and function Enhanced heme function and mitochondrial respiration promote the progression of lung cancer cells Carbon monoxide expedites metabolic exhaustion to inhibit tumor growth Reduced methylation of PFKFB3 in cancer cells shunts glucose towards the pentose phosphate pathway Development and application of high-performance affinity beads: toward chemical biology and drug discovery Membrane-bound progesterone receptors contain a cytochrome b5-like ligand-binding domain Possible role of PGRMC1 in breast cancer development PGRMC1: a new biomarker for the estrogen receptor in breast cancer Regulation of ovarian cancer cell viability and sensitivity to cisplatin by progesterone receptor membrane component-1 Progesterone receptor membrane component 1 deficiency attenuates growth while promoting chemosensitivity of human endometrial xenograft tumors Predictive toxicogenomics approaches reveal underlying molecular mechanisms of nongenotoxic carcinogenicity Elevated progesterone receptor membrane component 1/sigma-2 receptor levels in lung tumors and plasma from lung cancer patients Therapeutic targeting of pancreatic cancer utilizing sigma-2 ligands Overexpression of the cytochrome p450 activator hpr6 (heme-1 domain protein/human progesterone receptor) in tumors Pgrmc1 (progesterone receptor membrane component 1) associates with epidermal growth factor receptor and regulates erlotinib sensitivity Dap1/PGRMC1 binds and regulates cytochrome P450 enzymes Progesterone receptor membrane component 1 (Pgrmc1): a heme-1 domain protein that promotes tumorigenesis and is inhibited by a small molecule S2R(Pgrmc1): the cytochrome-related sigma-2 receptor that regulates lipid and drug metabolism and hormone signaling Spectroscopic and mutagenesis studies of human PGRMC1 Molecular identification of adrenal inner zone antigen as a heme-binding protein Inference of macromolecular assemblies from crystalline state TALOS+: a hybrid method for predicting protein backbone torsion angles from NMR chemical shifts Structural basis of CpG and inhibitory DNA recognition by Toll-like receptor 9 Structural basis for semaphorin signalling through the plexin receptor Involvement of heme regulatory motif in heme-mediated ubiquitination and degradation of IRP2 Replacement of the proximal histidine iron ligand by a cysteine or tyrosine converts heme oxygenase to an oxidase Semi-quantitative analyses of metabolic systems of human colon cancer metastatic xenografts in livers of superimmunodeficient NOG mice Energy management by enhanced glycolysis in G1-phase in human colon cancer cells in vitro and in vivo In vivo antitumor activity and host toxicity of methoxymorpholinyl doxorubicin: role of cytochrome P450 3A Cytochrome P450 CYP1B1 protein expression: a novel mechanism of anticancer drug resistance Progesterone receptor membrane component 1 inhibits the activity of drug-metabolizing cytochromes P450 and binds to cytochrome P450 reductase Involvement of NADPH in the interaction between heme oxygenase-1 and cytochrome P450 reductase Oxidized human neuroglobin acts as a heterotrimeric Galpha protein guanine nucleotide dissociation inhibitor Ligand binding reveals a role for heme in translationally-controlled tumor protein dimerization Structural basis for multimeric heme complexation through a specific protein-heme interaction: the case of the third neat domain of IsdH from Staphylococcus aureus Progesterone receptor membrane component 1: an integrative review Progesterone receptor membrane component-1 (PGRMC1) and PGRMC-2 interact to suppress entry into the cell cycle in spontaneously immortalized rat granulosa cells Regulation of iron homeostasis mediated by the heme-binding protein Dap1 (damage resistance protein 1) via the P450 protein Erg11/Cyp51 Progesterone receptor membrane component 1 modulates human cytochrome P450 activities in an isoform-dependent manner PGRMC1 contributes to doxorubicin-induced chemoresistance in MES-SA uterine sarcoma Involvement of let-7/miR-98 microRNAs in the regulation of progesterone receptor membrane component 1 expression in ovarian cancer cells Reversal of paclitaxel resistance in epithelial ovarian carcinoma cells by a MUC1 aptamer-let-7i chimera Interactions of multiple gas-transducing systems: hallmarks and uncertainties of CO, NO, and H2S gas biology Carbon monoxide from heme catabolism protects against hepatobiliary dysfunction in endotoxin-treated rat liver The heme oxygenase-carbon monoxide system: a regulator of hepatobiliary function Cellular and mitochondrial iron homeostasis in vertebrates Haem oxygenase is synthetically lethal with the tumour suppressor fumarate hydratase The induction of heme oxygenase-1 suppresses heat shock protein 90 and the proliferation of human breast cancer cells through its byproduct carbon monoxide Identification of the PGRMC1 protein complex as the putative sigma-2 receptor binding site Alzheimer's therapeutics targeting amyloid beta 1-42 oligomers II: Sigma-2/PGRMC1 receptors mediate Abeta 42 oligomer binding and synaptotoxicity Progesterone attenuates Abeta25-35-induced neuronal toxicity via JNK inactivation and progesterone receptor membrane component 1-dependent inhibition of mitochondrial apoptotic pathway Sigma-2 receptor binding is decreased in female, but not male, APP/PS1 mice XDS Decision-making in structure solution using Bayesian estimates of map quality: the PHENIX AutoSol wizard Coot: model-building tools for molecular graphics PHENIX: a comprehensive Python-based system for macromolecular structure solution Size-distribution analysis of macromolecules by sedimentation velocity ultracentrifugation and lamm equation modeling The implementation of SOMO (SOlution MOdeller) in the UltraScan analytical ultracentrifugation data analysis suite: enhanced capabilities allow the reliable hydrodynamic modeling of virtually any kind of biomacromolecule An improved diffusion-ordered spectroscopy experiment incorporating bipolar-gradient pulses The use of sample rotation for minimizing convection effects in self-diffusion NMR measurements Carbon monoxide: an endogenous modulator of sinusoidal tone in the perfused rat liver Distribution of heme oxygenase isoforms in rat liver. Topographic basis for carbon monoxide-mediated microvascular relaxation A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications Author contributions Y.K., T.K. and M.S. conceived the project; Y.K., T.N. and M.S. designed experiments and wrote the manuscript; Y.K., T.N. and T.S. performed X-ray crystallography. I.K., T.Y. and Y.S. performed cell biology experiments. E.H. and K.S. contributed to NMR experiments. E.K., M.N. and S.U. performed ESI-MS and SV-AUC analyses. M.O. and K.M. performed xenograft experiments in vivo. A.Y., T.U. and K.I. contributed to Raman spectrometric analyses. Y.Y., H.H. and S.I. contributed to consultations based on their expertise. X-ray crystal structure of PGRMC1. (a) Structure of the PGRMC1 dimer formed through stacked haems. Two PGRMC1 subunits (blue and green ribbons) dimerize via stacking of the haem molecules. (b) Haem coordination of PGRMC1 with Tyr113. Comparison of PGRMC1 (blue) and cytochrome b5 (yellow, ID: 3NER). (c) PGRMC1 has a longer helix (a.a.147–163), which is shifted away from the haem (arrow). PGRCM1 is dimerized by binding with haem. (a) Mass spectrometric analyses of the wild-type (wt) PGRMC1 or the C129S mutant in the presence or absence of haem under non-denaturing condition. Both proteins had identical lengths (a.a.44–195). Data of experimental mass show mean±s.d. (b) SV-AUC analyses of the wt-PGRMC1 and the C129S mutant (a.a.44–195) in the presence or absence of haem. SV-AUC experiments were performed with 1.5 mg ml−1 of PGRMC1 proteins. The major peak with sedimentation coefficient S20,w of 1.9∼2.0 S (monomer) or 3.1 S (dimer) was detected. (c) Difference absorption spectra of PGRMC1 (a.a.44–195) titrated with haem (left panel). The titration curve of haem to PGRMC1 (right panel). The absorbance difference at 400 nm is plotted against the haem concentration. Carbon monoxide inhibits haem-dependent PGRMC1 dimerization. (a) UV-visible absorption spectra of PGRMC1 (a.a.44–195). Measurements were performed in the presence of the oxidized form of haem (ferric), the reduced form of haem (ferrous) and the reduced form of haem plus CO gas (ferrous+CO). (b) Close-up view of haem stacking. Spheres are drawn with van der Waals radii. (c) Gel-filtration chromatography analyses of PGRMC1 (a.a.44–195) wild-type (wt) and the Y113F or C129S mutant in the presence or absence of haem, dithionite and/or CO. (d) Transition model for structural regulation of PGRMC1 in response to haem and CO. Haem-dependent dimerization of PGRMC1 is necessary for tumour proliferation mediated by EGFR signalling. (a) FLAG-PGRMC1 wild-type (wt) and Y113F mutant proteins (a.a.44–195), in either apo- or haem-bound form, were incubated with purified EGFR and co-immunoprecipitated with anti-FLAG antibody-conjugated beads. Input and bound proteins were detected by Western blotting. (b) In vitro binding assay was performed as in (a) using haem-bound FLAG-PGRMC1 wt (a.a.44–195) and purified EGFR with or without treatment of RuCl3 and CORM3. (c) FLAG-PGRMC1 wt or Y113F (full length) was over-expressed in HCT116 cells and immunoprecipitated with anti-FLAG antibody-conjugated beads. Co-immunoprecipitated proteins (FLAG-PGRMC1, endogenous PGRMC1 and EGFR) were detected with Western blotting by using anti-PGRMC1 or anti-EGFR antibody. (d) HCT116 cells were treated with or without 250 μmol l−1 of succinylacetone (SA) for 48 h. The intracellular haem was extracted and quantified by reverse-phase HPLC. The data represent mean±s.d. of four separate experiments. **P\u003c0.01 using unpaired Student's t-test. (e) Co-immunoprecipitation assay was performed as in (c) with or without SA treatment in HCT116 cells. (f) HCT116 cells expressing control shRNA or those knocking down PGRMC1 (PGRMC1-KD) were treated with EGF or left untreated, and components of the EGFR signaling pathway were detected by Western blotting. (g,h) HCT116 cells were treated with or without EGF, SA, RuCl3 and CORM3 as indicated, and components of the EGFR signaling pathway were detected by Western blotting. Haem-dependent dimerization of PGRMC1 accelerates tumour growth through the EGFR signaling pathway. (a) Nucleotide sequences of PGRMC1 targeted by shRNA and of the shRNA-resistant full length PGRMC1 expression vector. Stable PGRMC1-knockdown (PGRMC1-KD) HCT116 cells were transiently transfected with the shRNA-resistant expression vector of wild-type PGRMC1 (wt) or the Y113F mutant (Y113F). (b) Erlotinib was added to HCT116 (control) cells, PGRMC1-KD cells or PGRMC1-KD cells expressing shRNA-resistant PGRMC1 wt or Y113F, and cell viability was examined by MTT assay. The data represent mean±s.d. of four separate experiments. *P\u003c0.01 using ANOVA with Fischer's LSD test. (c) Spheroid formation in control and PGRMC1-KD HCT116 cells. Pictures indicate representative micrographs of spheroids formed under each condition. Spheroid size were measured and compared among groups. The graph represents mean±s.e. of each spheroid size. *P\u003c0.01 using ANOVA with Fischer's LSD test. Scale bar: 0.1 mm. (d) Tumour-bearing livers of NOG mice at 10 days after intrasplenic injection of HCT116 (control) or PGRMC1-KD cells. Percentages of cross-sectional areas showing metastatic tumours were calculated. Data represent mean±s.d. of 10 separate experiments. *P\u003c0.05 using unpaired Student's t-test. Scale bar: 5 mm. Haem-dependent PGRMC1 dimerization enhances tumour chemoresistance through interaction with cytochromes P450. (a,b) FLAG-PGRMC1 wild-type (wt) and Y113F mutant proteins (a.a.44–195), in either apo or haem-bound form, were incubated with CYP1A2 (a) or CYP3A4 (b) and immunoprecipitated with anti-FLAG antibody-conjugated beads. Input and bound proteins were detected by Western blotting. (c) Binding assay was performed as in (a) using haem-bound FLAG-PGRMC1 wt and CYP1A2 with or without RuCl3 and CORM3. (d) Schematic illustration of doxorubicin metabolism is shown on the left. Doxorubicin was incubated with HCT116 cells expressing control shRNA or shPGRMC1 (PGRMC1-KD), and the doxorubicinol/doxorubicin ratios in cell pellets were determined using LC-MS. Data represent mean±s.d. of four separate experiments. **P\u003c0.01 versus control using unpaired Student's t-test. (e) Indicated amounts of doxorubicin were added to HCT116 (control) cells, PGRMC1-KD cells, or PGRMC1-KD cells expressing shRNA-resistant full-length PGRMC1 wt or Y113F, and cell viability was examined by MTT assay. Data represent mean±s.d. of four separate experiments. *P\u003c0.01 using ANOVA with Fischer's LSD test. Schematic diagram for the regulation of PGRMC1 functions. Apo-PGRMC1 exists as an inactive monomer. On binding to haem, PGRMC1 forms a dimer through stacking interactions between the haem moieties, which enables PGRMC1 to interact with EGFR and cytochromes P450, leading to an enhanced proliferation and chemoresistance of cancer cells. CO interferes with the stacking interactions of the haems and thereby inhibits PGRMC1 functions. Data collection and refinement statistics.  \tNative\tPhasing\t \tData collection\t \t Space group\tI4122\tI4122\t \t Cell dimensions\t \t  a, b, c (Å)\t167.23, 167.23, 63.46\t168.11, 168.11, 63.65\t \t  α, β, γ (°)\t90, 90, 90\t90, 90, 90\t \t Wavelength (Å)\t1.000\t1.738\t \t Resolution (Å)\t20.0–1.95 (2.00–1.95)\t20.0–2.50 (2.56–2.50)\t \t Rmeas\t0.067 (1.168)\t0.010 (0.850)\t \t I/σI\t22.83 (2.39)\t22.43 (4.54)\t \t Completeness (%)\t97.8 (99.0)\t99.1 (97.6)\t \t Multiplicity\t11.2 (13.8)\t14.9 (15.2)\t \t CC1/2\t100 (81.8)\t99.9 (93.5)\t \t \t \t \t \tRefinement\t \t Resolution (Å)\t19.72–1.95\t \t \t Number of reflections\t32,298 (2,384)\t \t \t Rwork/Rfree\t0.1834/0.2123\t \t \t Number of atoms\t \t  Protein\t1,776\t \t \t  Ligand/ion\t86\t \t \t  Water\t109\t \t \t B-factors\t \t  Protein\t54.6\t \t \t  Ligand/ion\t42.9\t \t \t  Water\t46.6\t \t \t R.M.S deviations\t \t  Bond lengths (Å)\t0.008\t \t \t  Bond angles (°)\t1.164\t \t \t *Highest resolution shell is shown in parenthesis. PGRMC1 proteins exhibit haem-dependent dimerization in solution.  \tApo form\tHaem-bound form\t \t \t \tMass (Da)\t \tMass (Da)\t \taPGRMC1 wt (a.a.44–195)\t \t ESI-MS\t—\t17,844.14\t—\t36,920.19\t \t Theoretical\t \t17,843.65\t \t36,918.06\t \t \tHydrodynamic radius 10−9 (m)\tMW (kDa)\tHydrodynamic radius 10−9 (m)\tMW (kDa)\t \t DOSY\t2.04–2.15\t20\t2.94–3.02\t42\t \t \tS20,w (S)\tMW (kDa)\tS20,w (S)\tMW (kDa)\t \t SV-AUC\t1.9\t17.6\t3.1\t35.5\t \t \t \t \t \t \t \tbPGRMC1 C129S (a.a.44–195)\t \t ESI-MS\t—\t17,827.91\t—\t36,887.07\t \t Theoretical\t \t17,827.59\t \t36,885.6\t \t \tS20,w (S)\tMW (kDa)\tS20,w (S)\tMW (kDa)\t \t SV-AUC\t2.0\t18.1\t3.1\t35.8\t \t Differences in molecular weights of the wild-type (wt; a) and the C129S mutant (b) PGRMC1 proteins in the absence (apo form) or the presence of haem (haem-bound form). The protein sizes of the wt and C129S PGRMC1 cytosolic domains (a.a.44–195) in the presence or absence of haem were estimated by ESI-MS, DOSY and 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diff --git a/annotated_BioC_JSON/PMC4820378_ann.json b/annotated_BioC_JSON/PMC4820378_ann.json
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index 0000000000000000000000000000000000000000..71ca7d395623afbd70d43e80a716f345392afbd6
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+++ b/annotated_BioC_JSON/PMC4820378_ann.json
@@ -0,0 +1 @@
+[{"sourceid":"4820378","sourcedb":"","project":"","target":"","text":"Template-dependent nucleotide addition in the reverse (3′-5′) direction by Thg1-like protein Structures of Thg1-like proteins provide insight into the template-dependent nucleotide addition in the reverse (3′-5′) direction. Thg1-like protein (TLP) catalyzes the addition of a nucleotide to the 5′-end of truncated transfer RNA (tRNA) species in a Watson-Crick template–dependent manner. The reaction proceeds in two steps: the activation of the 5′-end by adenosine 5′-triphosphate (ATP)/guanosine 5′-triphosphate (GTP), followed by nucleotide addition. Structural analyses of the TLP and its reaction intermediates have revealed the atomic detail of the template-dependent elongation reaction in the 3′-5′ direction. The enzyme creates two substrate binding sites for the first- and second-step reactions in the vicinity of one reaction center consisting of two Mg2+ ions, and the two reactions are executed at the same reaction center in a stepwise fashion. When the incoming nucleotide is bound to the second binding site with Watson-Crick hydrogen bonds, the 3′-OH of the incoming nucleotide and the 5′-triphosphate of the tRNA are moved to the reaction center where the first reaction has occurred. That the 3′-5′ elongation enzyme performs this elaborate two-step reaction in one catalytic center suggests that these two reactions have been inseparable throughout the process of protein evolution. Although TLP and Thg1 have similar tetrameric organization, the tRNA binding mode of TLP is different from that of Thg1, a tRNAHis-specific G−1 addition enzyme. Each tRNAHis binds to three of the four Thg1 tetramer subunits, whereas in TLP, tRNA only binds to a dimer interface and the elongation reaction is terminated by measuring the accepter stem length through the flexible β-hairpin. Furthermore, mutational analyses show that tRNAHis is bound to TLP in a similar manner as Thg1, thus indicating that TLP has a dual binding mode. INTRODUCTION All polynucleotide chain elongation reactions, whether with DNA or RNA, proceed in the 5′-3′ direction. This reaction involves the nucleophilic attack of a 3′-OH of the terminal nucleotide in the elongating chain on the α-phosphate of an incoming nucleotide. The energy in the high-energy bond of the incoming nucleotide is used for its addition [termed tail polymerization ]. This elongation reaction of DNA/RNA chains is in clear contrast to the elongation of protein chains in which the high energy of the incoming aminoacyl-tRNA is not used for its own addition but for the addition of the next monomer (termed head polymerization). However, recent studies have shown that the Thg1/Thg1-like protein (TLP) family of proteins extends tRNA nucleotide chains in the reverse (3′-5′) direction. In this case, the 5′-end of tRNA is first activated using adenosine 5′-triphosphate (ATP)/guanosine 5′-triphosphate (GTP), followed by nucleophilic attack of a 3′-OH on the incoming nucleotide [nucleoside 5′-triphosphate (NTP)] to yield pppN-tRNA. Thus, the energy in the triphosphate bond of the incoming nucleotide is not used for its own addition but is reserved for subsequent polymerization (that is, head polymerization) (Fig. 1). Reaction schemes of 3′-5′ and 5′-3′ elongation. Top: Reaction scheme of 3′-5′ elongation by Thg1/TLP family proteins. Bottom: Reaction scheme of 5′-3′ elongation by DNA/RNA polymerases. In 3′-5′ elongation by Thg1/TLP family proteins, the 5′-monophosphate of the tRNA is first activated by ATP/GTP, followed by the actual elongation reaction. The energy of the incoming nucleotide is not used for its own addition but is reserved for the subsequent addition (head polymerization). In 5′-3′ elongation by DNA/RNA polymerases, the energy of the incoming nucleotide is used for its own addition (tail polymerization). The best-characterized member of this family of proteins is eukaryotic Thg1 (tRNAHis guanylyltransferase), which catalyzes the nontemplated addition of a guanylate to the 5′-end of immature tRNAHis. This guanosine at position −1 (G−1) of tRNAHis is a critical identity element for recognition by the histidyl-tRNA synthase. Therefore, Thg1 is essential to the fidelity of protein synthesis in eukaryotes. However, Thg1 homologs or TLPs are found in organisms in which G−1 is genetically encoded, and thus, posttranscriptional modification is not required. This finding suggests that TLPs may have potential functions other than tRNAHis maturation. TLPs have been shown to catalyze 5′-end nucleotide addition to truncated tRNA species in vitro in a Watson-Crick template–dependent manner. This function of TLPs is not limited to tRNAHis but occurs efficiently with other tRNAs. Furthermore, the yeast homolog (Thg1p) has been shown to interact with the replication origin recognition complex for DNA replication, and the plant homolog (ICA1) was identified as a protein affecting the capacity to repair DNA damage. These observations suggest that TLPs may have more general functions in DNA/RNA repair. The 3′-5′ addition reaction catalyzed by Thg1 occurs through three reaction steps. In the first step, the 5′-monophosphorylated tRNAHis, which is cleaved by ribonuclease P from pre-tRNAHis, is activated by ATP, creating a 5′-adenylylated tRNAHis intermediate. In the second step, the 3′-OH of the incoming GTP attacks the activated intermediate, yielding pppG−1-tRNAHis. Finally, the pyrophosphate is removed, and mature pG−1-tRNAHis is created. The crystal structure of human Thg1 (HsThg1) shows that its catalytic core shares structural homology with canonical 5′-3′ nucleotide polymerases, such as T7 DNA/RNA polymerases. This finding suggests that 3′-5′ elongation enzymes are related to 5′-3′ polymerases and raises important questions on why 5′-3′ polymerases predominate in nature. The crystal structure of TLP from Bacillus thuringiensis shows that it shares a similar tetrameric assembly and active-site architecture with HsThg1. Furthermore, the structure of Candida albicans Thg1 (CaThg1) complexed with tRNAHis reveals that the tRNA substrate accesses the reaction center from a direction opposite to that of canonical DNA/RNA polymerase. However, in this structural analysis, the 5′-end of tRNAHis was not activated and the second substrate (GTP) was not bound. Thus, a detailed reaction mechanism remains unknown. Here, we successfully solved the structure of TLP from the methanogenic archaeon Methanosarcina acetivorans (MaTLP) in complex with ppptRNAPheΔ1, which mimics the activated intermediate of the repair substrate. Although TLP and Thg1 have similar tetrameric organization, the mode of tRNA binding is different in TLP. Furthermore, we obtained the structure in which the GTP analog (GDPNP) was inserted into this complex to form a Watson-Crick base pair with C72 at the 3′-end region of the tRNA. On the basis of these structures, we discuss the reaction mechanism of template-dependent reverse (3′-5′) polymerization in comparison with canonical 5′-3′ polymerization. RESULTS Anticodon-independent binding of ppptRNAPheΔ1 to MaTLP Previous biochemical experiments have suggested that ppptRNAPheΔ1, in which the 5′-end of tRNAPhe was triphosphorylated and G1 was deleted, can be an efficient substrate for the repair reaction (guanylyl transfer) of Thg1/TLP. Therefore, we prepared a crystal of MaTLP complexed with ppptRNAPheΔ1 and solved its structure to study the template-directed 3′-5′ elongation reaction by TLP (fig. S1). The crystal contained a dimer of TLP (A and B molecules) and one tRNA in an asymmetric unit. Two dimers in the crystal further assembled as a dimer of dimers by the crystallographic twofold axis (Fig. 2). This tetrameric structure and 4:2 stoichiometry of the TLP-tRNA complex are the same as those of the CaThg1-tRNA complex. However, whereas the AB and CD dimers of tetrameric CaThg1 play different roles, respectively recognizing the accepter stem and anticodon of tRNAHis, the AB dimer and its symmetry mate (CD dimer) on tetrameric MaTLP independently bind one molecule of tRNA (fig. S2), recognizing the tRNA accepter stem and elbow region. Thus, consistent with the notion that MaTLP is an anticodon-independent repair enzyme, the anticodon was not recognized in the MaTLP-tRNA complex, whereas the binding mode of CaThg1 is for the G−1 addition reaction, therefore the His anticodon has to be recognized (see “Dual binding mode for tRNA repair”). Structure of the MaTLP complex with ppptRNAPheΔ1. Left: One molecule of the tRNA substrate (ppptRNAPheΔ1) is bound to the MaTLP dimer. The AB and CD dimers are further dimerized by the crystallographic twofold axis to form a tetrameric structure (dimer of dimers). Right: Left figure rotated by 90o. The CD dimer is omitted for clarity. The accepter stem of the tRNA is recognized by molecule A (yellow), and the elbow region by molecule B (blue). Residues important for binding are depicted in stick form. The β-hairpin region of molecule B is shown in red. The elbow region of the tRNA substrate was recognized by the β-hairpin of molecule B of MaTLP. The N atoms in the side chain of R215 in the β-hairpin region of MaTLP were hydrogen-bonded to the phosphate groups of U55 and G57. The O atom on the S213 side chain was also hydrogen-bonded to the phosphate moiety of G57 of the tRNA (Fig. 2). This β-hairpin region was disordered in the crystal structure of the CaThg1-tRNA complex. The accepter stem of the tRNA substrate was recognized by molecule A of MaTLP. The N7 atom of G2 at the 5′-end was hydrogen-bonded to the N atom of the R136 side chain, whereas the α-phosphate was bonded to the N137 side chain (Fig. 2). R136 was also hydrogen-bonded to the base of C72 (the Watson-Crick bond partner of ΔG1). The triphosphate moiety at the 5′-end of the tRNA was bonded to the D21-K26 region. These phosphates were also coordinated to two metal ions, presumably Mg2+ (Mg2+A and Mg2+B) because they were observed at the same positions (figs. S3 and S4) previously identified by CaThg1 and HsThg1 structures. These ions were in turn coordinated by the O atoms of the side chains of D21 and D69 and the main-chain O of G22 (fig. S3A). Mutation of D29 and D76 in HsThg1 (corresponding to D21 and D69 of MaTLP) has been shown to markedly decrease G−1 addition activity. Template-dependent binding of the GTP analog to the MaTLP-ppptRNAPheΔ1 complex Here, we successfully obtained the structure of the ternary complex of MaTLP, 5′-activated tRNA (ppptRNAPheΔ1), and the GTP analog (GDPNP) (Fig. 3 and fig. S4) by soaking the MaTLP-ppptRNAPheΔ1 complex crystal in a solution containing GDPNP. The obtained structure showed that the guanine base of the incoming GDPNP formed Watson-Crick hydrogen bonds with C72 and accompanied base-stacking interactions with G2 of the tRNA (Fig. 3B), whereas no interaction was observed between the guanine base and the enzyme. These features are consistent with the fact that this elongation reaction is template-dependent. The 5′-end (position 2) of the tRNA moved significantly (Fig. 3C) due to the insertion of GDPNP. Surprisingly, the 5′-triphosphate moiety after movement occupied the GTP/ATP triphosphate position during the activation step (Fig. 3D). Together with the observation that the 3′-OH of the incoming GTP analog was within coordination distance (2.8 Å) to Mg2+A (fig. S3B) and was able to execute a nucleophilic attack on the α-phosphate of the 5′-end, this structure indicates that the elongation reaction (second reaction) takes place at the same reaction center where the activation reaction (first reaction) occurs. Structural change of the tRNA (ppptRNAPheΔ1). Structural change of the tRNA (ppptRNAPheΔ1) accepter stem in MaTLP caused by insertion of GDPNP. (A) Structure before GDPNP binding. (B) Structure after GDPNP binding. (C) Superposition of the two structures showing movement of the 5′-end of the tRNA before (blue) and after (red) insertion of GDPNP. (D) Superposition of the 5′-end of the tRNA after GDPNP insertion (red) with GTP at the activation step (green), showing that both triphosphate moieties superpose well. The triphosphate moiety of GDPNP was at the interface between molecules A and B and was recognized by the side chains of both molecules, including R19 (molecule A), R83 (molecule B), K86 (molecule B), and R114 (molecule A) (Fig. 3B). All of these residues are well conserved (fig. S5), and mutation of corresponding residues in ScThg1 (R27, R93, K96, and R133) decreased the catalytic efficiency of G−1 addition. The triphosphate of the GDPNP was also bonded to the third Mg2+ (Mg2+C), which, unlike Mg2+A and Mg2+B, is not coordinated by the TLP molecule (fig. S3B). This triphosphate binding mode is the same as that for the second nucleotide binding site in Thg1. However, in previous analyses, the base moiety at the second site was either invisible or far beyond the reaction distance of the phosphate, and therefore, flipping of the base was expected to occur. tRNA binding and repair experiments of the β-hairpin mutants To confirm tRNA recognition by the β-hairpin, we created mutation variants with altered residues in the β-hairpin region. Then, tRNA binding and enzymatic activities were measured. β-Hairpin deletion variant delR198-R215 almost completely abolished the binding of tRNAPheΔ1 (fig. S6). Furthermore, the enzymatic activities of delR198-R215 and delG202-E210 were very weak (5.2 and 13.5%, respectively) compared with wild type, whereas mutations (N179A and F174A/N179A/R188A) on the anticodon recognition site [deduced from the Thg1-tRNAHis complex structure ] had no effect on the catalytic activity (Fig. 4A). Experiments using the tRNAHisΔ1 substrate gave similar results (Fig. 4A). All these results are consistent with the crystal structure and suggest that the β-hairpin plays an important role in anticodon-independent binding of the tRNA substrate. Residues in the β-hairpin are not well conserved, except for R215 (fig. S5). Mutants R215A and R215A/S213A, in which the completely conserved R215 was changed to alanine, showed a moderate effect on the activity (27.3 and 16.3%, respectively). Thus, specific interactions with the conserved R215 and van der Waals contacts to residues in the β-hairpin would be important for tRNA recognition. Mutational analysis of the β-hairpin and anticodon binding region. The rates of guanylylation by various mutants were measured. Error bars represent the SD of three independent experiments. (A) Guanylylation of ppptRNAPheΔ1 and ppptRNAHisΔ1 by various TLP mutants. The activity using [α-32P]GTP, wild-type MaTLP, and ppptRNAPheΔ1 is denoted as 100. (B) Guanylylation of tRNAPheΔ1, tRNAPhe, and tRNAHisΔ−1 by various TLP mutants. The activity to tRNAPheΔ1 is about 10% of ppptRNAPheΔ1. Termination of the elongation reaction by measuring the accepter stem TLPs catalyze the Watson-Crick template–dependent elongation or repair reaction for 5′-end truncated tRNAPhe substrates lacking G1 only (tRNAPheΔ1), or lacking both G1 and G2 (tRNAPheΔ1,2), whereas they do not show any activity with intact tRNAPhe (thus, repair is unnecessary). How TLP distinguishes between tRNAs that need 5′-end repair from ones that do not, or in other words, how the elongation reaction is properly terminated, remains unknown. The present structure of the MaTLP-ppptRNAPheΔ1 complex shows that, unlike Thg1, the TLP dimer binds one molecule of tRNA by recognizing the elbow region by the β-hairpin of molecule B and the 5′-end by molecule A. Therefore, we speculated that the flexible nature of the β-hairpin enables the recognition of tRNA substrates with different accepter stem lengths. To confirm this speculation, we used computer graphics to examine whether the β-hairpin region was able to bind tRNA substrates with different accepter stem lengths when the 5′-end was properly placed in the reaction site. When the 5′-end was placed in the reaction site, the body of the tRNA molecule shifted in a manner dependent on the accepter stem length. The tRNA body also rotated because of the helical nature of the accepter stem (fig. S7). This model structure showed that the accepter stem of intact tRNAPhe was too long for the β-hairpin to recognize its elbow region, whereas tRNAPheΔ1 and tRNAPheΔ1,2 were recognized by the β-hairpin region (fig. S7), which is consistent with previous experiments. On the basis of these model structures, we concluded that the TLP molecule can properly terminate elongation by measuring the accepter stem length of tRNA substrates. Dual binding mode for tRNA repair The present structural analysis revealed that although TLP and Thg1 have a similar tetrameric architecture, they have different binding modes for tRNAs: Thg1 is bound to tRNAHis as a tetramer, whereas TLP is bound to tRNAPhe as a dimer. This difference in the tRNA binding modes is closely related to their enzymatic functions. The tRNAHis-specific G−1 addition enzyme Thg1 needs to recognize both the accepter stem and anticodon of tRNAHis. The tetrameric architecture of the Thg1 molecule allows it to access both regions located at the opposite side of the tRNA molecule [the AB dimer recognizes the accepter stem and CD dimer anticodon ]. In contrast, the binding mode of TLP corresponds to the anticodon-independent repair reactions of 5′-truncated general tRNAs. This binding mode is also suitable for the correct termination of the elongation or repair reaction by measuring the length of the accepter stem by the flexible β-hairpin. Because tRNAHis requires an extra guanosine (G−1) at the 5′-end, the repair enzyme has to extend the 5′-end by one more nucleotide than other tRNAs. TLP has been shown to confer such catalytic activity on tRNAHisΔ−1 (Fig. 4B). Here, we showed that the TLP mutants, wherein the β-hairpin is truncated and tRNAPheΔ1 binding ability is lost, can still bind to tRNAPhe (GUG) whose anticodon is changed to that for His (fig. S6, C, H, and I). Also, the intact tRNAPhe, which is not recognized by TLP (Fig. 4B and fig. S6E), can be recognized when its anticodon is changed to that for His (fig. S6D). Furthermore, the TLP variant (F174A/N179A/R188A) whose anticodon recognition site [deduced from the Thg1-tRNAHis complex structure ] is disrupted has been shown to have a reduced catalytic activity to tRNAHisΔ−1 (Fig. 4B). All these experimental results indicate that TLP recognizes and binds tRNAs carrying the His anticodon in the same way that Thg1 recognizes tRNAHis. Thus, we concluded that TLP has two tRNA binding modes that are selectively used, depending on both the length of the accepter stem and the anticodon. The elongation or repair reaction normally terminates when the 5′-end reaches position 1, but when the His anticodon is present, TLP binds the tRNA in the second mode by recognizing the anticodon to execute the G−1 addition reaction. By having two different binding modes, TLP can manage this special feature of tRNAHis. DISCUSSION The Thg1/TLP family of proteins extends tRNA chains in the 3′-5′ direction. The reaction involves two steps. First, the 5′-phosphate is activated by GTP/ATP. Then, the activated phosphate is attacked by the incoming nucleotide, resulting in an extension by one nucleotide at the 5′-end. Here, we successfully solved for the first time the intermediate structures of the template-dependent 3′-5′ elongation complex of MaTLP. On the basis of these structures, we will discuss the 3′-5′ addition reaction compared with canonical 5′-3′ elongation by DNA/RNA polymerases. Figure 5 is a schematic diagram of the 3′-5′ addition reaction of TLP. This enzyme has two triphosphate binding sites and one reaction center at the position overlapping these two binding sites (Fig. 5A). In the first activation step, when GTP/ATP is bound to site 1 (Fig. 5B), the 5′-phosphate of the tRNA is deprotonated by Mg2+A and attacks the α-phosphate of the GTP/ATP, resulting in an activated intermediate (Fig. 5C). The structure of the MaTLP-ppptRNAPheΔ1 complex, wherein β- and γ-phosphates coordinate with Mg2+A and Mg2+B, respectively (Figs. 3A and 5C′), may represent this activated intermediate. Subsequent binding of an incoming nucleotide to site 2 followed by formation of the Watson-Crick base pair with a nucleotide in the template strand conveys the 3′-OH of the incoming nucleotide to the position of deprotonation by Mg2+A and the 5′-triphosphate of the tRNA to the reaction center (Figs. 3B and 5D). Then, the elongation reaction of step 2 occurs (Fig. 5E). Thus, the present structure shows that this 3′-5′ elongation enzyme utilizes a reaction center homologous to that of 5′-3′ elongation enzymes for both activation and elongation in a stepwise fashion. Although these two reactions are similar in chemistry, their substrate characteristics are very different. It should be noted that TLP has evolved to allow the occurrence of these two elaborate reaction steps within one reaction center. Schematic representation of the 3′-5′ elongation mechanism. (A) The reaction center overlapped with two triphosphate binding sites. A, B, and C (in green) represent binding sites for Mg2+A, Mg2+B, and Mg2+C. P (in blue) represents the phosphate binding sites; O− (in red) is the binding site for the deprotonated OH group. Important TLP residues for tRNA and Mg2+ binding are also shown. (B) Structure of the activation complex (corresponding to fig. S8). GTP/ATP binds to triphosphate binding site 1; the deprotonated OH group of the 5′-phosphate attacks the α-phosphate of GTP/ATP, and PPi (inorganic pyrophosphate) is released. (C) Possible structure after the activation step as suggested from the structure of (C′). (C′) Structure before the elongation reaction (corresponding to Fig. 3A). The 5′-triphosphate of the tRNA binds to the same site as for activation of the 5′-terminus of the tRNA in (B). (D) Structure of initiation of the elongation reaction (corresponding to Fig. 3B). The base of the incoming GTP forms a Watson-Crick hydrogen bond with the nucleotide at position 72 in the template chain and a base-stacking interaction with a neighboring base (G2). Movement of the 5′-terminal chain leaves the 5′-triphosphate of the tRNA in the same site as the activation step in (B). The 3′-OH of the incoming GTP is deprotonated by Mg2+A and attacks the α-phosphate to form a covalent bond. (E) After the elongation reaction, the triphosphate of the new nucleotide is placed on site 1, as in (C′), and is ready for the next reaction. Figure 6 compares the 3′-5′ and 5′-3′ elongation mechanisms, showing the symmetrical nature of both elongation reactions using a similar reaction center composed of Mg2+A and Mg2+B in the conserved catalytic core. In TLP, which carries out 3′-5′ elongation, the 3′-OH of the incoming nucleotide attacks the 5′-activated phosphate of the tRNA to form a phosphodiester bond, whereas in the T7 RNA polymerase, a representative 5′-3′ DNA/RNA polymerase, the 3′-OH of the 3′-terminal nucleotide of the RNA attacks the activated phosphate of the incoming nucleotide to form a phosphodiester bond. In these reactions, the roles of the two Mg ions are identical. Mg2+A activates the 3′-OH of the incoming nucleotide in TLP and the 3′-OH of the 3′-end of the RNA chain in T7 RNA polymerase. The role of Mg2+B is to position the 5′-triphosphate of the tRNA in TLP and the incoming nucleotide in T7 RNA polymerase. These two Mg2+ ions are coordinated by a conserved Asp (D21 and D69 in TLP) in the conserved catalytic core. Structures of template-dependent nucleotide elongation in the 3′-5′ and 5′-3′ directions. Symmetrical relationship between 3′-5′ elongation by TLP (this study) (left) and 5′-3′ elongation by T7 RNA polymerase [Protein Data Bank (PDB) ID: 1S76] (right). Red arrows represent elongation directions. In the 3′-5′ elongation reaction, the 3′-OH of the incoming nucleotide attacks the 5′-activated phosphate of the tRNA to form a phosphodiester bond, whereas in the 5′-3′ elongation reaction, the 3′-OH of the 3′-terminal nucleotide of the RNA attacks the activated phosphate of the incoming nucleotide to form a phosphodiester bond. Green spheres represent Mg2+ ions. Because the chemical roles of tRNA and the incoming nucleotide are reversed in these two reactions, these two substrates are inserted into a similar reaction center from opposite directions (Fig. 6). In spite of this difference, their fundamental reaction scheme is conserved. However, from an energetic viewpoint, these two reactions are clearly different: Whereas the high energy of the incoming nucleotide is used for its own addition in DNA/RNA polymerases, the high energy of the incoming nucleotide is used for subsequent addition in TLP. For this reason, TLP requires a mechanism that activates the 5′-terminus of the tRNA during the initial step of the reaction. Our analysis showed that the initial activation and subsequent elongation reactions occur sequentially at one reaction center. In this case, the enzyme needs to create two substrate binding sites for two different reactions in the vicinities of one reaction center. TLP has successfully created such sites by utilizing a conformational change in the tRNA through Watson-Crick base pairing (Fig. 3). These structural features of the TLP molecule suggest that development of an activation reaction site is a prerequisite for developing the 3′-5′ elongation enzyme. This is clearly more difficult than developing the 5′-3′ elongation enzyme, wherein the activation reaction site is not necessary, and which may be the primary reason why the 5′-3′ elongation enzyme has been exclusively developed. Here, we established a structural basis for 3′-5′ nucleotide elongation and showed that TLP has evolved to acquire a two-step Watson-Crick template–dependent 3′-5′ elongation reaction using the catalytic center homologous to 5′-3′ elongation enzymes. The active site of this enzyme is created at the dimerization interface. The dimerization also endows this protein with the ability to measure the length of the accepter stem of the tRNA substrate, so that the enzyme can properly terminate the elongation reaction. Furthermore, the dual binding mode of this protein suggests that it has further evolved to cover G−1 addition of tRNAHis by additional dimerization (dimer of dimers). Thus, the present structural analysis is consistent with the scenario in which TLP began as a 5′-end repair enzyme and evolved into a tRNAHis-specific G−1 addition enzyme. The detailed molecular mechanism of the Thg1/TLP family established by our analysis will open up new perspectives in our understanding of 3′-5′ versus 5′-3′ polymerization and the molecular evolution of template-dependent polymerases. MATERIALS AND METHODS Plasmid construction Genomic DNA from M. acetivorans NBRC100939 was obtained from the NITE Biological Resource Center. The MaTLP gene was amplified by polymerase chain reaction from genomic DNA. The DNA fragment encoding MaTLP was then cloned between the Nde I and Xho I restriction sites in a pET26b vector with a C-terminal His tag. In the MaTLP gene, the amber stop codon (UAG) at position 142 was translated as Pyl. To express the full-length MaTLP in Escherichia coli, the TAG codon was altered to TGG (encoding Trp) with the QuikChange Site-Directed Mutagenesis Kit (Agilent Technologies) as previously described. The inserted sequence was verified by DNA sequencing. Preparation of MaTLP and mutants Plasmids were transformed into E. coli strain BL21 (DE3) pLysSRARE by electroporation, and cells were grown in LB medium containing kanamycin (25 μg/ml) and chloramphenicol (34 μg/ml) at 37°C until reaching an optical density at 600 nm (OD600) of 0.45. The cells were then induced by the addition of isopropyl-β-d-thiogalactopyranoside to a final concentration of 250 μM and shifted to 18°C for approximately 20 hours before harvest. The cells were harvested and resuspended in buffer A [50 mM Hepes-NaOH (pH 7.5), 1 M NaCl, 4 mM MgCl2, 10% glycerol, 0.5 mM β-mercaptoethanol, lysozyme (0.5 mg/ml), and deoxyibonuclease (0.1 mg/ml)]. After sonication and centrifugation, the His6-tagged protein was purified by immobilized metal-ion affinity chromatography using a HisTrap HP column (GE Healthcare). The sample was washed with 75 mM imidazole and eluted with a 75 to 400 mM imidazole gradient in buffer B [50 mM tris-HCl (pH 7.5), 500 mM NaCl, 4 mM MgCl2, 20% glycerol, and 0.5 mM β-mercaptoethanol]. Then, the collected fractions were diluted in 300 mM NaCl with buffer C [25 mM tris-HCl (pH 7.5), 10% glycerol, 5 mM MgCl2, and 1 mM dithiothreitol (DTT)] and further purified on a HiTrap Heparin HP column (GE Healthcare) by elution with a 300 to 1000 mM NaCl gradient in buffer C. Finally, the protein was loaded onto a HiLoad 16/60 Superdex 200 prep grade column (GE Healthcare) equilibrated with buffer D [20 mM Hepes-NaOH (pH 7.5), 500 mM NaCl, 5 mM MgCl2, 10% glycerol, and 1 mM DTT]. The protein was concentrated to 3.9 mg/ml by ultrafiltration. All MaTLP mutants were constructed with the QuikChange Site-Directed Mutagenesis Kit. MaTLP mutants were purified by a HisTrap HP column for RNA binding assay and further purified by a HiLoad 16/60 Superdex 200 prep grade column for 3′-5′ nucleotide addition assay. Preparation of tRNA and its mutants tRNA transcripts derived from yeast tRNAPhe and tRNAHis were prepared using T7 RNA polymerase as previously described. ppptRNA transcripts were prepared by excluding guanosine 5′-monophosphate (GMP) from the reaction mixture. Transcribed tRNAs were purified by a HiTrap DEAE FF column (GE Healthcare) as previously described. Pooled tRNAs were precipitated with isopropanol and dissolved in buffer E [20 mM Hepes-NaOH (pH 7.5), 100 mM NaCl, and 10 mM MgCl2]. Preparation of the MaTLP-ppptRNAPheΔ1 complex MaTLP and ppptRNAPheΔ1 (tRNAPhe with a triphosphorylated 5′-end and deleted G1) were mixed in a molar ratio of 1.7:1 and incubated for 30 min at room temperature. The mixture was then loaded onto a HiLoad 16/60 Superdex 200 prep grade column equilibrated with buffer F [20 mM Hepes-NaOH (pH 7.5), 400 mM NaCl, 5 mM MgCl2, 10% glycerol, and 1 mM DTT]. Fractions containing the MaTLP-ppptRNAPheΔ1 complex were mixed with 1 mM spermine and concentrated to an OD280 of 16 by ultrafiltration. Crystallization and data collection All crystallization experiments were performed with the sitting-drop vapor diffusion method at 293 K. Initial crystals of MaTLP were obtained by mixing 1 μl of protein solution (3.9 mg/ml) with 1 μl of a reservoir solution containing 0.1 M Hepes-NaOH buffer (pH 7.5), 0.2 M magnesium chloride, and 30% polyethylene glycol 400 (PEG 400). MaTLP-GTP complex crystals were obtained by soaking the MaTLP crystals in the above reservoir solution supplemented with 1 mM GTP overnight. High-resolution crystals of MaTLP in apo form (MaTLP-apo) were obtained unexpectedly by mixing MaTLP with tRNAHis in 0.1 M sodium/potassium phosphate (pH 6.2) containing 2.5 M NaCl. Crystals of the MaTLP-ppptRNAPheΔ1 complex were obtained from a solution containing 0.2 M tripotassium citrate, 0.1 M tris (pH 8.0), 37% PEG3350, and 10 mM praseodymium (III) acetate. Crystals of the MaTLP-ppptRNAPheΔ1-GDPNP complex were obtained by soaking MaTLP-ppptRNAPheΔ1 complex crystals in a reservoir solution containing 0.2 M tripotassium citrate, 0.1 M tris (pH 8.0), 30% PEG3350, 5% glycerol, and 15 mM GDPNP overnight. Crystals of MaTLP-apo and MaTLP-GTP were cryoprotected with a reservoir solution containing 50% PEG400 before flash-cooling, whereas crystals of the MaTLP-ppptRNAPheΔ1-GDPNP and MaTLP-ppptRNAPheΔ1 complexes were flash-cooled without any cryoprotectant under a stream of liquid nitrogen at 100 K. X-ray diffraction data were collected from beamline BL41XU at SPring-8 (Hyogo, Japan) and beamlines BL5A and BL17A at Photon Factory (Ibaraki, Japan). All diffraction data were indexed, integrated, scaled, and merged using XDS. Structure determination and refinement The crystal structure of MaTLP-apo was determined by the molecular replacement (MR) method with Molrep, using the protomer structure of CaThg1 (PDB ID: 3WBZ) as a search model. The protomer structure of MaTLP-apo was then used as a search model to solve the structures of MaTLP-GTP. The crystal structure of the MaTLP-ppptRNAPheΔ1 complex was determined by the MR method with PHASER, using the protomer structures of MaTLP-apo and tRNAPhe from Saccharomyces cerevisiae (PDB ID: 1EHZ) as search models. The structure of the MaTLP-ppptRNAPheΔ1 complex was then used as a search model to solve the MaTLP-ppptRNAPheΔ1-GDPNP complex structure. Initial protein models were fitted manually using Coot, and tRNA models were automatically rebuilt by LAFIRE_NAFIT; these models were then refined using phenix.refine. The data collection and refinement statistics are summarized in Table 1. All structure figures were generated by PyMol. Summary of data collection and refinement statistics. Values in parentheses are for the highest-resolution shell. PF, Photon Factory; Rmsd, root-mean-square deviation. \tMaTLP-apo\tMaTLP-GTP\tMaTLP-ppptRNAPheΔ1\tMaTLP-ppptRNAPheΔ1-GDPNP\t \tPDB ID\t5AXK\t5AXL\t5AXM\t5AXN\t \tData collection\t \t  Beamline\tSPring-8 BL41XU\tSPring-8 BL41XU\tPF BL17A\tPF BL5A\t \t  Space group\tC2221\tC2221\tI222\tI222\t \t  Unit cell parameters a, b, c (Å)\t98.3, 120.5, 157.4\t103.1, 115.7, 144.9\t75.3, 127.6, 143.8\t82.3, 134.1, 147.4\t \t  Wavelength (Å)\t0.9780\t1.0000\t0.97319\t1.0000\t \t  Resolution range (Å)\t50.0–2.29 (2.43–2.29)\t50.0–2.99 (3.17–2.99)\t50.0–2.21 (2.34–2.21)\t50.0–2.70 (2.87–2.70)\t \t  Rmeas (%)*\t8.9 (76.3)\t15.2 (90.0)\t9.7 (74.4)\t11.0 (87.2)\t \t  CC1/2 (%)\t99.8 (80.4)\t99.5 (81.2)\t99.9 (83.6)\t99.9 (83.5)\t \t  〈I/σ(I)〉\t14.7 (2.8)\t12.0 (2.6)\t19.4 (3.2)\t16.9 (2.5)\t \t  Completeness (%)\t98.3 (93.8)\t98.8 (93.4)\t99.7 (98.6)\t99.7 (99.3)\t \t  Redundancy\t6.7 (6.6)\t7.2 (7.2)\t7.4 (7.3)\t8.1 (8.2)\t \tRefinement\t \t  No. of reflections\t41,650\t17,581\t35,102\t22,669\t \t  Rwork/Rfree (%)†\t20.6/24.0\t21.5/25.3\t21.6/24.3\t22.5/26.7\t \t  No. of atoms\t \t    Macromolecules\t3760\t3622\t5247\t5142\t \t    Ligand/ion\t30\t68\t36\t101\t \t    Water\t89\t8\t102\t16\t \t  B-factors (Å2)\t \t    Macromolecules\t57.0\t68.4\t45.3\t57.3\t \t    Ligand/ion\t60.5\t86.2\t46.6\t59.9\t \t    Water\t49.0\t59.5\t33.0\t38.1\t \t  Estimated coordinate error (Å)\t0.32\t0.48\t0.25\t0.41\t \t  Rmsd from ideal\t \t    Bond lengths (Å)\t0.009\t0.003\t0.003\t0.003\t \t    Bond angles (°)\t1.11\t0.92\t0.72\t0.80\t \t *Rmeas = Σhkl {N(hkl)/[N(hkl) − 1]}1/2 Σi | Ii(hkl) − 〈I(hkl)〉 |/Σhkl Σi\nIi(hkl), where 〈I(hkl)〉 and N(hkl) are the mean intensity of a set of equivalent reflections and the multiplicity, respectively. †Rwork\n= Σhkl ||Fobs| − |Fcalc||/Σhkl |Fobs|; Rfree was calculated for 5% randomly selected test sets that were not used in the refinement. Nucleotide addition assay Nucleotide addition assays were performed as previously described. A reaction mixture containing 25 mM Hepes-NaOH (pH 7.5), 400 mM NaCl, 10 mM MgCl2, 3 mM DTT, 5% glycerol, 0.1 μM [α-32P]GTP, 100 μM GTP, 1 μM MaTLP variants, and 10 μM tRNA transcript was incubated at 30°C for 2 hours. Then, the reaction was quenched with phenol/chloroform, and the supernatant was resolved on a 10% polyacrylamide gel containing 8 M urea. The radioactivity was visualized with a BAS-1800 II bioimaging analyzer (Fujifilm). tRNA binding assay A reaction mixture containing 34 μM MaTLP variants and 20 μM tRNA transcript was incubated in buffer F at room temperature for 30 min. Then, the mixture was loaded onto a Superdex 200 10/300 GL column (GE Healthcare) equilibrated with the same buffer. Supplementary Material SUPPLEMENTARY MATERIALS REFERENCES AND NOTES B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, P. Walter, Molecular Biology of the Cell (Garland Science, New York, ed. 5, 2008). tRNAHis-guanylyltransferase establishes tRNAHis identity Doing it in reverse: 3′-to-5′ polymerization by the Thg1 superfamily tRNAHis maturation: An essential yeast protein catalyzes addition of a guanine nucleotide to the 5′ end of tRNAHis Post-transcriptional nucleotide addition is responsible for the formation of the 5′ terminus of histidine tRNA Role of the extra G-C pair at the end of the acceptor stem of tRNAHb in aminoacylation Histidylation by yeast HisRS of tRNA or tRNA-like structure relies on residues −1 and 73 but is dependent on the RNA context Recognition of G-1:C73 atomic groups by Escherichia coli histidyl-tRNA synthetase Template-dependent 3′–5′ nucleotide addition is a shared feature of tRNAHis guanylyltransferase enzymes from multiple domains of life 3′–5′ tRNAHis guanylyltransferase in bacteria tRNA 5′-end repair activities of tRNAHis guanylyltransferase (Thg1)-like proteins from Bacteria and Archaea The highly conserved tRNAHis guanylyltransferase Thg1p interacts with the origin recognition complex and is required for the G2/M phase transition in the yeast Saccharomyces cerevisiae Natural variation identifies ICARUS1, a universal gene required for cell proliferation and growth at high temperatures in Arabidopsis thaliana Histidine tRNA guanylyltransferase from Saccharomyces cerevisiae. II. Catalytic mechanism tRNAHis guanylyltransferase (THG1), a unique 3′-5′ nucleotidyl transferase, shares unexpected structural homology with canonical 5′-3′ DNA polymerases Crystal structure of a bacteriophage T7 DNA replication complex at 2.2 Å resolution Structure of T7 RNA polymerase complexed to the transcriptional inhibitor T7 lysozyme Structural studies of a bacterial tRNAHis guanylyltransferase (Thg1)-like protein, with nucleotide in the activation and nucleotidyl transfer sites Structural basis of reverse nucleotide polymerization tRNAHis guanylyltransferase adds G-1 to the 5′ end of tRNAHis by recognition of the anticodon, one of several features unexpectedly shared with tRNA synthetases Identification of critical residues for G-1 addition and substrate recognition by tRNAHis guanylyltransferase The appearance of pyrrolysine in tRNAHis guanylyltransferase by neutral evolution Rapid, nondenaturing RNA purification using weak anion-exchange fast performance liquid chromatography XDS Molecular replacement with MOLREP Overview of the CCP4 suite and current developments Phaser crystallographic software. The crystal structure of yeast phenylalanine tRNA at 1.93 Å resolution: A classic structure revisited Coot: Model-building tools for molecular graphics New model-fitting and model-completion programs for automated iterative nucleic acid refinement PHENIX: A comprehensive Python-based system for macromolecular structure solution W. L. DeLano, The PyMOL Molecular Graphics System, Version 1.7.4 (Schrödinger, LLC, 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diff --git a/annotated_BioC_JSON/PMC4822050_ann.json b/annotated_BioC_JSON/PMC4822050_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..3a10d1037b5b4b27b5f0a8cd487ed59559423c8e
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4822050_ann.json
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+[{"sourceid":"4822050","sourcedb":"","project":"","target":"","text":"Hemi-methylated DNA opens a closed conformation of UHRF1 to facilitate its histone recognition UHRF1 is an important epigenetic regulator for maintenance DNA methylation. UHRF1 recognizes hemi-methylated DNA (hm-DNA) and trimethylation of histone H3K9 (H3K9me3), but the regulatory mechanism remains unknown. Here we show that UHRF1 adopts a closed conformation, in which a C-terminal region (Spacer) binds to the tandem Tudor domain (TTD) and inhibits H3K9me3 recognition, whereas the SET-and-RING-associated (SRA) domain binds to the plant homeodomain (PHD) and inhibits H3R2 recognition. Hm-DNA impairs the intramolecular interactions and promotes H3K9me3 recognition by TTD–PHD. The Spacer also facilitates UHRF1–DNMT1 interaction and enhances hm-DNA-binding affinity of the SRA. When TTD–PHD binds to H3K9me3, SRA-Spacer may exist in a dynamic equilibrium: either recognizes hm-DNA or recruits DNMT1 to chromatin. Our study reveals the mechanism for regulation of H3K9me3 and hm-DNA recognition by URHF1.  UHRF1 is involved in the maintenance of DNA methylation, but the regulatory mechanism of this epigenetic regulator is unclear. Here, the authors show that it has a closed conformation and are able to make conclusions about the mechanism of recognition of epigenetic marks. DNA methylation is an important epigenetic modification for gene repression, X-chromosome inactivation, genome imprinting and maintenance of genome stability. Mammalian DNA methylation is established by de novo DNA methyltransferases DNMT3A/3B, and DNA methylation patterns are maintained by maintenance DNA methyltransferase 1 (DNMT1) during DNA replication. Ubiquitin-like, containing PHD and RING fingers domains, 1 (UHRF1, also known as ICBP90 and NP95 in mouse) was shown to be essential for maintenance DNA methylation through recruiting DNMT1 to replication forks in S phase of the cell cycle. UHRF1 is essential for S phase entry and is involved in heterochromatin formation. UHRF1 also plays an important role in promoting proliferation and is shown to be upregulated in a number of cancers, suggesting that UHRF1 may serve as a potential drug target for therapeutic applications. UHRF1 is a multi-domain containing protein connecting histone modification and DNA methylation. As shown in Fig. 1a, UHRF1 is comprised of an N-terminal ubiquitin-like domain, followed by a tandem Tudor domain (TTD containing TTDN and TTDC sub-domains), a plant homeodomain (PHD), a SET-and-RING-associated (SRA) domain, and a C-terminal really interesting new gene (RING) domain. We and other groups demonstrated that the TTD and the PHD coordinately recognize histone H3K9me3, in which residue R2 is recognized by the PHD and tri-methylation of residue K9 (K9me3) is recognized by the TTD. The SRA preferentially binds to hemi-methylated DNA (hm-DNA). Recent studies show that the SRA directly binds to replication focus targeting sequence (RFTS) of DNMT1 (RFTSDNMT1). A spacer region (Fig. 1a, designated Spacer hereafter) connecting the SRA and the RING is rich in basic residues and predicted to be unstructured for unknown function. Recent study shows that phosphatidylinostiol phosphate PI5P binds to the Spacer and induces a conformational change of UHRF1 to allow the TTD to recognize H3K9me3 (ref.). These studies indicate that UHRF1 connects dynamic regulation of DNA methylation and H3K9me3, which are positively correlated in human genome. However, how UHRF1 regulates the recognition of these two repressive epigenetic marks and recruits DNMT1 for chromatin localization remain largely unknown. Here we report that UHRF1 adopts a closed conformation, in which the C-terminal Spacer binds to the TTD and inhibits its recognition of H3K9me3, whereas the SRA binds to the PHD and inhibits its recognition of H3R2 (unmethylated histone H3 at residue R2). Upon binding to hm-DNA, UHRF1 impairs the intramolecular interactions and promotes the H3K9me3 recognition by TTD–PHD, which may further enhance its genomic localization. As a result, UHRF1 is locked in the open conformation by the association of H3K9me3 by TTD–PHD, and thus SRA-Spacer either recognizes hm-DNA or recruits DNMT1 for DNA methylation. Therefore, UHRF1 may engage in a sophisticated regulation for its chromatin localization and recruitment of DNMT1 through a mechanism yet to be fully elucidated. Our study reveals the mechanism for regulation of H3K9me3 and hm-DNA recognition by UHRF1. Results Hm-DNA facilitates histone H3K9me3 recognition by UHRF1 To investigate how UHRF1 coordinates the recognition of H3K9me3 and hm-DNA, we purified recombinant UHRF1 (truncations and mutations) proteins from bacteria. We first performed an in vitro pull-down assay using biotinylated histone H3 peptides and hm-DNA (Supplementary Table 1). As shown in Fig. 1b, hm-DNA largely enhanced the interaction between full-length UHRF1 and unmethylated histone H3 (H3K9me0) or H3K9me3 peptide. Compared with hm-DNA, um-DNA (unmethylated DNA) or fm-DNA (fully methylated DNA) showed marginal effect on facilitating the interaction between UHRF1 and histone peptides, which is consistent with previous studies that UHRF1 prefers hm-DNA for chromatin association (Supplementary Fig. 1a). In contrast, histone peptides showed no enhancement on the interaction between hm-DNA and UHRF1 (Fig. 1c). These results suggest that hm-DNA facilitates histone recognition by UHRF1. Our previous studies show that the PHD recognizes H3K9me0 and the TTD and the PHD together (TTD–PHD) coordinately recognize H3K9me3 (refs.). We noticed that the isolated TTD–PHD showed much higher (∼31-fold) binding affinity to H3K9me3 peptide than full-length UHRF1 (Fig. 1d and Supplementary Table 2), and the isolated PHD showed much higher (∼34-fold) binding affinity to H3K9me0 peptide than full-length UHRF1 (Fig. 1e). The gel filtration analysis showed that UHRF1 is a monomer in solution (Supplementary Fig. 1b), indicating that the intramolecular (not intermolecular) interaction of UHRF1 regulates histone recognition. These results suggest that UHRF1 adopts an unfavourable conformation for histone H3 tails recognition, in which TTD–PHD might be blocked by other regions of UHRF1, and hm-DNA impairs this intramolecular interaction to facilitate its recognition of histone H3 tails. Intramolecular interaction within UHRF1 To test above hypothesis, we performed glutathione S-transferase (GST) pull-down assay using various truncations of UHRF1. Interestingly, the TTD directly bound to SRA-Spacer but not the SRA, suggesting that the Spacer (residues 587–674) is important for the intramolecular interaction (Fig. 2a). The isothermal titration calorimetry (ITC) measurements show that the TTD bound to the Spacer (but not the SRA) in a 1:1 stoichiometry with a binding affinity (KD) of 1.59 μM (Fig. 2b). The presence of the Spacer markedly impaired the interaction between TTD–PHD and H3K9me3 (Fig. 2c). The results indicate that the Spacer directly binds to the TTD and inhibits its interaction with H3K9me3. The GST pull-down assay also shows that the PHD bound to the SRA, which was further confirmed by the ITC measurements (KD=26.7 μM; Fig. 2a,d). Compared with the PHD alone, PHD-SRA showed decreased binding affinity to H3K9me0 peptide by a factor of eight (Fig. 2e). Pre-incubation of the SRA also modestly impaired PHD–H3K9me0 interaction. These results indicate that the SRA directly binds to the PHD and inhibits its binding affinity to H3K9me0. Taken together, UHRF1 seems to adopt a closed form through intramolecular interactions (TTD–Spacer and PHD-SRA), which inhibit histone H3 tail recognition by UHRF1. Overall structure of TTD–Spacer To investigate the intramolecular interaction within UHRF1, we first mapped the minimal regions within the Spacer for the interaction with the TTD (Supplementary Fig. 2a). Internal deletions of the Spacer, including SpacerΔ660–664, SpacerΔ665–669, SpacerΔ670–674 and Spacer642–674, bound to the TTD with comparable binding affinities to that of the Spacer, whereas Spacer587–641 showed no detectable interaction. SpacerΔ642–651, SpacerΔ650–654 and SpacerΔ655–659 also decreased binding affinities, indicating that residues 642–674 are important for TTD–Spacer interaction. We next determined the solution structure of the TTD (residues 134–285) bound to Spacer627–674 by conventional NMR techniques (Supplementary Table 3 and Supplementary Fig. 3a,b). In the complex structure, each Tudor domain adopts a ‘Royal' fold containing a characteristic five-stranded β-sheet and the two Tudor domains tightly pack against each other with a buried area of 573 Å2 (Fig. 3a). The TTD adopts similar fold to that in TTD–PHD–H3K9me3 complex structure (PDB: 4GY5) with a root-mean-square deviation of 1.09 Å for 128 Cα atoms, indicating that the Spacer does not result in obvious conformational change of the TTD (Fig. 3b). The Spacer (residues 643–655 were built in the model) adopts an extended conformation and binds to an acidic groove on the TTD (Fig. 3c). The TTD–Spacer interaction is mediated by a number of hydrogen bonds (Fig. 3d). The side chain of residue K648 forms hydrogen bonds with the carbonyl oxygen atom of D189 and side chain of D190 of the TTD. The side chain of residue R649 packs against an acidic surface mainly formed by residues D142 and E153. Residue S651 forms hydrogen bonds with the main chain of residues G236 and W238. The interaction is further supported by hydrogen bonds formed between residues K650, A652, G653 and G654 of the Spacer and residues N228, G236 and W238 of the TTD, respectively. In support of above structural analyses, mutation D142A/E153A of the TTD abolished its interaction with the Spacer (Fig. 3e). Mutations K648D and S651D of the Spacer decreased their binding affinities to the TTD, and mutation R649A of the Spacer showed more significant decrease (∼13-fold) in the binding affinity (Fig. 3f). As negative control, mutations S639D and S666D of the Spacer showed little effect on the interaction. Interestingly, phosphorylation at residue S651 of UHRF1 was observed in previous mass-spectrometry analyses. Compared with the unmodified peptide of Spacer642–664, a phosphorylation at S651 markedly decreased the binding affinity to the TTD (Supplementary Fig. 2b), suggesting that the phosphorylation may regulate the intramolecular interaction within UHRF1. The spacer binds to the TTD by competing with the linker Previous studies indicate that the TTD binds to a linker region connecting the TTD and PHD (residues 286–306, designated Linker, Fig. 1a), and TTD–Linker interaction is essential for H3K9me3 recognition by TTD–PHD. Comparison of TTD–Spacer and TTD–PHD–H3K9me3 (PDB: 4GY5) structures indicates that the Spacer and the Linker bind to the TTD in a similar manner in the two complexes (Fig. 3b). In TTD–PHD–H3K9me3 structure, residues R295, R296 and S298 of the Linker adopt almost identical conformation to residues K648, R649 and S651 of the Spacer in TTD–Spacer structure, respectively. Similar intramolecular contacts (TTD–Linker and TTD–Spacer) were observed in the two structures (Fig. 3b,d and Supplementary Fig. 4a). Thus, the Spacer may disrupt the TTD–Linker interaction and inhibits the recognition of H3K9me3 by TTD–PHD. To test this hypothesis, we first investigated the potential competition between the Linker and the Spacer for their interaction with the TTD. The ITC experiment shows that the Linker peptide (289–306) bound to the TTD with a binding affinity of 24.04 μM (Supplementary Fig. 4b), ∼15-fold lower than that of the Spacer peptide (KD=1.59 μM, Fig. 3e). The competitive ITC experiments show that TTD–Spacer binding affinity decreased by a factor of two in the presence of the Linker, whereas TTD–Linker interaction was abolished in the presence of the Spacer (Supplementary Fig. 4c). Compared with TTD–Spacer interaction (KD=1.48 μM), TTD–PHD decreased the binding affinity to the Spacer (KD=10.68 μM), whereas mutation R295D/R296D (within the Linker and important for TTD–Linker interaction) of TTD–PHD showed minor decrease in the binding affinity (KD=2.69 μM; Fig. 3g), indicating a competition between the Spacer and the Linker on the same binding site of the TTD. Notably, although the Linker (in the context of TTD-PHD) impairs the TTD–Spacer interaction to some extent, the isolated Spacer could still bind to TTD–PHD with moderate binding affinity (KD=10.68 μM), supporting the existence of the intramolecular interaction within UHRF1. To test whether TTD–Spacer association exists in the context of full-length UHRF1, we used various truncations of UHRF1 in the GST pull-down assay. As indicated in Fig. 3h, full-length UHRF1 and UHRF1ΔSRA showed no interaction with GST-tagged TTD, Linker or Spacer, suggesting that TTD–Spacer interaction in-cis within full-length UHRF1 or UHRF1ΔSRA prohibits TTD–Spacer complex formation in-trans. In contrast, UHRF1ΔTTD bound to GST-TTD, and UHRF1Δ627–674 bound to GST-Spacer, indicating that lack of TTD–Spacer interaction in-cis, TTD–Spacer complex could form in-trans, supporting that the TTD binds to the Spacer in the context of full-length UHRF1. Moreover, GST-Linker showed very weak if not undetectable interaction with wild-type or deletions of UHRF1, suggesting that TTD–Linker interaction is much weaker than that of TTD–Spacer. Taken together, UHRF1 adopts a closed conformation, in which the Spacer binds to the TTD through competing with the Linker, and therefore inhibits H3K9me3 recognition by UHRF1. The spacer inhibits H3K9me3 recognition by the isolated TTD Our previous study indicates that H3K9me3 binds to the TTD in different manner in TTD–PHD–H3K9me3 (ref.) and TTD-H3K9me3 (PDB: 2L3R) structures. Because the TTD is always associated with the PHD, whether the pattern of TTD–H3K9me3 interaction exists in vivo remains unknown. Nevertheless, comparison of TTD–H3K9me3 and TTD–Spacer structures indicates that H3K9me3 and the Spacer overlap on the surface of the TTD (Supplementary Fig. 4d), suggesting that the Spacer might block the H3K9me3 recognition by the isolated TTD. As shown in Supplementary Fig. 4e, the Spacer inhibited TTD–H3K9me3 interaction, whereas its TTD-binding defective mutants of the Spacer or the SRA (a negative control) markedly decreased the inhibition. We next tested whether such inhibition also occurs in the context of full-length UHRF1. Compared with full-length UHRF1, UHRF1Δ627–674 enhanced H3K9me3-binding affinity by a factor of four (Supplementary Fig. 4f). The restoration of H3K9me3-binding affinity is not dramatic because the PHD still binds to histone H3 in both proteins. To exclude this effect, we performed the assay using UHRF1D334A, which abolishes H3R2-binding affinity of the PHD. UHRF1D334A showed undetectable H3K9me3-binding affinity, whereas UHRF1D334A\u0026Δ627–674 dramatically restored its H3K9me3-binding affinity (KD=8.69 μM; Supplementary Fig. 4f), indicating that H3K9me3 recognition by the TTD is blocked by the Spacer through competitive interaction with the TTD. Moreover, the R295D/R296D mutant of full-length UHRF1 showed decreased binding affinity to H3K9me3 (eightfold lower than wild type), suggesting that mutation of R295D/R296D favours TTD–Spacer interaction and therefore promotes UHRF1 to exhibit a more stable closed conformation (Supplementary Fig. 4g). Taken together, the Spacer binds to the TTD and inhibits H3K9me3 recognition by UHRF1 through (i) disrupting TTD–Linker interaction, which is essential for H3K9me3 recognition by TTD–PHD, (ii) prohibiting H3K9me3 binding to the isolated TTD. TTD–PHD–H3K9me3 complex inhibits TTD–spacer interaction Interestingly, pre-incubation of H3K9me3 peptide completely blocked the interaction between the Spacer and the TTD alone or TTD–PHD (Supplementary Fig. 4h), whereas the presence of the Spacer partially impaired the interaction between TTD–PHD and H3K9me3 (Fig. 2c). The results are also consistent with the previous observation that the interaction between TTD–PHD and the Spacer is much weaker (KD=10.68 μM, Fig. 3g) than that between TTD–PHD and H3K9me3 (KD=0.15 μM, Fig. 1d). These results suggest that once TTD–PHD binds to H3K9me3, UHRF1 will be locked by H3K9me3 and the Spacer is unlikely to fold back for the intramolecular interaction. Hm-DNA disrupts intramolecular interaction within UHRF1 To investigate whether hm-DNA could open the closed conformation of UHRF1, we first measured the intramolecular interaction using UHRF1 truncations in the presence or absence of hm-DNA. The GST pull-down assays show that the PHD bound to the SRA and such interaction was impaired by the addition of hm-DNA (Fig. 4a). H3 peptide pull-down assays show that hm-DNA only enhanced the H3K9me0-binding affinities of UHRF1 truncations containing PHD-SRA, such as PHD-SRA, TTD-PHD-SRA, TTD-PHD-SRA-Spacer, UHRF1ΔTTD and UHRF1ΔSpacer (Fig. 4b). The result indicates that hm-DNA disrupts PHD–SRA interaction and facilitates H3K9me0-binding affinity of the PHD in a manner independent on the TTD or the Spacer. Moreover, the TTD or TTD–PHD bound to SRA–Spacer and the interaction was impaired by the addition of hm-DNA (Fig. 4c). The ITC measurements show that the presence of hm-DNA markedly impaired the interaction between the TTD and SRA–Spacer (Supplementary Fig. 5a). However, the TTD–Spacer interaction was not affected by the presence of the hm-DNA, indicating that hm-DNA displaces the Spacer from the TTD in a SRA-dependent manner (Supplementary Fig. 5b). To investigate whether hm-DNA disrupts TTD–Spacer interaction in the context of full-length UHRF1, we monitored the conformational changes of UHRF1 using its histone-binding affinity as read-out. UHRF1D334A was used to exclude the effect of H3K9me0 recognition by the PHD. As expected, all D334A-containing mutants showed undetectable interaction with H3K9me0 (Fig. 4d). UHRF1D334A bound to H3K9me3 peptide in the presence of hm-DNA, but showed no interaction in the absence of hm-DNA, which is consistent with the ITC experiments (Supplementary Fig. 4f). In contrast, UHRF1D334A\u0026Δ627–674 strongly bound to H3K9me3 even in the absence of hm-DNA (Fig. 4d), indicating that the deletion of the Spacer releases otherwise blocked TTD–PHD for H3K9me3 recognition. The results further support the conclusion that the Spacer binds to the TTD in the context of full-length UHRF1 and the intramolecular interactions are disrupted by hm-DNA. We next performed similar peptide pull-down assay using two mutants (N228C/G653C and R235C/G654C) generated on UHRF1D334A. Residues N228/R235 from the TTD and G653/G654 from the Spacer were chosen according to the TTD–Spacer complex structure (Supplementary Fig. 5c) so that the replaced Cysteine residues (one from the TTD and one from the Spacer) are physically close enough to each other to form a disulphide bond in the absence of reducing reagent (dithiothreitol, DTT). As shown in Fig. 4d, hm-DNA largely enhanced the H3K9me3-binding affinities of both mutants in the presence of DTT, but not in the absence of DTT, indicating that the disulphide bond formation (in the absence of DTT) disallows hm-DNA to disrupt TTD–Spacer interaction for H3K9me3 recognition. As negative controls, H3K9me3 recognition by UHRF1D334A or UHRF1D334A\u0026Δ627–674 is not affected by DTT. The above results collectively demonstrate that (i) full-length UHRF1 adopts a closed form, in which the Spacer binds to the TTD and H3K9me3 recognition is inhibited; (ii) hm-DNA displaces the Spacer from the TTD in the context of full-length UHRF1 and therefore largely enhances its histone H3K9me3-binding activity in a manner independent on the PHD (SRA is required). We have previously demonstrated that hm-DNA also disrupts PHD–SRA interaction and facilitates H3K9me0-binding affinity of the PHD in a manner independent on the TTD or the Spacer. Taken together, hm-DNA disrupts the intramolecular interactions within UHRF1, and therefore facilitates the coordinate recognition of H3K9me3 by TTD–PHD. The spacer enhances hm-DNA-binding affinity of the SRA To investigate how hm-DNA impairs TTD–Spacer interaction, we tested whether the Spacer is involved in hm-DNA recognition by the SRA, which is the only known domain for hm-DNA recognition within UHRF1. In the electrophoretic mobility-shift assay, SRA–Spacer showed higher hm-DNA-binding affinity than the SRA alone (Supplementary Fig. 6a). ITC measurements show that SRA–Spacer bound to hm-DNA with a much higher binding affinity (KD=1.75 μM) than the SRA (KD=25.12 μM), whereas the Spacer alone showed no interaction with hm-DNA (Fig. 5a). In the fluorescence polarization (FP) measurements, SRA–Spacer, full-length UHRF1 and UHRF1ΔTTD showed comparable hm-DNA-binding affinities (Fig. 5b and Supplementary Table 4), suggesting that UHRF1 binds to hm-DNA no matter UHRF1 adopts a closed form or not. In contrast, UHRF1ΔSRA abolished hm-DNA-binding affinity, indicating that the SRA is essential for hm-DNA recognition. Compared with full-length UHRF1, UHRF1Δ627–674 decreased the hm-DNA-binding affinity by a factor of 14 (Fig. 5b), further supporting that the Spacer plays an important role in hm-DNA recognition in the context of full-length UHRF1. In addition, hm-DNA-binding affinities of SRA or SRA–Spacer did not obviously vary upon the change of DNA lengths but did decrease with the increasing salt concentrations (Supplementary Fig. 6b,c and Supplementary Table 5). These results indicate that the Spacer not only binds to the TTD and inhibits H3K9me3 recognition when UHRF1 adopts closed conformation, but also facilitates hm-DNA recognition by the SRA when UHRF1 binds to hm-DNA. We next mapped the minimal region of the Spacer for the enhancement of hm-DNA-binding affinity. SRA–Spacer-661 (residues 414–661) still maintained strong hm-DNA-binding affinity comparable to that of SRA–Spacer (residues 414–674), whereas SRA–Spacer-652 and SRA–Spacer-642 markedly decreased their hm-DNA-binding affinities (Fig. 5c), indicating that residues 642–661 are important for enhancing hm-DNA-binding affinity of the SRA. This minimal region largely overlaps with the Spacer region (643–655) essential for TTD interaction. We also determined the crystal structure of SRA–Spacer bound to hm-DNA at 3.15 Å resolution (Supplementary Table 6 and Supplementary Fig. 7a). The structure shows that the SRA binds to hm-DNA in a manner similar to that observed in the previously reported SRA-hm-DNA structures. Intriguingly, no electron density was observed for the Spacer. A possible explanation is that the Spacer facilitates SRA–hm-DNA interaction through nonspecific salt bridge contacts because DNA is rich in acidic groups and the Spacer is rich in basic residues (Supplementary Fig. 7b). The nonspecific interaction is consistent with the previous observation that UHRF1 has no DNA sequence selectivity besides hm-CpG dinucleotide. The spacer is important for PCH localization of UHRF1 To investigate the role of the Spacer in the regulation of UHRF1 function, we transiently overexpressed GFP-tagged wild type or mutants of UHRF1 in NIH3T3 cells to determine their subcellular localization. For the NIH3T3 cells expressing wild-type UHRF1, most cells (∼74.6%) showed a focal pattern of protein that is co-localized with 4,6-diamidino-2-phenylindole (DAPI) foci (Fig. 5d), whereas the rest cells showed a diffuse nuclear staining pattern. The result is consistent with the previous studies that UHRF1 is mainly localized to highly methylated pericentromeric heterochromatin (PCH). In contrast, for the cells expressing UHRF1Δ627–674, a spacer deletion mutant with decreased hm-DNA-binding affinity (Fig. 5b), only ∼22.1% cells showed co-localization with DAPI. Previous reports have shown that the H3K9me3 recognition of UHRF1 also plays an important role in its heterochromatin localization. For example, UHRF1 mutant (within TTD domain) lacking H3K9me3-binding affinity largely reduces its co-localization with heterochromatin. Because manipulation of endogenous hm-DNA in cells is technically challenging, we used UHRF1ΔSRA (lacks hm-DNA-binding affinity but maintains closed conformation, Figs 3h and 5b) to test whether closed conformation of UHRF1 exists in vivo. In NIH3T3 cells, UHRF1ΔSRA largely decreased chromatin association (Fig. 5d). Only ∼4.8% cells expressing UHRF1ΔSRA showed an intermediate enrichment, but not characteristic focal pattern, at DAPI foci, whereas the majority of the cells showed a diffuse nuclear staining pattern. The results suggest that UHRF1ΔSRA adopts closed conformation so that H3K9me3 recognition by TTD–PHD is blocked by the intramolecular interaction, and support the regulatory role of the Spacer in PCH localization of UHRF1 in vivo. The spacer facilitates UHRF1–DNMT1 interaction Previous studies show that UHRF1 recruits DNMT1 to hm-DNA for maintenance DNA methylation through the interaction between the SRA and RFTSDNMT1 (refs). We confirmed the direct interaction between RFTSDNMT1 and the SRA in a solution with low salt concentration (50 mM NaCl), but observed weak or undetectable interaction in a solution with higher salt concentrations (100 or 150 mM NaCl) (Supplementary Fig. 8a). Compared with the SRA, SRA–Spacer exhibited stronger interaction with RFTSDNMT1. In addition, RFTSDNMT1 bound to SRA–Spacer with a binding affinity of 7.09 μM, but showed no detectable interaction with the SRA (Supplementary Fig. 8b). Interestingly, the addition of hm-DNA abolished the interaction between RFTSDNMT1 and SRA–Spacer, suggesting that hm-DNA also regulates UHRF1–DNMT1 interaction (Supplementary Fig. 8c). These results indicate that the Spacer facilitates the interaction between RFTSDNMT1 and the SRA, and the interaction is impaired by the presence of hm-DNA. We next tested whether the UHRF1–DNMT1 interaction is regulated by the conformational change of UHRF1. Because the addition of hm-DNA disrupts the interaction between the SRA–Spacer and RFTSDNMT1, we used various truncations to mimic open and closed forms of UHRF1. In the absence of hm-DNA, only UHRF1ΔTTD bound to RFTSDNMT1, whereas full-length UHRF1, UHRF1ΔSRA and UHRF1Δ627–674 showed undetectable interaction (Fig. 5e). As the deletion of the TTD allows UHRF1 to adopt an open conformation, the results suggest that RFTSDNMT1 binds to SRA–Spacer when UHRF1 adopts an open conformation in the absence of hm-DNA. In support of above observations, the addition of large amount of RFTSDNMT1 impaired the interaction between UHRF1 and hm-DNA (Supplementary Fig. 8d), suggesting an existence of dynamic equilibrium between UHRF1–hm-DNA and UHRF1–DNMT1 complexes. Discussion According to the above results, we here proposed a working model for hm-DNA-mediated regulation of UHRF1 conformation (Fig. 5f). In the absence of hm-DNA (A), UHRF1 prefers a closed conformation, in which the Spacer binds to the TTD by competing with the Linker and the SRA binds to the PHD. As a result, the recognition of histone H3K9me3 by the TTD is blocked by the Spacer, and recognition of unmodified histone H3 (H3R2) by the PHD is inhibited by the SRA. The interaction between UHRF1 and DNMT1 is also weak because the Spacer is unable to facilitate the intermolecular interaction. In the presence of hm-DNA (B), UHRF1 prefers an open conformation, in which the SRA binds to the hm-DNA; the Spacer dissociates from the TTD and facilitates the interaction between the SRA and hm-DNA; the Linker binds to the TTD and allows TTD–PHD to recognize histone H3K9me3. When UHRF1 adopts an open conformation and has already bound to H3K9me3 (B), the interaction between H3K9me3 and TTD–PHD further prevents the Spacer from folding back to interact with the TTD, and therefore locks UHRF1 in an open conformation. The association of UHRF1 to the histone may facilitate the ubiquitination of histone tail (mediated by RING domain) for DNMT1 targeting. Moreover, through a mechanism yet to be fully elucidated, DNMT1 targets hm-DNA for maintenance DNA methylation, probably through interaction with the histone ubiquitylation and/or SRA-Spacer. This cartoon summarizes our findings in this study. The P(r) function obtained from small-angle X-ray scattering (SAXS) measurements of TTD–PHD–SRA–Spacer–hm-DNA complex showed a broader distribution than that of the TTD–PHD–SRA–Spacer alone, supporting the proposed model that UHRF1 adopts an open conformation in the presence of hm-DNA (Supplementary Fig. 8e). Many questions need to be further clarified. We have tried crystallizing more than three sub-constructs with and without DNA across over 1,200 crystallization conditions but failed to determine the structure of TTD–PHD–SRA–Spacer in the absence or presence of hm-DNA. Getting these structures would greatly help for understanding the hm-DNA-mediated regulation of UHRF1. In addition, this regulatory process should be further characterized using advanced techniques, such as single molecular measurement. Our previous studies show that phosphorylation at S639 within the Spacer disrupts interaction between UHRF1 and deubiquitylase USP7 and decreases UHRF1 stability in the M phase of the cell cycle. The Spacer was predicted to contain two nuclear localization signals, residues 581–600 and 648-670 (ref.). In this report, we found that the Spacer (i) binds to the TTD in the closed form of UHRF1 and inhibits its interaction with H3K9me3; (ii) facilitates hm-DNA recognition by the SRA and (iii) facilitates the interaction between the SRA and RFTSDNMT1. These findings together indicate that the Spacer plays a very important role in the dynamic regulation of UHRF1. When our manuscript was in preparation, Gelato et al. reported that binding of PI5P to the Spacer opens the closed conformation of UHRF1 and increases H3K9me3-binding affinity of the TTD. The result suggests that PI5P may facilitate the conformational change of UHRF1 induced by hm-DNA when UHRF1 is recruited to chromatin. In addition, mass-spectrometry analyses have identified several phosphorylation sites (S639, S651, S661) within the Spacer, suggesting that post-translational modification may add another layer of regulation of UHRF1 (refs). It has been well characterized that the SRA of UHRF1 preferentially recognizes hm-DNA through a base-flipping mechanism. Our study demonstrates that the Spacer markedly enhances the hm-DNA-binding affinity of the SRA and the deletion of the Spacer impairs heterochromatin localization of UHRF1, indicating that the Spacer is essential for recognition of hm-DNA in the context of full-length UHRF1. Interestingly, variant in methylation 1 (VIM1, a UHRF1 homologue in Arabidopsis) contains an equivalent spacer region, which was shown to be required for hm-DNA recognition by its SRA domain, suggesting a conserved regulatory mechanism in SRA domain-containing proteins. Intriguingly, UHRF2 (the only mammalian homologue of UHRF1) and UHRF1 show very high sequence similarities for all the domains but very low similarity for the Spacer (Supplementary Fig. 7c). Thus, although UHRF2 exhibits the histone- and hm-DNA-binding activities, the difference in the Spacer region may contribute to the functional differences between UHRF1 and UHRF2. This is also consistent with previous finding that UHRF2 is unable to replace UHRF1 to maintain the DNA methylation. One of the key questions in the field of DNA methylation is why UHRF1 contains modules recognizing two repressive epigenetic marks: H3K9me3 (by TTD–PHD) and hm-DNA (by the SRA). Previous studies show that chromatin localization of UHRF1 is dependent on hm-DNA, whereas other studies indicate that histone H3K9me3 recognition and hm-DNA association are both required for UHRF1-mediated maintenance DNA methylation. However, little is known about the crosstalk between these two epigenetic marks within UHRF1. In this study, we provide an explanation. As shown in the proposed model, recognition of H3K9me3 by full-length UHRF1 is blocked to avoid its miss-localization to unmethylated genomic region, in which chromatin contains H3K9me3 (KD=4.61 μM) or H3K9me0 (KD=25.99 μM). We have shown that full-length UHRF1 and SRA–Spacer strongly bind to hm-DNA (0.35 and 0.49 μM, respectively) and the Spacer plays an important role in PCH localization (Fig. 5d). Therefore, genomic localization of UHRF1 is primarily determined by its recognition of hm-DNA, which allows UHRF1 to adopt an open form and promotes its histone tail recognition for proper genomic localization and function. As a result, when SRA–Spacer dissociates from hm-DNA and binds to DNMT1 with a currently unknown mechanism, UHRF1 may keep the complex associated with chromatin through the interaction between TTD–PHD and H3K9me3 (or PHD-H3), and make it possible for DNMT1 to target proper DNA substrate for methylation. This explanation agrees nicely with previous observations and clarifies the importance of coordinate recognition of H3K9me3 and hm-DNA by UHRF1 for maintenance DNA methylation. UHRF1 is essential for maintenance DNA methylation through recruiting DNMT1 to DNA replication forks during S phase. This function is probably induced by a direct interaction between the SRA and RFTSDNMT1 (refs) or interaction between DNMT1 and ubiquitylation of histione tail. Recent study indicates that histone tail association of UHRF1 (by the PHD domain) is required for histone H3 ubiquitylation, which is dependent on ubiquitin ligase activity of the RING domain of UHRF1 (ref.). DNMT1 binds to ubiquitylated histone H3 and ubiquitylation is required for maintenance of DNA methylation in vivo. In this study, we found that both TTD and PHD are regulated by hm-DNA to recognize histone tail. Thus, the closed form UHRF1 may prevent miss localization of URHF1, whereas only the UHRF1 in open conformation (induced by hm-DNA) could properly binds to histone tail for ubiquitylation and subsequent DNA methylation. Moreover, structural analyses of DNMT1–DNA and SRA–DNA complexes also indicate that it is impossible for DNMT1 to methylate the hm-DNA that UHRF1 binds to because of steric hindrance. In our in vitro assays, we could detect interaction between SRA–Spacer and RFTSDNMT1, but not the interaction between full-length UHRF1 and RFTSDNMT1 (Supplementary Fig. 8a,b and Fig. 5e). The results suggest that UHRF1 adopts multiple conformations. Binding of UHRF1 to hm-DNA may serve as a switch for its recruitment of DNMT1. The S phase-dependent interaction between UHRF1 and DNMT1 (refs) suggest that DNMT1 may also undergo conformation changes so that RFTSDNMT1 binds to UHRF1 and the catalytic domain of DNMT1 binds to hm-DNA for reaction. Methods Protein expression and purification The ubiquitin-like domain (residues 1–133), TTD (residues 134–285), PHD (residues 307–366), TTD–PHD (residues 134–366), SRA (residues 414–617), SRA–Spacer (residues 414–674), Spacer (residues 587–674), RING (residues 675–793), UHRF1 (residues 1–793) and other mutants or truncations of human UHRF1 were sub-cloned in a pGEX-6p-1 derivative vector. The truncated Spacer (residues 627–674) used for NMR analyses was inserted into modified pRSF-Duet-1 vector. All the proteins were expressed in E. coli strain BL21 (DE3) and purified as described previously. In brief, the transformants were grown at 37 °C in 2X YT medium and induced by adding isopropyl-β-D-thiogalactopyranoside (IPTG) to 0.1 mM when the OD600 reached 0.6 and further incubated at 15 °C overnight. The cells were harvested and disrupted. After centrifugation, the supernatant of GST-tagged proteins was purified by GST affinity column (GE Healthcare) and the His-tagged truncated Spacer (residues 627–674) was purified by Nickel Nitrilotriacetic Acid affinity chromatography (GE Healthcare). The GST-tagged proteins used for GST pull-down experiment were eluted directly. The fusion proteins were digested with PreScission protease and further purified by ion exchange and gel filtration chromatography. The proteins were concentrated to 5–20 mg ml−1 for the following biochemical and structural analyses. To purify 15N- and 13C-labelled proteins, the transformants were grown in M9 medium containing 15N-labelled NH4Cl (1 g l−1) and 13C-labelled glucose (2 g l−1). The isotope-labelled TTD and truncated Spacer were purified as described above. Pull-down experiments For GST pull-down assays, 15 μg GST-tagged proteins were incubated with 40 μg recombinant proteins in 500 μl pull-down buffer (20 mM HEPES-NaOH, pH 8.0, 100 mM NaCl, 5% glycerol and 0.1% Triton X-100) for 1 h at 4 °C. Glutathione resins (GE Healthcare) were washed six times with pull-down buffer then mixed with the proteins for 1 h at 4 °C. After washed three times with pull-down buffer, the bound proteins were analysed by SDS–PAGE Coomassie blue staining. For competitive pull-down experiments: purified proteins were pre-incubated with hemi-methylated-DNA (12 bp, upper strand: 5′-GGGCCXGCAGGG-3′, X=5-methyldeoxycytosine) at the indicated molar ratios for 10 min at 4 °C. For salt concentration-dependent pull-down experiments, the pull-down buffer contains 50, 100 or 150 mM NaCl, respectively. For histone peptide or hm-DNA pull-down, 1 μg biotinylated histone H3 peptide (residues 1–21) or hm-DNA (12-bp, upper strand 5′-GAGGCXGCCTGC-3′ and lower strand 5′-biotin-GCAGGCGGCCTC-3′, X=5-methyldeoxycytosine) were incubated with 20 μg wild type or mutants of UHRF1 proteins in 500 μl pull-down buffer (20 mM HEPES-NaOH, pH 8.0, 100 mM NaCl, 5% glycerol and 0.1% Triton X-100) for 1 h at 4 °C. The proteins were pre-incubated with hm-DNA (12- bp, upper strand: 5′-GGGCCXGCAGGG-3′, X=5-methyldeoxycytosine) or indicated H3 peptide, or binding buffer as a control, at 1:2 molar ratios for 10 min at 4 °C. Then, 20 μl streptavidin beads were washed six times with pull-down buffer and incubated with the mixture for 1 h at 4 °C. The bound proteins were analysed as described above. The results are summarized in Supplementary Fig. 10. ITC measurements The binding affinity of protein/protein, protein/peptide or protein/DNA was measured by adding 0.05 mM protein in cell and titrated with 0.5 mM protein, peptide or hm-DNA (12 bp, upper strand: 5′-GGGCCXGCAGGG-3′, X=5-methyldeoxycytosine) in the syringe using iTC200 microcalorimeter (GE Healthcare) at 18 °C. For competition ITC experiments: the indicated proteins were pre-incubated with competitive peptide, protein or hm-DNA (at 1:2 molar ratio if not specified) for 10 min followed by ITC measurements as described above. Proteins, DNA and peptides were prepared within ITC buffer containing 10 mM HEPES, pH 8.0, 100 mM NaCl. The data were fitted by software Origin 7.0. All ITC results were summarized in Supplementary Table 2 and raw data were shown in Supplementary Fig. 9. NMR titration assay To determine the mole ratio of TTD versus Spacer peptide (residues 627–674) in the complex for NMR studies, NMR stepwise titration assay was performed at 20 °C in a PBS buffer supplemented with 0.01% NaN3, pH 7.4 and 10% D2O. The Spacer peptide was added into 15N-labelled TTD solution with an increasing molar ratio of TTD/Spacer as follows: 1:0.0, 1:0.2, 1:0.4, 1:0.6, 1:0.8, 1:1.2 and 1:1.5. The 1H–15N heteronuclear single-quantum correlation (HSQC) spectra of the TTD were collected after each addition. NMR spectroscopy and analysis Two NMR samples were prepared in a mole ratio of 1:1.2 (TTD/Spacer). One is 0.7 mM uniformly 13C/15N-labelled TTD in complex with unlabelled Spacer peptide in NMR buffer (PBS buffer, 0.01% NaN3, pH 7.4 and 10% D2O). The other is 0.5 mM 15N-labelled Spacer peptide mixed with unlabelled TTD protein. All NMR experiments were performed at 20 °C on a Varian Unity Inova 600 NMR spectrometer equipped with a triple resonances cryoprobe and pulsed field gradients. The standard suite of experiments for assigning the 1H, 13C and 15N backbone, determining the side-chain chemical shifts of the TTD in complex with the Spacer peptide and collecting the Nuclear Overhauser effect (NOE)-based distance restraints were measured, including two-dimensional (2D) 13C-edited HSQC and 15N-edited HSQC; three-dimensional (3D) HNCA, HNCO, HN(CO)CA, HNCACB, CBCA(CO)NH, 15N-resolved HSQC-total correlation spectroscopy (TOCSY) and 13C-resolved HSQC-TOCSY in both aliphatic and aromatic regions; 15N-resolved HSQC-NOESY; 13C-resolved HSQC-NOESY for both aliphatic and aromatic resonances and 2D hbcbcgcdceheA and hbcbcgcdhdA spectra for the correlation of Cβ and Hδ or Hɛ in the aromatic ring that is used for aromatic proton assignment. The NMR signals of bound TTD were assigned according to the previously report. The proton NMR signals of the bound Spacer peptide were assigned by analysing the 2D 13C-filtered, 15N-filtered and J-resolved NOE spectroscopy (NOESY) and TOCSY spectra recorded for the 13C- and 15N-labelled protein with the unlabelled Spacer peptide and the 2D 1H–1H COSY, NOESY and TOCSY spectra recorded for the unlabelled free Spacer peptide, and 15N-edited HSQC, 3D 15N-resolved HSQC-TOCSY for the 15N-labelled Spacer in complex with the TTD protein in the NMR buffer described above, respectively. The intermolecular NOEs between the labelled protein and the unlabelled Spacer peptide were obtained by analysing the 3D 13C-F1-edited and 13C/15N-F3-filtered NOESY spectra. The spectra were processed with the NMRPipe programme and analysed using Sparky 3 (http://www.cgl.ucsf.edu/home/sparky/). Determining the NMR structure The calculations were performed using a standard simulated annealing protocol implemented in the XPLOR-2.29 programme (NIH version). The inter-proton distance restraints derived from the NOE intensities were grouped into three distance ranges, namely 1.8–2.9, 1.8–3.5and 1.8–6.0 Å, which corresponds to strong, medium and weak NOEs, respectively. The dihedral angles phi and psi were derived from the backbone chemical shifts (HN, HA, CO and CA) using the programme TALOS. The hydrogen-bond constraints were generated based on the observed NOE pattern between anti-β-sheets in the TTD domain, confirmed by H-D exchange experiments, and used in structural calculation. A total of ten iterations were performed (50 structures in the initial eight iterations). In total, 100 structures were computed during the last two iterations, and the 20 conformers with the lowest energy were used to represent the 3D structures. The conformers of these bundles (TTD in complex with the Spacer peptide) do not violate the following constraints: NOE \u003e0.3 Å and dihedral angle \u003e3o. The entire structure statistics were evaluated with PROCHECK and PROCHECK-NMR and are summarized in Supplementary Table 3. All of the structure figures were generated using the PyMOL and MOLMOL programmes. Electrophoretic mobility-shift assay A 6-carboxy-fluorescein (FAM)-labelled primer, 5′-CCATGCGCTGAC-3′, was annealed to a primer 5′-GTCAGXGCATGG-3′ (X=5-methyldeoxycytosine). The hemi-methylated double-strand DNA was used in both electrophoretic mobility-shift assay and FP assays. 50 nM FAM-hm-DNA (1 pmole per lane) was pre-incubated with indicated amount of proteins in reaction buffer (20 mM HEPES, pH 7.5, 100 mM NaCl, 8% glycerol and 1 mM DTT) for 20 min on ice. The samples were subjected to a 10% polyacrylamide gel electrophoresis and run in 0.5 × Tris-borate-EDTA buffer at 100 V for 1 h at 4 °C. The results were visualized on Tanon-5200 Chemiluminescent Imaging System (Tanon Science \u0026 Technology Co., Ltd). FP measurements The 12-bp FAM-labelled hm-DNA (as described above) was incubated with increasing amount of indicated proteins for 20 min at 25 °C in reaction buffer containing 20 mM HEPES, pH 7.5, 175 mM NaCl, 8% glycerol and 1 mM DTT. FP measurements were performed at 25 °C on Synergy 4 Microplate Reader (BioTek). The 16-bp and 20-bp FAM-labelled hm-DNA (lower strand: 5′-GTGTCAGXGCATGGCC-3′ and 5′-CCGTGTCAGXGCATGGCCAT-3′, respectively. X=5-methyldeoxycytosine) were used in the FP experiment to test the effect of DNA length on the protein/DNA interaction. All experiments were performed in triplicate. The curves were fitted by GraphPad Prism 5. For salt concentration-dependent FP experiments, the reaction buffer contains 50 mM or 150 mM NaCl, respectively. Crystallization and data collection Crystals of SRA–Spacer in complex with an 18-bp hm-DNA (upper strand: 5′-CATCGTCCCTGCGGGCCC-3′, lower strand: 5′-GGGCCXGCAGGGACGATG-3′. X=5-methyldeoxycytosine) were grown at 18 °C using the hanging drop vapour diffusion method by mixing an equal volume of protein–DNA complex and crystallization buffer containing 12% PEG 3350, 45 mM citric acid/55 mM BIS-TRIS propane (pH 6.9). Protein and hm-DNA were mixed at the molar ratio of 1:1.5 and incubated for 0.5 h on ice before crystallization. Crystals were flash frozen in a cold nitrogen stream at −173 °C. All data sets were collected on beamline BL17U at the SSRF (Shanghai Synchrotron Radiation Facility, China). The data were processed using the programme HKL2000 (ref.). Structure determination The structure of SRA–Spacer–hm-DNA complex was determined by molecular replacement using structure of the SRA (PDB:3BI7) as a searching model. Rotation and translation function searches were performed with the programme PHASER. The model was manually built with COOT. All refinements were performed using the refinement module phenix.refine of PHENIX package. The model quality was checked with the PROCHECK programme and all structure figures were generated by PyMol. Cell culture, transient transfection and images capture NIH3T3 cells were obtained from the Shanghai Institute of Biochemistry and Cell Biology. Wild type and mutants of UHRF1 were sub-cloned into pEGFP-C1 vector. Transient transfections of NIH3T3 cells were carried out using Lipofectamine 2000 (Invitrogen). The NIH3T3 cells were grown on glass coverslips and harvested in 36 h after transfection. The images were acquired and examined as previously described. Briefly, cells were fixed with 4% paraformaldehyde for 25 min, then washed with PBS three times. Coverslips were mounted with Antifade reagent containing DAPI (Molecular Probes) on slides and examined with a confocal microscopy. SAXS measurements SAXS measurements were performed with Anton Paar SAXSess mc2 instrument with linecolimation and charge-coupled-device detection. The X-ray wavelength was 1.5418 Å (CuKα), the sample to detector was 306.8 mm and the sample slit width was 10 mm. Each sample was prepared in 300 μl solution contained 150 mM NaCl, 10 mM HEPES, pH=8.0, 5 mM DTT and 5% glycerol. For the hm-DNA-bound form, the protein was pre-incubated with hm-DNA (12-bp, upper strand: 5′-GGGCCmCGCAGGG-3′, mC=5-methyldeoxycytosine) at 1:1.2 molar ratio for 10 min on ice. To correct for interparticle interference, the data of protein sample were collected twice and each time for 1 h. The solution containing no protein sample was also tested as background. The initial data were first processed using SAXSquant and the further analysis with ATSAS software. The SAXS data were only analysed these were collected in the first hour because there was no time effect on the samples. The radius of gyration Rg was estimated from primus. The distance distribution function P(r) was calculated in PCG package. The maximum particle dimension Dmax was estimated from the P(r) function as the r for which P(r)=0. Additional information Accession codes: The coordinate and structure factor for the TTD–Spacer complex structure have been deposited in the Protein Data Bank under accession code 5IAY. The chemical shift assignment of TTD–Spacer was deposited with BMRB ADIT-NMR online deposition system under the accession number 30019. How to cite this article: Fang, J. et al. Hemi-methylated DNA opens a closed conformation of UHRF1 to facilitate its histone recognition. Nat. Commun. 7:11197 doi: 10.1038/ncomms11197 (2016). Supplementary Material Epigenetic regulation of gene expression: how the genome integrates intrinsic and environmental signals DNA methylation: roles in mammalian development Establishing, maintaining and modifying DNA methylation patterns in plants and animals Dynamic regulation of DNA methylation during mammalian development DNA methylation patterns and epigenetic memory Two DNA methyltransferases from murine erythroleukemia cells: purification, sequence specificity, and mode of interaction with DNA DNA methyltransferases Dnmt3a and Dnmt3b are essential for de novo methylation and mammalian development The DNA methyltransferases of mammals UHRF1 plays a role in maintaining DNA methylation in mammalian cells The SRA protein Np95 mediates epigenetic inheritance by recruiting Dnmt1 to methylated DNA Np95 is regulated by E1A during mitotic reactivation of terminally differentiated cells and is essential for S phase entry The retinoblastoma gene and its product are targeted by ICBP90: a key mechanism in the G1/S transition during the cell cycle ICBP90, a novel methyl K9 H3 binding protein linking protein ubiquitination with heterochromatin formation The UHRF family: oncogenes that are drugable targets for cancer therapy in the near future? Critical role of the ubiquitin ligase activity of UHRF1, a nuclear RING finger protein, in tumor cell growth ICBP90 belongs to a new family of proteins with an expression that is deregulated in cancer cells UHRF1 is a novel diagnostic marker of lung cancer Recognition of modification status on a histone H3 tail by linked histone reader modules of the epigenetic regulator UHRF1 Structural insight into coordinated recognition of trimethylated histone H3 lysine 9 (H3K9me3) by the plant homeodomain (PHD) and tandem tudor domain (TTD) of UHRF1 (ubiquitin-like, containing PHD and RING finger domains, 1) protein Crystal structure of PHD domain of UHRF1 and insights into recognition of unmodified histone H3 arginine residue 2 Recognition of multivalent histone states associated with heterochromatin by UHRF1 protein PHD finger recognition of unmodified histone H3R2 links UHRF1 to regulation of euchromatic gene expression Multivalent histone engagement by the linked tandem Tudor and PHD domains of UHRF1 is required for the epigenetic inheritance of DNA methylation Structural basis for site-specific reading of unmodified R2 of histone H3 tail by UHRF1 PHD finger Recognition of hemi-methylated DNA by the SRA protein UHRF1 by a base-flipping mechanism Structural basis for recognition of hemi-methylated DNA by the SRA domain of human UHRF1 The SRA domain of UHRF1 flips 5-methylcytosine out of the DNA helix The interaction of the SRA domain of ICBP90 with a novel domain of DNMT1 is involved in the regulation of VEGF gene expression The UHRF1 protein stimulates the activity and specificity of the maintenance DNA methyltransferase DNMT1 by an allosteric mechanism The DNA methyltransferase Dnmt1 directly interacts with the SET and RING finger-associated (SRA) domain of the multifunctional protein Uhrf1 to facilitate accession of the catalytic center to hemi-methylated DNA Accessibility of different histone H3-binding domains of UHRF1 is allosterically regulated by phosphatidylinositol 5-phosphate System-wide temporal characterization of the proteome and phosphoproteome of human embryonic stem cell differentiation Np95 is implicated in pericentromeric heterochromatin replication and in major satellite silencing UHRF1 targets DNMT1 for DNA methylation through cooperative binding of hemi-methylated DNA and methylated H3K9 Uhrf1-dependent H3K23 ubiquitylation couples maintenance DNA methylation and replication DNA methylation requires a DNMT1 ubiquitin interacting motif (UIM) and histone ubiquitination M phase phosphorylation of the epigenetic regulator UHRF1 regulates its physical association with the deubiquitylase USP7 and stability ICBP90, a novel human CCAAT binding protein, involved in the regulation of topoisomerase IIalpha expression Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during mitosis UHRF1 phosphorylation by cyclin A2/cyclin-dependent kinase 2 is required for zebrafish embryogenesis VIM1, a methylcytosine-binding protein required for centromeric heterochromatinization Cooperative DNA and histone binding by Uhrf2 links the two major repressive epigenetic pathways S phase-dependent interaction with DNMT1 dictates the role of UHRF1 but not UHRF2 in DNA methylation maintenance DNA methylation requires a DNMT1 ubiquitin interacting motif (UIM) and histone ubiquitination Structure-based mechanistic insights into DNMT1-mediated maintenance DNA methylation Structure of DNMT1-DNA complex reveals a role for autoinhibition in maintenance DNA methylation Determining the structures of large proteins and protein complexes by NMR Two-dimensional NMR experiments for correlating carbon-13. beta. and proton. delta./. epsilon. chemical shifts of aromatic residues in 13C-labeled proteins via scalar couplings NMRPipe: a multidimensional spectral processing system based on UNIX pipes Sources of and solutions to problems in the refinement of protein NMR structures against torsion angle potentials of mean force Protein backbone angle restraints from searching a database for chemical shift and sequence homology PROCHECK: a program to check the stereochemical quality of protein structures AQUA and PROCHECK-NMR: programs for checking the quality of protein structures solved by NMR  MOLMOL: a program for display and analysis of macromolecular structures Processing of X-ray diffraction data collected in oscillation mode Likelihood-enhanced fast translation functions Coot: model-building tools for molecular graphics PHENIX: building new software for automated crystallographic structure determination Author contributions J.F., H.Y., C.C. and Y.X. designed the experiments. J.F., J.C., M.L., R.G., X.Z. and Y.F. performed protein purification and crystallization of SRA-Spacer-DNA complex. J.F., J.C. and P.W. collected the data and determined the crystal structure. Jia.W. and W.L. performed the NMR titration assay. Jiaolong W. determined the NMR structure of TTD-Spacer complex. Q.Z. performed all the cell-based assays. J.F., M.L. and J.C. performed protein interaction assays. J.F. and Z.G. performed the SAXS measurements. C.T., W.L., C.C. and Jie.W. helped in the data analysis. J.F., H.Y. and Y.X. analysed the data and wrote the manuscript. Y.X. supervised the project. Hm-DNA facilities histione tails recognition by full-length UHRF1. (a) Colour-coded domain structure of human UHRF1. The boundaries of the domains are indicated with the numbers representing the amino-acid positions. Note that the conserved motif (green background) of the Linker (residues 286–306) and the Spacer (residues 587–674) bind to the TTD in a similar manner (Fig. 3b). (b) Hm-DNA facilities histone H3 and H3K9me3 recognition by UHRF1. Purified full-length UHRF1 was incubated with biotinylated H3 (1–21) or H3K9me3 (1–21) peptides in the presence or absence of hm-DNA (molar ratio UHRF1/hm-DNA=1:2). The mixture was immobilized onto streptavidin Sepharose beads. The bound proteins were analysed in SDS–PAGE followed by Coomassie blue staining. Sequences of the peptides are indicated in Supplementary Table 1. (c) Histone peptides do not affect hm-DNA-binding affinity of UHRF1. Full-length UHRF1 was incubated with biotinylated hm-DNA in the presence or absence of H3 (1–17) or H3K9me3 (1–17) peptides and analysed as in b. (d,e) Superimposed ITC enthalpy plots for binding of H3K9me3 peptide (1–17) to TTD–PHD and full-length UHRF1 (d), and H3 peptide (1–17) to the PHD and full-length UHRF1 (e). The estimated binding affinities (KD) are listed. The samples in the syringe (designated Sy hereafter) and cell are indicated. Intramolecular interactions inhibit histone recognition by UHRF1. (a) GST pull-down assays for the intramolecular interactions. The isolated domains of UHRF1 were incubated with GST-tagged TTD or PHD immobilized on glutathione resin. The bound proteins were analysed by SDS–PAGE and Coomassie blue staining. (b,d) Superimposed ITC enthalpy plots for the intramolecular interactions of isolated UHRF1 domains. The estimated binding affinities (KD) were listed. ND, not detectable. (c) Superimposed ITC enthalpy plots for the binding of H3K9me3 to TTD–PHD in the absence or presence of the Spacer (molar ratio TTD–PHD/Spacer=1:2). (e) Superimposed ITC enthalpy plots for the binding of H3 to PHD–SRA or PHD in the absence or presence of the SRA (molar ratio PHD/SRA=1:1 or 1:2). ND, not determined; Sy., syringe. NMR structure of the TTD bound to the Spacer. (a) Ribbon representation of TTD–Spacer structure. N- and C-termini of the Spacer are indicated. The TTD is coloured in green, and the Spacer is coloured in yellow. The colour scheme is used the all the structure figures. (b) Superimposition of TTD–Spacer and TTD–PHD–H3K9me3 (4GY5.PDB) structures shown in ribbon representations. The TTD is coloured in green and the Spacer in yellow in TTD–Spacer structure. TTD–PHD–H3K9me3 complex is coloured in grey, and the PHD and H3K9me3 are omitted for simplicity. Residues for the interactions are shown in stick representation. (c) Electrostatic potential surface representation of the TTD with the Spacer shown in ribbon representation. The critical residues on the Spacer for the interaction are shown in stick representation. (d) Close-up view of TTD–Spacer interaction. Critical residues for the interaction are shown in stick representation. Hydrogen bonds are indicated as dashed lines. (e–g) Superimposed ITC enthalpy plots for the interaction between the Spacer and the TTD (or TTD–PHD) with the estimated binding affinity (KD) indicated. Wild-type and mutant proteins for the measurements are indicated. (h) GST pull-down assays for the intramolecular interactions. The wild-type or indicated truncations of UHRF1 were incubated with GST-tagged TTD, Linker or Spacer. The mixtures were immobilized on glutathione resin. The bound proteins were analysed by SDS–PAGE and Coomassie blue staining. ND, not determined; Sy., syringe. Hm-DNA impairs the intramolecular interaction of UHRF1 and facilitates its histone recognition. (a) Hm-DNA impairs the intramolecular interaction of PHD–SRA. The SRA was incubated with GST-tagged PHD in the presence of increasing concentrations of hm-DNA and immobilized on glutathione resin. The bound proteins were analysed in SDS–PAGE and Coomassie blue staining (left) and quantified by band densitometry (right). Error bars, s.d. for triplicate experiments. (b) Purified fragments of UHRF1 were analysed by histone peptide (H3K9me0) pull-down assay as described in Fig. 1b. (c) Hm-DNA impairs the intramolecular interaction of TTD–Spacer. SRA–Spacer was incubated with GST-tagged TTD–PHD or TTD in the presence of increasing concentrations of hm-DNA and analysed in pull-down experiment as described in a. The quantified band densitometries are indicated below the Coomassie blue staining. Error bars, s.d. for triplicate experiments. (d) Histone peptide pull-down assay using UHRF1 mutants as indicated. The assays were performed in the presence (+DTT) or absence (−DTT) of 15 mM DTT. The Spacer facilitates hm-DNA–SRA interaction and DNMT1–UHRF1 interaction. (a) Superimposed ITC enthalpy plots for hm-DNA-binding affinities of the SRA, the Spacer and SRA–Spacer. (b,c) Superimposed fluorescence polarization (FP) plots for hm-DNA-binding affinities of truncations or full-length UHRF1. The estimated binding affinities (KD) are listed above. (d) Subcellular localization of GFP-tagged wild-type or indicated mutants of UHRF1 in NIH3T3 cells. The percentages of cells showing co-localization with DAPI foci were counted from at least 100 cells and shown on the left of the corresponding representative confocal microscopy. The experiment was repeated three times in the same condition. Scale bar, 5 μm. (e) GST pull-down experiment for the interactions between wild-type or truncations of UHRF1 and RFTSDNMT1 as described in Fig. 2a. (f) Working model for hm-DNA-mediated conformational changes of UHRF1, as described in the Discussion. ND, not determined; Sy., 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diff --git a/annotated_BioC_JSON/PMC4822561_ann.json b/annotated_BioC_JSON/PMC4822561_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..2fa577db099676e629f865005e7edff4109801d8
--- /dev/null
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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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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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+[{"sourceid":"4831588","sourcedb":"","project":"","target":"","text":"X-ray Crystallographic Structures of a Trimer, Dodecamer, and Annular Pore Formed by an Aβ17–36 β-Hairpin High-resolution structures of oligomers formed by the β-amyloid peptide Aβ are needed to understand the molecular basis of Alzheimer’s disease and develop therapies. This paper presents the X-ray crystallographic structures of oligomers formed by a 20-residue peptide segment derived from Aβ. The development of a peptide in which Aβ17–36 is stabilized as a β-hairpin is described, and the X-ray crystallographic structures of oligomers it forms are reported. Two covalent constraints act in tandem to stabilize the Aβ17–36 peptide in a hairpin conformation: a δ-linked ornithine turn connecting positions 17 and 36 to create a macrocycle and an intramolecular disulfide linkage between positions 24 and 29. An N-methyl group at position 33 blocks uncontrolled aggregation. The peptide readily crystallizes as a folded β-hairpin, which assembles hierarchically in the crystal lattice. Three β-hairpin monomers assemble to form a triangular trimer, four trimers assemble in a tetrahedral arrangement to form a dodecamer, and five dodecamers pack together to form an annular pore. This hierarchical assembly provides a model, in which full-length Aβ transitions from an unfolded monomer to a folded β-hairpin, which assembles to form oligomers that further pack to form an annular pore. This model may provide a better understanding of the molecular basis of Alzheimer’s disease at atomic resolution. Introduction High-resolution structures of oligomers formed by the β-amyloid peptide Aβ are desperately needed to understand the molecular basis of Alzheimer’s disease and ultimately develop preventions or treatments. In Alzheimer’s disease, monomeric Aβ aggregates to form soluble low molecular weight oligomers, such as dimers, trimers, tetramers, hexamers, nonamers, and dodecamers, as well as high molecular weight aggregates, such as annular protofibrils. Over the last two decades the role of Aβ oligomers in the pathophysiology of Alzheimer’s disease has begun to unfold. Mouse models for Alzheimer’s disease have helped shape our current understanding about the Aβ oligomerization that precedes neurodegeneration. Aβ isolated from the brains of young plaque-free Tg2576 mice forms a mixture of low molecular weight oligomers. A 56 kDa soluble oligomer identified by SDS-PAGE was found to be especially important within this mixture. This oligomer was termed Aβ*56 and appears to be a dodecamer of Aβ. Purified Aβ*56 injected intercranially into healthy rats was found to impair memory, providing evidence that this Aβ oligomer may cause memory loss in Alzheimer’s disease. Smaller oligomers with molecular weights consistent with trimers, hexamers, and nonamers were also identified within the mixture of low molecular weight oligomers. Treatment of the mixture of low molecular weight oligomers with hexafluoroisopropanol resulted in the dissociation of the putative dodecamers, nonamers, and hexamers into trimers and monomers, suggesting that trimers may be the building block of the dodecamers, nonamers, and hexamers. Recently, Aβ trimers and Aβ*56 were identified in the brains of cognitively normal humans and were found to increase with age. A type of large oligomers called annular protofibrils (APFs) have also been observed in the brains of transgenic mice and isolated from the brains of Alzheimer’s patients. APFs were first discovered in vitro using chemically synthesized Aβ that aggregated into porelike structures that could be observed by atomic force microscopy (AFM) and transmission electron microscopy (TEM). The sizes of APFs prepared in vitro vary among different studies. Lashuel et al. observed APFs with an outer diameter that ranged from 7–10 nm and an inner diameter that ranged from 1.5–2 nm, consistent with molecular weights of 150–250 kDa. Quist et al. observed APFs with an outer diameter of 16 nm embedded in a lipid bilayer. Kayed et al. observed APFs with an outer diameter that ranged from 8–25 nm, which were composed of small spherical Aβ oligomers, 3–5 nm in diameter. Although the APFs in these studies differ in size, they share a similar annular morphology and appear to be composed of smaller oligomers. APFs have also been observed in the brains of APP23 transgenic mice by immunofluorescence with an anti-APF antibody and were found to accumulate in neuronal processes and synapses. In a subsequent study, APFs were isolated from the brains of Alzheimer’s patients by immunoprecipitation with an anti-APF antibody. These APFs had an outer diameter that ranged from 11–14 nm and an inner diameter that ranged from 2.5–4 nm. Dimers of Aβ have also been isolated from the brains of Alzheimer’s patients.− Aβ dimers inhibit long-term potentiation in mice and promote hyperphosphorylation of the microtubule-associated protein tau, leading to neuritic damage. Aβ dimers have only been isolated from human or transgenic mouse brains that contain the pathognomonic fibrillar Aβ plaques associated with Alzheimer’s disease. Furthermore, the endogenous rise of Aβ dimers in the brains of Tg2576 and J20 transgenic mice coincides with the deposition of Aβ plaques. These observations suggest that the Aβ trimers, hexamers, dodecamers, and related assemblies may be associated with presymptomatic neurodegeneration, while Aβ dimers are more closely associated with fibril formation and plaque deposition during symptomatic Alzheimer’s disease.− The approach of isolating and characterizing Aβ oligomers has not provided any high-resolution structures of Aβ oligomers. Techniques such as SDS-PAGE, TEM, and AFM have only provided information about the molecular weights, sizes, morphologies, and stoichiometry of Aβ oligomers. High-resolution structural studies of Aβ have primarily focused on Aβ fibrils and Aβ monomers. Solid-state NMR spectroscopy studies of Aβ fibrils revealed that Aβ fibrils are generally composed of extended networks of in-register parallel β-sheets.− X-ray crystallographic studies using fragments of Aβ have provided additional information about how Aβ fibrils pack. Solution-phase NMR and solid-state NMR have been used to study the structures of the Aβ monomers within oligomeric assemblies.− A major finding from these studies is that oligomeric assemblies of Aβ are primarily composed of antiparallel β-sheets. Many of these studies have reported the monomer subunit as adopting a β-hairpin conformation, in which the hydrophobic central and C-terminal regions form an antiparallel β-sheet. In 2008, Hoyer et al. reported the NMR structure of an Aβ monomer bound to an artificial binding protein called an affibody (PDB 2OTK). The structure revealed that monomeric Aβ forms a β-hairpin when bound to the affibody. This Aβ β-hairpin encompasses residues 17–37 and contains two β-strands comprising Aβ17–24 and Aβ30–37 connected by an Aβ25–29 loop. Sequestering Aβ within the affibody prevents its fibrilization and reduces its neurotoxicity, providing evidence that the β-hairpin structure may contribute to the ability of Aβ to form neurotoxic oligomers. In a related study, Sandberg et al. constrained Aβ in a β-hairpin conformation by mutating residues A21 and A30 to cysteine and forming an intramolecular disulfide bond. Locking Aβ into a β-hairpin structure resulted in the formation Aβ oligomers, which were observed by size exclusion chromatography (SEC) and SDS-PAGE. The oligomers with a molecular weight of ∼100 kDa that were isolated by SEC were toxic toward neuronally derived SH-SY5Y cells. This study provides evidence for the role of β-hairpin structure in Aβ oligomerization and neurotoxicity. Inspired by these β-hairpin structures, our laboratory developed a macrocyclic β-sheet peptide derived from Aβ17–36 designed to mimic an Aβ β-hairpin and reported its X-ray crystallographic structure. This peptide (peptide 1) consists of two β-strands comprising Aβ17–23 and Aβ30–36 covalently linked by two δ-linked ornithine (δOrn) β-turn mimics. The δOrn that connects residues D23 and A30 replaces the Aβ24–29 loop. The δOrn that connects residues L17 and V36 enforces β-hairpin structure. We incorporated an N-methyl group at position G33 to prevent uncontrolled aggregation and precipitation of the peptide. To improve the solubility of the peptide we replaced M35 with the hydrophilic isostere of methionine, ornithine (α-linked) (Figure 1B). The X-ray crystallographic structure of peptide 1 reveals that it folds to form a β-hairpin that assembles to form trimers and that the trimers further assemble to form hexamers and dodecamers. (A) Cartoon\nillustrating the design of peptides 1 and 2 and their relationship to an Aβ17–36 β-hairpin.\n(B) Chemical structure of peptide 1 illustrating Aβ17–23 and Aβ30–36, M35Orn, the N-methyl group, and the δ-linked\nornithine turns. (C) Chemical structure of peptide 2 illustrating\nAβ17–36, the N-methyl group,\nthe disulfide bond across positions 24 and 29, and the δ-linked\nornithine turn. Our design of peptide 1 omitted the Aβ24–29 loop. To visualize the Aβ24–29 loop, we performed replica-exchange molecular dynamics (REMD) simulations on Aβ17–36 using the X-ray crystallographic coordinates of Aβ17–23 and Aβ30–36 from peptide 1. These studies provided a working model for a trimer of Aβ17–36 β-hairpins and demonstrated that the trimer should be capable of accommodating the Aβ24–29 loop. In the current study we set out to restore the Aβ24–29 loop, reintroduce the methionine residue at position 35, and determine the X-ray crystallographic structures of oligomers that form. We designed peptide 2 as a homologue of peptide 1 that embodies these ideas. Peptide 2 contains a methionine residue at position 35 and an Aβ24–29 loop with residues 24 and 29 (Val and Gly) mutated to cysteine and linked by a disulfide bond (Figure 1C). Here, we describe the development of peptide 2 and report the X-ray crystallographic structures of the trimer, dodecamer, and annular pore observed within the crystal structure. Results Development of Peptide 2 We developed peptide 2 from peptide 1 by an iterative process, in which we first attempted to restore the Aβ24–29 loop without a disulfide linkage. We envisioned peptide 3 as a homologue of peptide 1 with the Aβ24–29 loop in place of the δOrn that connects D23 and A30 and p-iodophenylalanine (FI) in place of F19. We routinely use p-iodophenylalanine to determine the X-ray crystallographic phases. After determining the X-ray crystallographic structure of the p-iodophenylalanine variant we attempt to determine the structure of the native phenylalanine compound by isomorphous replacement. Upon synthesizing peptide 3, we found that it formed an amorphous precipitate in most crystallization conditions screened and failed to afford crystals in any condition. We postulate that the loss of the δOrn constraint leads to conformational heterogeneity that prevents peptide 3 from crystallizing. To address this issue, we next incorporated a disulfide bond between residues 24 and 29 as a conformational constraint that serves as a surrogate for δOrn. We designed peptide 4 to embody this idea, mutating Val24 and Gly29 to cysteine and forming an interstrand disulfide linkage. We mutated these residues because they occupy the same position as the δOrn that connects D23 and A30 in peptide 1. Residues V24 and G29 form a non-hydrogen-bonded pair, which can readily accommodate disulfide linkages in antiparallel β-sheets. Disulfide bonds across non-hydrogen-bonded pairs stabilize β-hairpins, while disulfide bonds across hydrogen-bonded pairs do not. Although the disulfide bond between positions 24 and 29 helps stabilize the β-hairpin, it does not alter the charge or substantially change the hydrophobicity of the Aβ17–36 β-hairpin. We were gratified to find that peptide 4 afforded crystals suitable for X-ray crystallography. As the next step in the iterative process, we determined the X-ray crystallographic structure of this peptide (PDB 5HOW). After determining the X-ray crystallographic structure of peptide 4 we reintroduced the native phenylalanine at position 19 and the methionine at position 35 to afford peptide 2. We completed the iterative process—from 1 to 3 to 4 to 2—by successfully determining the X-ray crystallographic structure of peptide 2 (PDB 5HOX and 5HOY). The following sections describe the synthesis of peptides 2–4 and the X-ray crystallographic structure of peptide 2. Synthesis of Peptides 2–4 We synthesized peptides 2–4 by similar procedures to those we have developed for other macrocyclic peptides. Our laboratory routinely prepares macrocyclic peptides by solid-phase synthesis of the corresponding linear peptide on 2-chlorotrityl resin, followed by cleavage of the protected linear peptide from the resin, solution-phase macrolactamization, and deprotection of the resulting macrocyclic peptide. In synthesizing peptides 2 and 4 we formed the disulfide linkage after macrolactamization and deprotection of the acid-labile side chain protecting groups. We used acid-stable Acm-protected cysteine residues at positions 24 and 29 and removed the Acm groups by oxidation with I2 in aqueous acetic acid to afford the disulfide linkage. Peptides 2–4 were purified by RP-HPLC. Crystallization, X-ray Crystallographic Data Collection, Data Processing, and Structure Determination of Peptides 2 and 4 We screened crystallization conditions for peptide 4 in a 96-well-plate format using three different Hampton Research crystallization kits (Crystal Screen, Index, and PEG/Ion) with three ratios of peptide and mother liquor per condition (864 experiments). Peptide 4 afforded crystals in a single set of conditions containing HEPES buffer and Jeffamine M-600—the same crystallization conditions that afforded crystals of peptide 1. Peptide 2 also afforded crystals in these conditions. We further optimized these conditions to rapidly (∼72 h) yield crystals suitable for X-ray crystallography. The optimized conditions consist of 0.1 M HEPES at pH 6.4 with 31% Jeffamine M-600 for peptide 4 and 0.1 M HEPES pH 7.1 with 29% Jeffamine M-600 for peptide 2. Crystal diffraction data for peptides 4 and 2 were collected in-house with a Rigaku MicroMax 007HF X-ray diffractometer at 1.54 Å wavelength. Crystal diffraction data for peptide 2 were also collected at the Advanced Light Source at Lawrence Berkeley National Laboratory with a synchrotron source at 1.00 Å wavelength to achieve higher resolution. Data from peptides 4 and 2 suitable for refinement at 2.30 Å were obtained from the diffractometer; data from peptide 2 suitable for refinement at 1.90 Å were obtained from the synchrotron. Data for peptides 4 and 2 were scaled and merged using XDS. Phases for peptide 4 were determined by single-wavelength anomalous diffraction (SAD) phasing by using the coordinates of the iodine anomalous signal from p-iodophenylalanine. Phases for peptide 2 were determined by isomorphous replacement of peptide 4. The structures of peptides 2 and 4 were solved and refined in the P6122 space group. Coordinates for hydrogens were generated by phenix.refine during refinement. The asymmetric unit of each peptide consists of six monomers, arranged as two trimers. Peptides 2 and 4 form morphologically identical structures and assemblies in the crystal lattice. X-ray Crystallographic Structure of Peptide 2 and the Oligomers It Forms The X-ray crystallographic structure of peptide 2 reveals that it folds to form a twisted β-hairpin comprising two β-strands connected by a loop (Figure 2A). Eight residues make up each surface of the β-hairpin: L17, F19, A21, D23, A30, I32, L34, and V36 make up one surface; V18, F20, E22, C24, C29, I31, G33, and M35 make up the other surface. The β-strands of the monomers in the asymmetric unit are virtually identical, differing primarily in rotamers of F20, E22, C24, C29, I31, and M35 (Figure S1). The disulfide linkages suffered radiation damage under synchrotron radiation. We refined three of the β-hairpins with intact disulfide linkages and three with thiols to represent cleaved disulfide linkages in the synchrotron data set (PDB 5HOX). No evidence for cleavage of the disulfides was observed in the refinement of the data set collected on the X-ray diffractometer, and we refined all disulfide linkages as intact (PDB 5HOY). X-ray crystallographic\nstructure of peptide 2 (PDB 5HOX, synchrotron data\nset). (A) X-ray crystallographic structure of a representative β-hairpin\nmonomer formed by peptide 2. (B) Overlay of the six β-hairpin\nmonomers in the asymmetric unit. The β-hairpins are shown as\ncartoons to illustrate the differences in the Aβ25–28 loops. The Aβ25–28 loops of the six monomers within the asymmetric unit vary substantially in backbone geometry and side chain rotamers (Figures 2B and S1). The electron density for the loops is weak and diffuse compared to the electron density for the β-strands. The B values for the loops are large, indicating that the loops are dynamic and not well ordered. Thus, the differences in backbone geometry and side chain rotamers among the loops are likely of little significance and should be interpreted with caution. Peptide 2 assembles into oligomers similar in morphology to those formed by peptide 1. Like peptide 1, peptide 2 forms a triangular trimer, and four trimers assemble to form a dodecamer. In the higher-order assembly of the dodecamers formed by peptide 2 a new structure emerges, not seen in peptide 1, an annular pore consisting of five dodecamers. Trimer Peptide 2 forms a trimer, much like that which we observed previously for peptide 1, in which three β-hairpins assemble to form an equilateral triangle (Figure 3A). The trimer maintains all of the same stabilizing contacts as those of peptide 1. Hydrogen bonding and hydrophobic interactions between residues on the β-strands comprising Aβ17–23 and Aβ30–36 stabilize the core of the trimer. The disulfide bonds between residues 24 and 29 are adjacent to the structural core of the trimer and do not make any substantial intermolecular contacts. Two crystallographically distinct trimers comprise the peptide portion of the asymmetric unit. The two trimers are almost identical in structure, differing slightly among side chain rotamers and loop conformations. X-ray crystallographic structure of the trimer formed\nby peptide 2. (A) Triangular trimer. The three water\nmolecules in the\ncenter hole of the trimer are shown as spheres. (B) Detailed view\nof the intermolecular hydrogen bonds between the main chains of V18 and E22 and δOrn and C24, at the three corners of the triangular trimer. (C) The F19 face of the trimer, with key side chains shown as spheres. (D) The\nF20 face of the trimer, with key side chains as spheres. A network of 18 intermolecular hydrogen bonds helps stabilize the trimer. At the corners of the trimer, the pairs of β-hairpin monomers form four hydrogen bonds: two between the main chains of V18 and E22 and two between δOrn and the main chain of C24 (Figure 3B). Three ordered water molecules fill the hole in the center of the trimer, hydrogen bonding to each other and to the main chain of F20 (Figure 3A). Hydrophobic contacts between residues at the three corners of the trimer, where the β-hairpins meet, further stabilize the trimer. At each corner, the side chains of residues L17, F19, and V36 of one β-hairpin pack against the side chains of residues A21, I32, L34, and also D23 of the adjacent β-hairpin to create a hydrophobic cluster (Figure 3C). The three hydrophobic clusters create a large hydrophobic surface on one face of the trimer. The other face of the trimer displays a smaller hydrophobic surface, which includes the side chains of residues V18, F20, and I31 of the three β-hairpins (Figure 3D). In subsequent discussion, we designate the former surface the “F19 face” and the latter surface the “F20 face”. Dodecamer Four trimers assemble to form a dodecamer. The four trimers arrange in a tetrahedral fashion, creating a central cavity inside the dodecamer. Because each trimer is triangular, the resulting arrangement resembles an octahedron. Each of the 12 β-hairpins constitutes an edge of the octahedron, and the triangular trimers occupy four of the eight faces of the octahedron. Figure 4A illustrates the octahedral shape of the dodecamer. Figure 4B illustrates the tetrahedral arrangement of the four trimers. X-ray\ncrystallographic structure of the dodecamer formed by peptide 2. (A) View of the dodecamer that illustrates the octahedral\nshape. (B) View of the dodecamer that illustrates the tetrahedral\narrangement of the four trimers that comprise the dodecamer. (C) View\nof two trimer subunits from inside the cavity of the dodecamer. Residues\nL17, L34, and V36 are shown as spheres,\nillustrating the hydrophobic packing that occurs at the six vertices\nof the dodecamer. (D) Detailed view of one of the six vertices of\nthe dodecamer. The F19 faces of the trimers line the interior of the dodecamer. At the six vertices, hydrophobic packing between the side chains of L17, L34, and V36 helps stabilize the dodecamer (Figures 4C and D). Salt bridges between the side chains of D23 and δOrn at the vertices further stabilize the dodecamer. Each of the six vertices includes two Aβ25–28 loops that extend past the core of the dodecamer without making any substantial intermolecular contacts. The exterior of the dodecamer displays four F20 faces (Figure S3). In the crystal lattice, each F20 face of one dodecamer packs against an F20 face of another dodecamer. Although the asymmetric unit comprises half a dodecamer, the crystal lattice may be thought of as being built of dodecamers. The electron density map for the X-ray crystallographic structure of peptide 2 has long tubes of electron density inside the central cavity of the dodecamer. The shape and length of the electron density is consistent with the structure of Jeffamine M-600, which is an essential component of the crystallization conditions. Jeffamine M-600 is a polypropylene glycol derivative with a 2-methoxyethoxy unit at one end and a 2-aminopropyl unit at the other end. Its average molecular weight is about 600 Da, which corresponds to nine propylene glycol units. Although Jeffamine M-600 is a heterogeneous mixture with varying chain lengths and stereochemistry, we modeled a single stereoisomer with nine propylene glycol units (n = 9) to fit the electron density. The Jeffamine M-600 appears to stabilize the dodecamer by occupying the central cavity and making hydrophobic contacts with residues lining the cavity (Figure S3). In a dodecamer formed by full-length Aβ, the hydrophobic C-terminal residues (Aβ37–40 or Aβ37–42) might play a similar role in filling the dodecamer and thus create a packed hydrophobic core within the central cavity of the dodecamer. Annular Pore Five dodecamers assemble to form an annular porelike structure (Figure 5A). Hydrophobic packing between the F20 faces of trimers displayed on the outer surface of each dodecamer stabilizes the porelike assembly. Two morphologically distinct interactions between trimers occur at the interfaces of the five dodecamers: one in which the trimers are eclipsed (Figure 5B), and one in which the trimers are staggered (Figure 5C). Hydrophobic packing between the side chains of F20, I31, and E22 stabilizes these interfaces (Figure 5D and E). The annular pore contains three eclipsed interfaces and two staggered interfaces. The eclipsed interfaces occur between dodecamers 1 and 2, 1 and 5, and 3 and 4, as shown in Figure 5A. The staggered interfaces occur between dodecamers 2 and 3 and 4 and 5. The annular pore is not completely flat, instead, adopting a slightly puckered shape, which accommodates the eclipsed and staggered interfaces. Ten Aβ25–28 loops from the vertices of the five dodecamers line the hole in the center of the pore. The hydrophilic side chains of S26, N27, and K28 decorate the hole. X-ray crystallographic structure of the annular pore formed by\npeptide 2. (A) Annular porelike structure illustrating\nthe relationship of the five dodecamers that form the pore (top view).\n(B) Eclipsed interface between dodecamers 1 and 2 (side view). The\nsame eclipsed interface also occurs between dodecamers 1 and 5 and\n3 and 4. (C) Staggered interface between dodecamers 2 and 3 (side\nview). The same staggered interface also occurs between dodecamers\n4 and 5. (D) Eclipsed interface between dodecamers 1 and 5 (top view).\nResidues F20, I31, and E22 are shown\nas spheres to detail the hydrophobic packing. (E) Staggered interface\nbetween dodecamers 2 and 3 (top view). Residues F20, I31, and E22 are shown as spheres to detail the hydrophobic\npacking. The annular pore is comparable in size to other large protein assemblies. The outer diameter is ∼11–12 nm. The diameter of the hole in the center of the pore is ∼2 nm. The thickness of the pore is ∼5 nm, which is comparable to that of a lipid bilayer membrane. It is important to note that the annular pore formed by peptide 2 is not a discrete unit in the crystal lattice. Rather, the crystal lattice is composed of conjoined annular pores in which all four F20 faces on the surface of each dodecamer contact F20 faces on other dodecamers (Figure S4). The crystal lattice shows how the dodecamers can further assemble to form larger structures. Each dodecamer may be thought of as a tetravalent building block with the potential to assemble on all four faces to form higher-order supramolecular assemblies. Discussion The X-ray crystallographic study of peptide 2 described here provides high-resolution structures of oligomers formed by an Aβ17–36 β-hairpin. The crystallographic assembly of peptide 2 into a trimer, dodecamer, and annular pore provides a model for the assembly of the full-length Aβ peptide to form oligomers. In this model Aβ folds to form a β-hairpin comprising the hydrophobic central and C-terminal regions. Three β-hairpins assemble to form a trimer, and four trimers assemble to form a dodecamer. The dodecamers further assemble to form an annular pore (Figure 6). Model for the hierarchical assembly of an Aβ\nβ-hairpin\ninto a trimer, dodecamer, and annular pore based on the crystallographic\nassembly of peptide 2. Monomeric Aβ folds to form\na β-hairpin in which the hydrophobic central and C-terminal regions form an antiparallel β-sheet. Three β-hairpin\nmonomers assemble to form a triangular trimer. Four triangular trimers\nassemble to form a dodecamer. Five dodecamers assemble to form an\nannular pore. The molecular weights shown correspond to an Aβ42 monomer (∼4.5 kDa), an Aβ42 trimer\n(∼13.5 kDa), an Aβ42 dodecamer (∼54\nkDa), and an Aβ42 annular pore composed of five dodecamers\n(∼270 kDa). The model put forth in Figure 6 is consistent with the current understanding of endogenous Aβ oligomerization and explains at atomic resolution many key observations about Aβ oligomers. Two general types of endogenous Aβ oligomers have been observed: Aβ oligomers that occur on a pathway to fibrils, or “fibrillar oligomers”, and Aβ oligomers that evade a fibrillar fate, or “nonfibrillar oligomers”.− Fibrillar oligomers accumulate in Alzheimer’s disease later than nonfibrillar oligomers and coincide with the deposition of plaques. Nonfibrillar oligomers accumulate early in Alzheimer’s disease before plaque deposition. Fibrillar and nonfibrillar oligomers have structurally distinct characteristics, which are reflected in their reactivity with the fibril-specific OC antibody and the oligomer-specific A11 antibody. Fibrillar oligomers are recognized by the OC antibody but not the A11 antibody, whereas nonfibrillar oligomers are recognized by the A11 antibody but not the OC antibody. These criteria have been used to classify the Aβ oligomers that accumulate in vivo. Aβ dimers have been classified as fibrillar oligomers, whereas Aβ trimers, Aβ*56, and APFs have been classified as nonfibrillar oligomers. Larson and Lesné proposed a model for the endogenous production of nonfibrillar oligomers that explains these observations. In this model, folded Aβ monomer assembles into a trimer, the trimer further assembles into hexamers and dodecamers, and the dodecamers further assemble to form annular protofibrils. The hierarchical assembly of peptide 2 is consistent with this model; and the trimer, dodecamer, and annular pore formed by peptide 2 may share similarities to the trimers, Aβ*56, and APFs observed in vivo. At this point, we can only speculate whether the trimer and dodecamer formed by peptide 2 share structural similarities to Aβ trimers and Aβ*56, as little is known about the structure of Aβ trimers and Aβ*56. The crystallographically observed annular pore formed by peptide 2 is morphologically similar to the APFs formed by full-length Aβ. The annular pore formed by peptide 2 is comparable in size to the APFs prepared in vitro or isolated from Alzheimer’s brains (Figure 7 and Table 1). The varying sizes of APFs formed by full-length Aβ might result from differences in the number of oligomer subunits comprising each APF. Although the annular pore formed by peptide 2 contains five dodecamer subunits, pores containing fewer or more subunits can easily be envisioned. The dodecamers that comprise the annular pore exhibit two modes of assembly—eclipsed interactions and staggered interactions between the F20 faces of trimers within dodecamers. These two modes of assembly might reflect a dynamic interaction between dodecamers, which could permit assemblies of more dodecamers into larger annular pores. Surface views of the annular pore formed\nby peptide 2. (A) Top view. (B) Side view. Annular Pores Formed by Aβ and\nPeptide 2 annular pore\t \tsource\touter diameter\tinner diameter\tobservation\t \tmethod\t \tpeptide 2\t ∼11–12 nm\t∼2 nm\tX-ray crystallography\t \tsynthetic Aβ\t7–10 nm\t1.5–2 nm\tTEM\t \tsynthetic Aβ\t16 nm\tnot reported\tAFM\t \tsynthetic Aβ\t8–25 nm\tnot reported\tTEM\t \tAlzheimer’s brain\t11–14 nm\t2.5–4 nm\tTEM\t \t Dot blot analysis shows that peptide 2 is reactive toward the A11 antibody (Figure S5). This reactivity suggests that peptide 2 forms oligomers in solution that share structural similarities to the nonfibrillar oligomers formed by full-length Aβ. Further studies are needed to elucidate the species that peptide 2 forms in solution and to study their biological properties. This is an active area of research in our laboratory. Preliminary attempts to study these species by SEC and SDS-PAGE have not provided a clear measure of the structures formed in solution. The difficulty in studying the oligomers formed in solution may reflect the propensity of the dodecamer to assemble on all four F20 faces. The X-ray crystallographic structure and A11 reactivity of peptide 2 support the model proposed by Larsen and Lesné and suggest that β-hairpins constitute a fundamental building block for nonfibrillar oligomers. What makes β-hairpins special is that three β-hairpins can nestle together to form trimers, stabilized by a network of hydrogen bonds and hydrophobic interactions. This mode of assembly is not unique to Aβ. The foldon domain of bacteriophage T4 fibritin is composed of three β-hairpins that assemble into a triangular trimer similar to the triangular trimer formed by peptide 2. Additionally, our research group has observed a similar assembly of a β-hairpin peptide derived from β2-microglobulin. Conclusion Although we began these studies with a relatively simple hypothesis—that the trimers and dodecamers formed by peptide 1 could accommodate the Aβ24–29 loop—an even more exciting finding has emerged—that the dodecamers can assemble to form annular pores. This finding could not have been anticipated from the X-ray crystallographic structure of peptide 1 and reveals a new level of hierarchical assembly that recapitulates micrographic observations of annular protofibrils. The crystallographically observed dodecamer, in turn, recapitulates the observation of Aβ*56, which appears to be a dodecamer of Aβ. The crystallographically observed trimer recapitulates the Aβ trimers that are observed even before the onset of symptoms in Alzheimer’s disease. Our approach of constraining Aβ17–36 into a β-hairpin conformation and blocking aggregation with an N-methyl group has allowed us to crystallize a large fragment of what is generally considered to be an uncrystallizable peptide. We believe this iterative, “bottom up” approach of identifying the minimal modification required to crystallize Aβ peptides will ultimately allow larger fragments of Aβ to be crystallized, thus providing greater insights into the structures of Aβ oligomers. Supporting Information Available Procedures for the synthesis and crystallization of peptides 2–4; details of X-ray crystallographic data collection, processing, and refinement; procedure and data for dot blot analysis (PDF) Crystallographic coordinates of peptide 2 deposited into the Protein Data Bank (PDB) with code 5HOX (data collected on a synchrotron at 1.00 Å wavelength) (PDB) Crystallographic coordinates of peptide 2 deposited into the Protein Data Bank (PDB) with code 5HOY (data collected on an X-ray diffractometer at 1.54 Å wavelength) (PDB) Crystallographic coordinates of peptide 4 deposited into the Protein Data Bank (PDB) with code 5HOW (PDB) Crystallographic coordinates of the dodecamer formed by peptide 2 (PDB) Crystallographic coordinates of the annular pore formed by peptide 2 (PDB) Crystallographic data for 5HOW (CIF) Crystallographic data for 5HOX (CIF) Crystallographic data for 5HOY (CIF) The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/jacs.6b01332. Supplementary Material The authors declare no competing financial interest. References                                      recently reported the NMR structure of a hexameric peptide barrel formed by this disulfide constrained Aβ: Lendel, C.; Bjerring, M.; Dubnovitsky, A.; Kelly, R. T.; Filippov, A.; Antzutkin, O. N.; Nielsen, N. C.; Härd, T                  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diff --git a/annotated_BioC_JSON/PMC4832331_ann.json b/annotated_BioC_JSON/PMC4832331_ann.json
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@@ -0,0 +1 @@
+[{"sourceid":"4832331","sourcedb":"","project":"","target":"","text":"Structural insights into the Escherichia coli lysine decarboxylases and molecular determinants of interaction with the AAA+ ATPase RavA The inducible lysine decarboxylase LdcI is an important enterobacterial acid stress response enzyme whereas LdcC is its close paralogue thought to play mainly a metabolic role. A unique macromolecular cage formed by two decamers of the Escherichia coli LdcI and five hexamers of the AAA+ ATPase RavA was shown to counteract acid stress under starvation. Previously, we proposed a pseudoatomic model of the LdcI-RavA cage based on its cryo-electron microscopy map and crystal structures of an inactive LdcI decamer and a RavA monomer. We now present cryo-electron microscopy 3D reconstructions of the E. coli LdcI and LdcC, and an improved map of the LdcI bound to the LARA domain of RavA, at pH optimal for their enzymatic activity. Comparison with each other and with available structures uncovers differences between LdcI and LdcC explaining why only the acid stress response enzyme is capable of binding RavA. We identify interdomain movements associated with the pH-dependent enzyme activation and with the RavA binding. Multiple sequence alignment coupled to a phylogenetic analysis reveals that certain enterobacteria exert evolutionary pressure on the lysine decarboxylase towards the cage-like assembly with RavA, implying that this complex may have an important function under particular stress conditions. Enterobacterial inducible decarboxylases of basic amino acids lysine, arginine and ornithine have a common evolutionary origin and belong to the α-family of pyridoxal-5′-phosphate (PLP)-dependent enzymes. They counteract acid stress experienced by the bacterium in the host digestive and urinary tract, and in particular in the extremely acidic stomach. Each decarboxylase is induced by an excess of the target amino acid and a specific range of extracellular pH, and works in conjunction with a cognate inner membrane antiporter. Decarboxylation of the amino acid into a polyamine is catalysed by a PLP cofactor in a multistep reaction that consumes a cytoplasmic proton and produces a CO2 molecule passively diffusing out of the cell, while the polyamine is excreted by the antiporter in exchange for a new amino acid substrate. Consequently, these enzymes buffer both the bacterial cytoplasm and the local extracellular environment. These amino acid decarboxylases are therefore called acid stress inducible or biodegradative to distinguish them from their biosynthetic lysine and ornithine decarboxylase paralogs catalysing the same reaction but responsible for the polyamine production at neutral pH. Inducible enterobacterial amino acid decarboxylases have been intensively studied since the early 1940 because the ability of bacteria to withstand acid stress can be linked to their pathogenicity in humans. In particular, the inducible lysine decarboxylase LdcI (or CadA) attracts attention due to its broad pH range of activity and its capacity to promote survival and growth of pathogenic enterobacteria such as Salmonella enterica serovar Typhimurium, Vibrio cholerae and Vibrio vulnificus under acidic conditions. Furthermore, both LdcI and the biosynthetic lysine decarboxylase LdcC of uropathogenic Escherichia coli (UPEC) appear to play an important role in increased resistance of this pathogen to nitrosative stress produced by nitric oxide and other damaging reactive nitrogen intermediates accumulating during the course of urinary tract infections (UTI). This effect is attributed to cadaverine, the diamine produced by decarboxylation of lysine by LdcI and LdcC, that was shown to enhance UPEC colonisation of the bladder. In addition, the biosynthetic E. coli lysine decarboxylase LdcC, long thought to be constitutively expressed in low amounts, was demonstrated to be strongly upregulated by fluoroquinolones via their induction of RpoS. A direct correlation between the level of cadaverine and the resistance of E. coli to these antibiotics commonly used as a first-line treatment of UTI could be established. Both acid pH and cadaverine induce closure of outer membrane porins thereby contributing to bacterial protection from acid stress, but also from certain antibiotics, by reduction in membrane permeability. The crystal structure of the E. coli LdcI as well as its low resolution characterisation by electron microscopy (EM) showed that it is a decamer made of two pentameric rings. Each monomer is composed of three domains – an N-terminal wing domain (residues 1–129), a PLP-binding core domain (residues 130–563), and a C-terminal domain (CTD, residues 564–715). Monomers tightly associate via their core domains into 2-fold symmetrical dimers with two complete active sites, and further build a toroidal D5-symmetrical structure held by the wing and core domain interactions around the central pore, with the CTDs at the periphery. Ten years ago we showed that the E. coli AAA+ ATPase RavA, involved in multiple stress response pathways, tightly interacted with LdcI but was not capable of binding to LdcC. We described how two double pentameric rings of the LdcI tightly associate with five hexameric rings of RavA to form a unique cage-like architecture that enables the bacterium to withstand acid stress even under conditions of nutrient deprivation eliciting stringent response. Furthermore, we recently solved the structure of the E. coli LdcI-RavA complex by cryo-electron microscopy (cryoEM) and combined it with the crystal structures of the individual proteins. This allowed us to make a pseudoatomic model of the whole assembly, underpinned by a cryoEM map of the LdcI-LARA complex (with LARA standing for LdcI associating domain of RavA), and to identify conformational rearrangements and specific elements essential for complex formation. The main determinants of the LdcI-RavA cage assembly appeared to be the N-terminal loop of the LARA domain of RavA and the C-terminal β-sheet of LdcI. In spite of this wealth of structural information, the fact that LdcC does not interact with RavA, although the two lysine decarboxylases are 69% identical and 84% similar, and the physiological significance of the absence of this interaction remained unexplored. To solve this discrepancy, in the present work we provided a three-dimensional (3D) cryoEM reconstruction of LdcC and compared it with the available LdcI and LdcI-RavA structures. Given that the LdcI crystal structures were obtained at high pH where the enzyme is inactive (LdcIi, pH 8.5), whereas the cryoEM reconstructions of LdcI-RavA and LdcI-LARA were done at acidic pH optimal for the enzymatic activity, for a meaningful comparison, we also produced a 3D reconstruction of the LdcI at active pH (LdcIa, pH 6.2). This comparison pinpointed differences between the biodegradative and the biosynthetic lysine decarboxylases and brought to light interdomain movements associated to pH-dependent enzyme activation and RavA binding, notably at the predicted RavA binding site at the level of the C-terminal β-sheet of LdcI. Consequently, we tested the capacity of cage formation by LdcI-LdcC chimeras where we interchanged the C-terminal β-sheets in question. Finally, we performed multiple sequence alignment of 22 lysine decarboxylases from Enterobacteriaceae containing the ravA-viaA operon in their genome. Remarkably, this analysis revealed that several specific residues in the above-mentioned β-sheet, independently of the rest of the protein sequence, are sufficient to define if a particular lysine decarboxylase should be classified as an “LdcC-like” or an “LdcI-like”. Moreover, this classification perfectly agrees with the genetic environment of the lysine decarboxylase genes. This fascinating parallelism between the propensity for RavA binding and the genetic environment of an enterobacterial lysine decarboxylase, as well as the high degree of conservation of this small structural motif, emphasize the functional importance of the interaction between biodegradative enterobacterial lysine decarboxylases and the AAA+ ATPase RavA. Results and Discussion CryoEM 3D reconstructions of LdcC, LdcIa and LdcI-LARA In the frame of this work, we produced two novel subnanometer resolution cryoEM reconstructions of the E. coli lysine decarboxylases at pH optimal for their enzymatic activity – a 5.5 Å resolution cryoEM map of the LdcC (pH 7.5) for which no 3D structural information has been previously available (Figs 1A,B and S1), and a 6.1 Å resolution cryoEM map of the LdcIa, (pH 6.2) (Figs 1C,D and S2). In addition, we improved our earlier cryoEM map of the LdcI-LARA complex from 7.5 Å to 6.2 Å resolution (Figs 1E,F and S3). Based on these reconstructions, reliable pseudoatomic models of the three assemblies were obtained by flexible fitting of either the crystal structure of LdcIi or a derived structural homology model of LdcC (Table S1). Significant differences between these pseudoatomic models can be interpreted as movements between specific biological states of the proteins as described below. The wing domains as a stable anchor at the center of the double-ring As a first step of a comparative analysis, we superimposed the three cryoEM reconstructions (LdcIa, LdcI-LARA and LdcC) and the crystal structure of the LdcIi decamer (Fig. 2 and Movie S1). This superposition reveals that the densities lining the central hole of the toroid are roughly at the same location, while the rest of the structure exhibits noticeable changes. Specifically, at the center of the double-ring the wing domains of the subunits provide the conserved basis for the assembly with the lowest root mean square deviation (RMSD) (between 1.4 and 2 Å for the Cα atoms only), whereas the peripheral CTDs containing the RavA binding interface manifest the highest RMSD (up to 4.2 Å) (Table S2). In addition, the wing domains of all structures are very similar, with the RMSD after optimal rigid body alignment (RMSDmin) less than 1.1 Å. Thus, taking the limited resolution of the cryoEM maps into account, we consider that the wing domains of all the four structures are essentially identical and that in the present study the RMSD of less than 2 Å can serve as a baseline below which differences may be assumed as insignificant. This preservation of the central part of the double-ring assembly may help the enzymes to maintain their decameric state upon activation and incorporation into the LdcI-RavA cage. The core domain and the active site rearrangements upon pH-dependent enzyme activation and LARA binding Both visual inspection (Fig. 2) and RMSD calculations (Table S2) show that globally the three structures at active pH (LdcIa, LdcI-LARA and LdcC) are more similar to each other than to the structure determined at high pH conditions (LdcIi). The decameric enzyme is built of five dimers associating into a 5-fold symmetrical double-ring (two monomers making a dimer are delineated in Fig. 1). As common for the α family of the PLP-dependent decarboxylases, dimerization is required for the enzymatic activity because the active site is buried in the dimer interface (Fig. 3A,B). This interface is formed essentially by the core domains with some contribution of the CTDs. The core domain is built by the PLP-binding subdomain (PLP-SD, residues 184–417) flanked by two smaller subdomains rich in partly disordered loops – the linker region (residues 130–183) and the subdomain 4 (residues 418–563). Zooming in the variations in the PLP-SD shows that most of the structural changes concern displacements in the active site (Fig. 3C–F). The most conspicuous differences between the PLP-SDs can be linked to the pH-dependent activation of the enzymes. The resolution of the cryoEM maps does not allow modeling the position of the PLP moiety and calls for caution in detailed mechanistic interpretations in terms of individual amino acids. Therefore we restrict our analysis to secondary structure elements. In particular, transition from LdcIi to LdcI-LARA involves ~3.5 Å and ~4.5 Å shifts away from the 5-fold axis in the active site α-helices spanning residues 218–232 and 246–254 respectively (Fig. 3C–E). Between these two extremes, the PLP-SDs of LdcIa and LdcC are similar both in the context of the decamer (Fig. 3F) and in terms of RMSDmin = 0.9 Å, which probably reflects the fact that, at the optimal pH, these lysine decarboxylases have a similar enzymatic activity. In addition, our earlier biochemical observation that the enzymatic activity of LdcIa is unaffected by RavA binding is consistent with the relatively small changes undergone by the active site upon transition from LdcIa to LdcI-LARA. Worthy of note, our previous comparison of the crystal structure of LdcIi with that of the inducible arginine decarboxylase AdiA revealed high conservation of the PLP-coordinating residues and identified a patch of negatively charged residues lining the active site channel as a potential binding site for the target amino acid substrate (Figs S3 and S4 in ref.). Rearrangements of the ppGpp binding pocket upon pH-dependent enzyme activation and LARA binding An inhibitor of the LdcI and LdcC activity, the stringent response alarmone ppGpp, is known to bind at the interface between neighboring monomers within each ring (Fig. S4). The ppGpp binding pocket is made up by residues from all domains and is located approximately 30 Å away from the PLP moiety. Whereas the crystal structure of the ppGpp-LdcIi was solved to 2 Å resolution, only a 4.1 Å resolution structure of the ppGpp-free LdcIi could be obtained. At this resolution, the apo-LdcIi and ppGpp-LdcIi structures (both solved at pH 8.5) appeared indistinguishable except for the presence of ppGpp (Fig. S11 in ref. ). Thus, we speculated that inhibition of LdcI by ppGpp would be accompanied by a transduction of subtle structural changes at the level of individual amino acid side chains between the ppGpp binding pocket and the active site of the enzyme. All our current cryoEM reconstructions of the lysine decarboxylases were obtained in the absence of ppGpp in order to be closer to the active state of the enzymes under study. While differences in the ppGpp binding site could indeed be visualized (Fig. S4), the level of resolution warns against speculations about their significance. The fact that interaction with RavA reduces the ppGpp affinity for LdcI despite the long distance of ~30 Å between the LARA domain binding site and the closest ppGpp binding pocket (Fig. S5) seems to favor an allosteric regulation mechanism. Interestingly, although a number of ppGpp binding residues are strictly conserved between LdcI and AdiA that also forms decamers at low pH optimal for its arginine decarboxylase activity, no ppGpp regulation of AdiA could be demonstrated. Swinging and stretching of the CTDs upon pH-dependent LdcI activation and LARA binding Inspection of the superimposed decameric structures (Figs 2 and S6) suggests a depiction of the wing domains as an anchor around which the peripheral CTDs swing. This swinging movement seems to be mediated by the core domains and is accompanied by a stretching of the whole LdcI subunits attracted by the RavA magnets. Indeed, all CTDs have very similar structures (RMSDmin \u003c1 Å). Yet the superposition of the decamers lays bare a progressive movement of the CTD as a whole upon enzyme activation by pH and the binding of LARA. The LdcIi monomer is the most compact, whereas LdcIa and especially LdcI-LARA gradually extend their CTDs towards the LARA domain of RavA (Figs 2 and 4). These small but noticeable swinging and stretching (up to ~4 Å) may be related to the incorporation of the LdcI decamer into the LdcI-RavA cage. The C-terminal β-sheet of a lysine decarboxylase as a major determinant of the interaction with RavA In our previous contribution, based on the fit of the LdcIi and the LARA crystal structures into the LdcI-LARA cryoEM density, we predicted that the LdcI-RavA interaction should involve the C-terminal two-stranded β-sheet of the LdcI. Our present cryoEM maps and pseudoatomic models provide first structure-based insights into the differences between the inducible and the constitutive lysine decarboxylases. However, at the level of this structural element the two proteins are actually surpisingly similar. Therefore, we wanted to check the influence of the primary sequence of the two proteins in this region on their ability to interact with RavA. To this end, we swapped the relevant β-sheets of the two proteins and produced their chimeras, namely LdcIC (i.e. LdcI with the C-terminal β-sheet of LdcC) and LdcCI (i.e. LdcC with the C-terminal β-sheet of LdcI) (Fig. 5A–C). Both constructs could be purified and could form decamers visually indistinguishable from the wild-type proteins. As expected, binding of LdcI to RavA was completely abolished by this procedure and no LdcIC-RavA complex could be detected. On the contrary, introduction of the C-terminal β-sheet of LdcI into LdcC led to an assembly of the LdcCI-RavA complex. On the negative stain EM grid, the chimeric cages appeared less rigid than the native LdcI-RavA, which probably means that the environment of the β-sheet contributes to the efficiency of the interaction and the stability of the entire architecture (Fig. 5D–F). The C-terminal β-sheet of a lysine decarboxylase is a highly conserved signature allowing to distinguish between LdcI and LdcC Alignment of the primary sequences of the E. coli LdcI and LdcC shows that some amino acid residues of the C-terminal β-sheet are the same in the two proteins, whereas others are notably different in chemical nature. Importantly, most of the amino acid differences between the two enzymes are located in this very region. Thus, to advance beyond our experimental confirmation of the C-terminal β-sheet as a major determinant of the capacity of a particular lysine decarboxylase to form a cage with RavA, we set out to investigate whether certain residues in this β-sheet are conserved in lysine decarboxylases of different enterobacteria that have the ravA-viaA operon in their genome. We inspected the genetic environment of lysine decarboxylases from 22 enterobacterial species referenced in the NCBI database, corrected the gene annotation where necessary (Tables S3 and S4), and performed multiple sequence alignment coupled to a phylogenetic analysis (see Methods). This procedure yielded several unexpected and exciting results. First of all, consensus sequence for the entire lysine decarboxylase family was derived. Second, the phylogenetic analysis clearly split the lysine decarboxylases into two groups (Fig. 6A). All lysine decarboxylases predicted to be “LdcI-like” or biodegradable based on their genetic environment, as for example their organization in an operon with a gene encoding the CadB antiporter (see Methods), were found in one group, whereas all enzymes predicted as “LdcC-like” or biosynthetic partitioned into another group. Thus, consensus sequences could also be determined for each of the two groups (Figs 6B,C and S7). Inspection of these consensus sequences revealed important differences between the groups regarding charge, size and hydrophobicity of several residues precisely at the level of the C-terminal β-sheet that is responsible for the interaction with RavA (Fig. 6B–D). For example, in our previous study, site-directed mutations identified Y697 as critically required for the RavA binding. Our current analysis shows that Y697 is strictly conserved in the “LdcI-like” group whereas the “LdcC-like” enzymes always have a lysine in this position; it also uncovers several other residues potentially essential for the interaction with RavA which can now be addressed by site-directed mutagenesis. The third and most remarkable finding was that exactly the same separation into “LdcI-like” and “LdcC”-like groups can be obtained based on a comparison of the C-terminal β-sheets only, without taking the rest of the primary sequence into account. Therefore the C-terminal β-sheet emerges as being a highly conserved signature sequence, sufficient to unambiguously discriminate between the “LdcI-like” and “LdcC-like” enterobacterial lysine decarboxylases independently of any other information (Figs 6 and S7). Our structures show that this motif is not involved in the enzymatic activity or the oligomeric state of the proteins. Thus, enterobacteria identified here (Fig. 6, Table S4) appear to exert evolutionary pressure on the biodegradative lysine decarboxylase towards the RavA binding. One of the elucidated roles of the LdcI-RavA cage is to maintain LdcI activity under conditions of enterobacterial starvation by preventing LdcI inhibition by the stringent response alarmone ppGpp. Furthermore, the recently documented interaction of both LdcI and RavA with specific subunits of the respiratory complex I, together with the unanticipated link between RavA and maturation of numerous iron-sulfur proteins, tend to suggest an additional intriguing function for this 3.5 MDa assembly. The conformational rearrangements of LdcI upon enzyme activation and RavA binding revealed in this work, and our amazing finding that the molecular determinant of the LdcI-RavA interaction is the one that straightforwardly determines if a particular enterobacterial lysine decarboxylase belongs to “LdcI-like” or “LdcC-like” proteins, should give a new impetus to functional studies of the unique LdcI-RavA cage. Besides, the structures and the pseudoatomic models of the active ppGpp-free states of both the biodegradative and the biosynthetic E. coli lysine decarboxylases offer an additional tool for analysis of their role in UPEC infectivity. Together with the apo-LdcI and ppGpp-LdcIi crystal structures, our cryoEM reconstructions provide a structural framework for future studies of structure-function relationships of lysine decarboxylases from other enterobacteria and even of their homologues outside Enterobacteriaceae. For example, the lysine decarboxylase of Eikenella corrodens is thought to play a major role in the periodontal disease and its inhibitors were shown to retard gingivitis development. Finally, cadaverine being an important platform chemical for the production of industrial polymers such as nylon, structural information is valuable for optimisation of bacterial lysine decarboxylases used for its production in biotechnology. Methods Protein expression and purification LdcI and LdcC were expressed and purified as described from an E. coli strain that cannot produce ppGpp (MG1655 ΔrelA ΔspoT strain). LdcI was stored in 20 mM Tris-HCl, 100 mM NaCl, 1 mM DTT, 0.1 mM PLP, pH 6.8 (buffer A) and LdcC in 20 mM Tris-HCl, 100 mM NaCl, 1 mM DTT, 0.1 mM PLP, pH 7.5 (buffer B). Chimeric LdcIC and LdcCI were constructed, expressed and purified as follows. The chimeras were designed by exchange, between LdcI and LdcC, of residues from 631 to 640 and from 697 to the C-terminus, corresponding to the regions around the two strands of the C-terminal β-sheet (Fig. 5B,C), while leaving the rest of the sequence unaltered. The synthetic ldcIC and ldcCI genes (2148 bp and 2154 bp respectively), provided within a pUC57 vector (GenScript) were subcloned into pET-TEV vector based on pET-28a (Invitrogen) containing an N-terminal TEV-cleavable 6x-His-Tag. Proteins were expressed in Rosetta 2 (DE3) cells (Novagen) in LB medium supplemented with kanamycin and chloramphenicol at 37 °C, upon induction with 0.5 mM IPTG at 18 °C. Cells were harvested by centrifugation, the pellet resuspended in a 50 mM Tris-HCl, 150 mM NaCl, pH 8 buffer supplemented with Complete EDTA free (Roche) and 0.1 mM PMSF (Sigma), and disrupted by sonication at 4 °C. After centrifugation at 75000 g at 4 °C for 20 min, the supernatant was loaded on a Ni-NTA column. The eluted protein-containing fractions were pooled and the His-Tag removed by incubation with the TEV protease at 1/100 ratio and an extensive dialysis in a 50 mM Tris-HCl, 150 mM NaCl, 1 mM DTT, 5 mM EDTA, pH 8 buffer. After a second dialysis in a 50 mM Tris-HCl, 150 mM NaCl, pH 8 buffer supplemented with 10 mM imidazole, the sample was loaded on a Ni-NTA column in the same buffer, which allowed to separate the TEV protease and LdcCI/LdcIC. Finally, the pure proteins were obtained by size exclusion chromatography on a Superdex-S200 column in buffer A. LdcIa -cryoEM data collection and 3D reconstruction LdcI was prepared at 2 mg/ml in a buffer containing 25 mM MES, 100 mM NaCl, 0.2 mM PLP, 1 mM DTT, pH 6.2. 3 μl of sample were applied to glow-discharged quantifoil grids 300 mesh 2/1 (Quantifoil Micro Tools GmbH, Germany), excess solution was blotted during 2.5 s with a Vitrobot (FEI) and the grid frozen in liquid ethane. Data collection was performed on a FEI Polara microscope operated at 300 kV under low dose conditions. Micrographs were recorded on Kodak SO-163 film at 59,000 magnification, with defocus ranging from 0.6 to 4.9 μm. Films were digitized on a Zeiss scanner (Photoscan) at a step size of 7 μm giving a pixel size of 1.186 Å. The contrast transfer function (CTF) for each micrograph was determined with CTFFIND3. Initially ~2500 particles of 256 × 256 pixels were extracted manually, binned 4 times and subjected to one round of multivariate statistical analysis and classification using IMAGIC. Representative class averages corresponding to projections in different orientations were used as input for an ab-initio 3D reconstruction by RICOserver (rico.ibs.fr/). The resulting 3D reconstruction was refined using RELION, which yielded an 18 Å resolution map. Using projections of this model as a template, particles of size 256 × 256 pixels were semi-automatically selected from all the micrographs using the Fast Projection Matching (FPM) algorithm. The resulting dataset of ~46000 particles was processed in RELION with the previous map as an initial model and with a full CTF correction after the first peak. The final map comprised 44207 particles with a resolution of 7.4 Å as per the gold-standard FSC = 0.143 criterion. It was sharpened with EMBfactor using calculated B-factor of −350 Å2 and masked with a soft mask to obtain a final map with a resolution of 6.1 Å (Fig. S3, Table S1). LdcC - cryoEM data collection and 3D reconstruction LdcC was prepared at 2 mg/ml in a buffer containing 25 mM HEPES, 100 mM NaCl, 0.2 mM PLP, 1 mM DTT, pH 7.2. Grids were prepared and sample imaged as LdcIa. Data were processed essentially as LdcIa described above. Briefly, an initial ~20 Å resolution model was generated by angular reconstitution after manual picking of circa 3000 particles from the first micrographs, filtered to 60 Å resolution, refined with RELION and used for a semi-automatic selection with FPM. The dataset was processed in RELION with a full CTF correction to yield a final reconstruction comprising 61000 particles. The map was sharpened with Relion postprocessing, using a soft mask and automated B-factor estimation (−690 Å2), yielding a map at 5.5 Å resolution (Fig. S1, Table S1). LdcI-LARA - 3D reconstruction In our first study, the dataset was processed in SPIDER and the CTF correction involved a simple phase-flipping. Therefore, for consistency with the present work, we revisited the dataset and processed it in RELION with a full CTF correction after the first peak. It was sharpened with EMBfactor using calculated B-factor of −350 Å2 and masked with a soft mask to obtain a final map with a resolution of 6.2 Å (Fig. S2). This reconstruction is of a slightly better quality in terms of the continuity of the internal density. Therefore we used this improved map for fitting of the atomic model and further analysis (Fig. S2, Table S1). Additional image processing As a crosscheck, each data set was also refined either from other initial models: a “featureless donut” with approximate dimensions of the decamer, and low pass-filtered reconstructions from the two other data sets (i.e. the LdcC reconstruction was used as a model for the LdcIa and LdcI-LARA data sets, etc). All refinements converged to the same solutions independently of the starting model. Local resolution of all maps was determined with ResMap. LdcCI and LdcIC chimeras —negative stain EM and 2D image analysis 0.4 mg/ml of RavA (in a 20 mM Tris-HCl, 500 mM NaCl, 10 mM MgCl2, 1 mM DTT, 5% glycerol, pH 6.8 buffer) was mixed with 0.3 mg/ml of either LdcI, LdcC, LdcCI or LdcIC in the presence of 2 mM ADP and 10 mM MgCl2 in a buffer containing 20 mM Hepes and 150 mM NaCl at pH 7.4. After 10 minutes incubation at room temperature, 3 μl of mixture were applied to the clear side of the carbon on a carbon-mica interface and negatively stained with 2% uranyl acetate. Images were recorded with a JEOL 1200 EX II microscope at 100 kV at a nominal magnification of 15000 on a CCD camera yielding a pixel size of 4.667 Å. No complexes between RavA and LdcC or LdcIC could be observed, whereas the LdcCI-RavA preparation manifested cage-like particles similar to the previously published LdcI-RavA, but also unbound RavA and LdcCI, which implies that the affinity of RavA to the LdcCI chimera is lower than its affinity to the native LdcI. 1260 particles of 96 × 96 pixels were extracted interactively from several micrographs. 2D centering, multivariate statistical analysis and classification were performed using IMAGIC. Class-averages similar to the cage-like LdcI-RavA complex were used as references for multi-reference alignment followed by multivariate statistical analysis and classification. Fitting of atomic models into cryoEM maps A homology model of LdcC was obtained using the atomic coordinates of the LdcI monomer (3N75) as the template in SWISS-MODEL server. The RMSD between the template and the resulting model was 0.26 Å. The LdcC model was then fitted as a rigid body into the LdcC cryoEM map using the fit-in-map module of UCSF Chimera. This rigid fit indicated movements of several parts of the protein. Therefore, the density corresponding to one LdcC monomer was extracted and flexible fitting was performed using IMODFIT at 8 Å resolution. This monomeric model was then docked into the decameric cryoEM map with URO and its graphical version VEDA that use symmetry information for fitting in Fourier space. The Cα RMSDmin between the initial model of the LdcC monomer and the final IMODFIT LdcC model is 1.2 Å. In the case of LdcIa, the density corresponding to an individual monomer was extracted and the fit performed similarly to the one described above, with the final model of the decameric particle obtained with URO and VEDA. The Cα RMSDmin between the LdcIi monomer and the final IMODFIT model is 1.4 Å. For LdcI-LARA, the density accounting for one LdcI monomer bound to a LARA domain was extracted and further separated into individual densities corresponding to LdcI and to LARA. The fit of LdcI was performed as for LdcC and LdcIa, while the crystal structure of LARA was docked into the monomeric LdcI-LARA map as a rigid body using SITUS. The resulting pseudoatomic models were used to create the final model of the LdcI-LARA decamer with URO and VEDA. The Cα RMSDmin between the LdcIi monomer and the final IMODFIT model is 1.4 Å. A brief summary of relevant parameters is provided in Table S1. Sequence analysis Out of a non-exhaustive list of 50 species of Enterobacteriaceae (Table S3), 22 were found to contain genes annotated as ldcI or ldcC and containing the ravA-viaA operon (Table S4). An analysis using MUSCLE with default parameters showed that these genes share more than 65% identity. To verify annotation of these genes, we compared their genetic environment with that of E. coli ldcI and ldcC. Indeed, in E. coli the ldcI gene is in operon with the cadB gene encoding a lysine-cadaverine antiporter, whereas the ldcC gene is present between the accA gene (encoding an acetyl-CoA carboxylase alpha subunit carboxyltransferase) and the yaeR gene (coding for an unknown protein belonging to the family of Glyoxalase/Dioxygenase/Bleomycin resistance proteins). Compared with this genetic environment, the annotation of several ldcI and ldcC genes in enterobacteria was found to be inconsistent (Table S4). For example, several strains contain genes annotated as ldcC in the genetic background of ldcI and vice versa, as in the case of Salmonella enterica and Trabulsiella guamensi. Furthermore, the gene with an “ldcC-like” environment was found to be annotated as cadA in particular strains of Citrobacter freundii, Cronobacter sakazakii, Enterobacter cloacae subsp. Cloaca, Erwinia amylovora, Pantoea agglomerans, Rahnella aquatilis, Shigella dysenteriae, and Yersinia enterocolitica subsp. enterocolitica, whereas in Hafnia alvei, Kluyvera ascorbata, and Serratia marcescens subsp. marcescens, the gene with an “ldcI-like” environment was found to be annotated as ldcC. In addition, as far as the genetic environment is concerned, Plesiomonas appears to have two ldc genes with the organization of the E. coli ldcI (operon cadA-cadB). Consequently, we corrected for gene annotation consistent with the genetic environment and made multiple sequence alignments using version 8.0.1 of the CLC Genomics Workbench software. A phylogenetic tree was generated based on Maximum Likelihood and following the Neighbor-Joining method with the WAG protein substitution model. The reliability of the generated phylogenetic tree was assessed by bootstrap analysis. The presented unrooted phylogenetic tree shows the nodes that are reliable over 95% (Fig. 6A). Remarkably, the multiple sequence alignment and the resulting phylogenetic tree clearly grouped together all sequences annotated as ldcI on the one side, and all sequences annotated as ldcC on the other side. Thus, we conclude that all modifications in gene annotations that we introduced for the sake of consistency with the genetic environment are perfectly corroborated by the multiple sequence alignment and the phylogenetic analysis. Since the regulation of the lysine decarboxylase gene (i.e. inducible or constitutive) cannot be assessed by this analysis, we call the resulting groups “ldcI-like” and “ldcC-like” as referred to the E. coli enzymes, and make a parallel between the membership in a given group and the ability of the protein to form a cage complex with RavA. Additional Information Accession codes: CryoEM maps and corresponding fitted atomic structures (main chain atoms) have been deposited in the Electron Microscopy Data Bank and Protein Data Bank, respectively, with accession codes EMD-3205 and 5FKZ for LdcC, EMD-3204 and 5FKX for LdcIa and EMD-3206 and 5FL2 for LdcI-LARA. How to cite this article: Kandiah, E. et al. Structural insights into the Escherichia coli lysine decarboxylases and molecular determinants of interaction with the AAA+ ATPase RavA. Sci. Rep. 6, 24601; doi: 10.1038/srep24601 (2016). Supplementary Material From cofactor to enzymes. The molecular evolution of pyridoxal-5′-phosphate-dependent enzymes Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations Acid stress response in enteropathogenic gammaproteobacteria: an aptitude for survival Mechanisms of acid resistance in Escherichia coli Sensing and adaptation to low pH mediated by inducible amino acid decarboxylases in Salmonella The effect of the pH of the medium during growth on the enzymic activities of bacteria (Escherichia coli and Micrococcus lysodeikticus) and the biological significance of the changes produced Studies on bacterial amino-acid decarboxylases: 1. l(+)-lysine decarboxylase The cadA gene of Vibrio cholerae is induced during infection and plays a role in acid tolerance Lysine decarboxylase expression by Vibrio vulnificus is induced by SoxR in response to superoxide stress Polyamine-mediated resistance of uropathogenic Escherichia coli to nitrosative stress Conditioning of uropathogenic Escherichia coli for enhanced colonization of host Lysine Decarboxylase Activity as a Factor of Fluoroquinolone Resistance in Escherichia coli RpoS-dependent expression of the second lysine decarboxylase gene in Escherichia coli Cadaverine inhibition of porin plays a role in cell survival at acidic pH The role of OmpC and OmpF in acidic resistance in Escherichia coli Role of polyamines in formation of multiple antibiotic resistance of Escherichia coli under stress conditions Linkage between the bacterial acid stress and stringent responses: the structure of the inducible lysine decarboxylase Purification and physical properties of inducible Escherichia coli lysine decarboxylase Formation of a distinctive complex between the inducible bacterial lysine decarboxylase and a novel AAA+ ATPase Novel structural and functional insights into the MoxR family of AAA+ ATPases Structure of RavA MoxR AAA+ protein reveals the design principles of a molecular cage modulating the inducible lysine decarboxylase activity The MoxR ATPase RavA and its cofactor ViaA interact with the NADH:ubiquinone oxidoreductase I in Escherichia coli Assembly principles of a unique cage formed by hexameric and decameric E. coli proteins The Escherichia coli ldcC gene encodes another lysine decarboxylase, probably a constitutive enzyme Crystal structure of diaminopelargonic acid synthase: evolutionary relationships between pyridoxal-5′-phosphate-dependent enzymes The enzymatic activities of the Escherichia coli basic aliphatic amino acid decarboxylases exhibit a pH zone of inhibition Disruption of individual nuo-genes leads to the formation of partially assembled NADH:ubiquinone oxidoreductase (complex I) in Escherichia coli Biofilm Lysine Decarboxylase, a New Therapeutic Target for Periodontal Inflammation Effects of immunization with natural and recombinant lysine decarboxylase on canine gingivitis development Bacterial lysine decarboxylase influences human dental biofilm lysine content, biofilm accumulation, and subclinical gingival inflammation Optimization of Direct Lysine Decarboxylase Biotransformation for Cadaverine Production with Whole-Cell Biocatalysts at High Lysine Concentration Enhanced cadaverine production from L-lysine using recombinant Escherichia coli co-overexpressing CadA and CadB Cadaverine production by heterologous expression of Klebsiella oxytoca lysine decarboxylase Cryo-electron microscopy of vitrified specimens Accurate determination of local defocus and specimen tilt in electron microscopy A new generation of the IMAGIC image processing system On the three-dimensional reconstruction of icosahedral particles RELION: implementation of a Bayesian approach to cryo-EM structure determination Ab initio high-resolution single-particle 3D reconstructions: the symmetry adapted functions way Prevention of overfitting in cryo-EM structure determination Sharpening high resolution information in single particle electron cryomicroscopy Quantifying the local resolution of cryo-EM density maps SWISS-MODEL: An automated protein homology-modeling server UCSF Chimera–a visualization system for exploratory research and analysis iMODFIT: efficient and robust flexible fitting based on vibrational analysis in internal coordinates On the fitting of model electron densities into EM reconstructions: a reciprocal-space formulation UROX 2.0: an interactive tool for fitting atomic models into electron-microscopy reconstructions MUSCLE: multiple sequence alignment with high accuracy and high throughput A general empirical model of protein evolution derived from multiple protein families using a maximum-likelihood approach Author Contributions E.K., H.M. and I.G. carried out EM data collection with assistance of M.B. and analyzed the data. D.C. performed cloning, multiple sequence alignment and phylogenetic analysis under the direction of S.E. and I.G., J.P. cloned and purified chimeric proteins under the direction of S.O.C., K.L. and S.W.S.C. purified LdcI, LdcC and LARA under the direction of W.A.H., I.G. conceived and directed the studies and wrote the manuscript with input from E.K. 3D cryoEM reconstructions of LdcC, LdcI-LARA and LdcIa. (A,C,E) cryoEM map of the LdcC (A), LdcIa\n(C) and LdcI-LARA (E) decamers with one protomer in light\ngrey. In the rest of the protomers, the wing, core and C-terminal domains\nare colored from light to dark in shades of green for LdcC (A), pink\nfor LdcIa (C) and blue for LdcI in LdcI-LARA (E).\nIn (E), the LARA domain density is shown in dark grey. Two monomers\nmaking a dimer are delineated. Scale bar 50 Å.\n(B,D,F) One protomer from the cryoEM map of the LdcC (B),\nLdcIa (D) and LdcI-LARA (F) in light grey with\nthe pseudoatomic model represented as cartoons and colored as the densities\nin (A,C,E). Each domain is indicated for clarity. Scale bar\n50 Å. See also Figs S1 and S3. Analysis of conformational rearrangements. Superposition of the pseudoatomic models of LdcC, LdcI from LdcI-LARA and\nLdcIa colored as in Fig. 1, and the\ncrystal structure of LdcIi in shades of yellow. Only one of the\ntwo rings of the double toroid is shown for clarity. The dashed circle\nindicates the central region that remains virtually unchanged between all\nthe structures, while the periphery undergoes visible movements. Scale bar\n50 Å. Conformational rearrangements in the enzyme active site. (A) LdcIi crystal structure, with one ring represented as a\ngrey surface and the second as a cartoon. A monomer with its PLP cofactor is\ndelineated. The PLP moieties of the cartoon ring are shown in red.\n(B) The LdcIi dimer extracted from the crystal structure\nof the decamer. One monomer is colored in shades of yellow as in Figs 1 and 2, while the monomer\nrelated by C2 symmetry is grey. The PLP is red. The active site is boxed.\n(C–F) Close-up views of the active site. The PLP\nmoiety in red is from the LdcIi crystal structure. We did not\nattempt to model it in the cryoEM maps. The dimer interface is shown as a\ndashed line and the active site α-helices mentioned in the text\nare highlighted. (C) Compares LdcIi (yellow) and\nLdcIa (pink), (D) compares LdcIa (pink) and\nLdcI-LARA (blue), and (E) compares LdcIi (yellow),\nLdcIa (pink) and LdcI-LARA (blue) simultaneously in order to\nshow the progressive shift described in the text. (F) Shows the\nsimilarity between LdcIa and LdcC at the level of the secondary\nstructure elements composing the active site. Colors are as in the other\nfigures. Stretching of the LdcI monomer upon pH-dependent enzyme activation and LARA\nbinding. (A–C) A slice through the pseudoatomic models of the LdcI\nmonomers extracted from the superimposed decamers (Fig.\n2) The rectangle indicates the regions enlarged in\n(D–F). (A) compares LdcIi (yellow)\nand LdcIa (pink), (B) compares LdcIa (pink) and\nLdcI-LARA (blue), and (C) compares LdcIi (yellow),\nLdcIa (pink) and LdcI-LARA (blue) simultaneously in order to\nshow the progressive stretching described in the text. The cryoEM density of\nthe LARA domain is represented as a grey surface to show the position of the\nbinding site and the direction of the movement. (D–F)\nInserts zooming at the CTD part in proximity of the LARA binding site. Loop\nregions are removed for a clearer visual comparison. An arrow indicates a\nswinging movement. Analysis of the LdcIC and LdcCI chimeras. (A) A slice through the pseudoatomic models of the LdcIa\n(purple) and LdcC (green) monomers extracted from the superimposed decamers\n(Fig. 2). (B) The C-terminal\nβ-sheet in LdcIa and LdcC enlarged from\n(A,C) Exchanged primary sequences (capital letters) and\ntheir immediate vicinity (lower case letters) colored as in\n(A,B), with the corresponding secondary structure elements\nand the amino acid numbering shown. (D,E) A gallery of negative stain\nEM images of (D) the wild type LdcI-RavA cage and (E) the\nLdcCI-RavA cage-like particles. (F) Some representative class\naverages of the LdcCI-RavA cage-like particles. Scale bar\n20 nm. Sequence analysis of enterobacterial lysine decarboxylases. (A) Maximum likelihood tree with the\n“LdcC-like” and the\n“LdcI-like” groups highlighted in green and pink,\nrespectively. Only nodes with higher values than 95% are shown (500\nreplicates of the original dataset, see Methods for details). Scale bar\nindicates the average number of substitutions per site. (B) Analysis\nof consensus “LdcI-like” and\n“LdcC-like” sequences around the first and second\nC-terminal β-strands. The height of the bars and the letters\nrepresenting the amino acids reflects the degree of conservation of each\nparticular position is in the alignment. Amino acids are colored according\nto a polarity color scheme with hydrophobic residues in black, hydrophilic\nin green, acidic in red and basic in blue. Numbering as in E. coli.\n(C) Signature sequences of LdcI and LdcC in the C-terminal\nβ-sheet. Polarity differences are highlighted. (D)\nPosition and nature of these differences at the surface of the respective\ncryoEM maps with the color code as in B. See also Fig. S7 and Tables S3 and 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diff --git a/annotated_BioC_JSON/PMC4833862_ann.json b/annotated_BioC_JSON/PMC4833862_ann.json
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@@ -0,0 +1 @@
+[{"sourceid":"4833862","sourcedb":"","project":"","target":"","text":"The dynamic organization of fungal acetyl-CoA carboxylase Acetyl-CoA carboxylases (ACCs) catalyse the committed step in fatty-acid biosynthesis: the ATP-dependent carboxylation of acetyl-CoA to malonyl-CoA. They are important regulatory hubs for metabolic control and relevant drug targets for the treatment of the metabolic syndrome and cancer. Eukaryotic ACCs are single-chain multienzymes characterized by a large, non-catalytic central domain (CD), whose role in ACC regulation remains poorly characterized. Here we report the crystal structure of the yeast ACC CD, revealing a unique four-domain organization. A regulatory loop, which is phosphorylated at the key functional phosphorylation site of fungal ACC, wedges into a crevice between two domains of CD. Combining the yeast CD structure with intermediate and low-resolution data of larger fragments up to intact ACCs provides a comprehensive characterization of the dynamic fungal ACC architecture. In contrast to related carboxylases, large-scale conformational changes are required for substrate turnover, and are mediated by the CD under phosphorylation control.  Acetyl-CoA carboxylases are central regulatory hubs of fatty acid metabolism and are important targets for drug development in obesity and cancer. Here, the authors demonstrate that the regulation of these highly dynamic enzymes in fungi is governed by a mechanism based on phosphorylation-dependent conformational variability. Biotin-dependent acetyl-CoA carboxylases (ACCs) are essential enzymes that catalyse the ATP-dependent carboxylation of acetyl-CoA to malonyl-CoA. This reaction provides the committed activated substrate for the biosynthesis of fatty acids via fatty-acid synthase. By catalysing this rate-limiting step in fatty-acid biosynthesis, ACC plays a key role in anabolic metabolism. ACC inhibition and knock-out studies show the potential of targeting ACC for treatment of the metabolic syndrome. Furthermore, elevated ACC activity is observed in malignant tumours. A direct link between ACC and cancer is provided by cancer-associated mutations in the breast cancer susceptibility gene 1 (BRCA1), which relieve inhibitory interactions of BRCA1 with ACC. Thus, ACC is a relevant drug target for type 2 diabetes and cancer. Microbial ACCs are also the principal target of antifungal and antibiotic compounds, such as Soraphen A. The principal functional protein components of ACCs have been described already in the late 1960s for Escherichia coli (E. coli) ACC: Biotin carboxylase (BC) catalyses the ATP-dependent carboxylation of a biotin moiety, which is covalently linked to the biotin carboxyl carrier protein (BCCP). Carboxyltransferase (CT) transfers the activated carboxyl group from carboxybiotin to acetyl-CoA to yield malonyl-CoA. Prokaryotic ACCs are transient assemblies of individual BC, CT and BCCP subunits. Eukaryotic ACCs, instead, are multienzymes, which integrate all functional components into a single polypeptide chain of ∼2,300 amino acids. Human ACC occurs in two closely related isoforms, ACC1 and 2, located in the cytosol and at the outer mitochondrial membrane, respectively. In addition to the canonical ACC components, eukaryotic ACCs contain two non-catalytic regions, the large central domain (CD) and the BC–CT interaction domain (BT). The CD comprises one-third of the protein and is a unique feature of eukaryotic ACCs without homologues in other proteins. The function of this domain remains poorly characterized, although phosphorylation of several serine residues in the CD regulates ACC activity. The BT domain has been visualized in bacterial carboxylases, where it mediates contacts between α- and β-subunits. Structural studies on the functional architecture of intact ACCs have been hindered by their huge size and pronounced dynamics, as well as the transient assembly mode of bacterial ACCs. However, crystal structures of individual components or domains from prokaryotic and eukaryotic ACCs, respectively, have been solved. The structure determination of the holoenzymes of bacterial biotin-dependent carboxylases, which lack the characteristic CD, such as the pyruvate carboxylase (PC), propionyl-CoA carboxylase, 3-methyl-crotonyl-CoA carboxylase and a long-chain acyl-CoA carboxylase revealed strikingly divergent architectures despite a general conservation of all functional components. In these structures, the BC and CT active sites are at distances between 40 and 80 Å, such that substrate transfer could be mediated solely by the mobility of the flexibly tethered BCCP. Human ACC1 is regulated allosterically, via specific protein–protein interactions, and by reversible phosphorylation. Dynamic polymerization of human ACC1 is linked to increased activity and is regulated allosterically by the activator citrate and the inhibitor palmitate, or by binding of the small protein MIG-12 (ref.). Human ACC1 is further regulated by specific phosphorylation-dependent binding of BRCA1 to Ser1263 in the CD. BRCA1 binds only to the phosphorylated form of ACC1 and prevents ACC activation by phosphatase-mediated dephosphorylation. Furthermore, phosphorylation by AMP-activated protein kinase (AMPK) and cAMP-dependent protein kinase (PKA) leads to a decrease in ACC1 activity. AMPK phosphorylates ACC1 in vitro at Ser80, Ser1201 and Ser1216 and PKA at Ser78 and Ser1201. However, regulatory effects on ACC1 activity are mainly mediated by phosphorylation of Ser80 and Ser1201 (refs). Phosphorylated Ser80, which is highly conserved only in higher eukaryotes, presumably binds into the Soraphen A-binding pocket. The regulatory Ser1201 shows only moderate conservation across higher eukaryotes, while the phosphorylated Ser1216 is highly conserved across all eukaryotes. However, no effect of Ser1216 phosphorylation on ACC activity has been reported in higher eukaryotes. For fungal ACC, neither spontaneous nor inducible polymerization has been detected despite considerable sequence conservation to human ACC1. The BRCA1-interacting phosphoserine position is not conserved in fungal ACC, and no other phospho-dependent protein–protein interactions of fungal ACC have been described. In yeast ACC, phosphorylation sites have been identified at Ser2, Ser735, Ser1148, Ser1157 and Ser1162 (ref.). Of these, only Ser1157 is highly conserved in fungal ACC and aligns to Ser1216 in human ACC1. Its phosphorylation by the AMPK homologue SNF1 results in strongly reduced ACC activity. Despite the outstanding relevance of ACC in primary metabolism and disease, the dynamic organization and regulation of the giant eukaryotic, and in particular fungal ACC, remain poorly characterized. Here we provide the structure of Saccharomyces cerevisiae (Sce) ACC CD, intermediate- and low-resolution structures of human (Hsa) ACC CD and larger fragments of fungal ACC from Chaetomium thermophilum (Cth; Fig. 1a). Integrating these data with small-angle X-ray scattering (SAXS) and electron microscopy (EM) observations yield a comprehensive representation of the dynamic structure and regulation of fungal ACC. Results The organization of the yeast ACC CD First, we focused on structure determination of the 82-kDa CD. The crystal structure of the CD of SceACC (SceCD) was determined at 3.0 Å resolution by experimental phasing and refined to Rwork/Rfree=0.20/0.24 (Table 1). The overall extent of the SceCD is 70 by 75 Å (Fig. 1b and Supplementary Fig. 1a,b), and the attachment points of the N-terminal 26-residue linker to the BCCP domain and the C-terminal CT domain are separated by 46 Å (the N- and C termini are indicated with spheres in Fig. 1b). SceCD comprises four distinct domains, an N-terminal α-helical domain (CDN), and a central four-helix bundle linker domain (CDL), followed by two α–β-fold C-terminal domains (CDC1/CDC2). CDN adopts a letter C shape, where one of the ends is a regular four-helix bundle (Nα3-6), the other end is a helical hairpin (Nα8,9) and the bridging region comprises six helices (Nα1,2,7,10–12). CDL is composed of a small, irregular four-helix bundle (Lα1–4) and tightly interacts with the open face of CDC1 via an interface of 1,300 Å2 involving helices Lα3 and Lα4. CDL does not interact with CDN apart from the covalent linkage and forms only a small contact to CDC2 via a loop between Lα2/α3 and the N-terminal end of Lα1, with an interface area of 400 Å2. CDC1/CDC2 share a common fold; they are composed of six-stranded β-sheets flanked on one side by two long, bent helices inserted between strands β3/β4 and β4/β5. CDC2 is extended at its C terminus by an additional β-strand and an irregular β-hairpin. On the basis of a root mean square deviation of main chain atom positions of 2.2 Å, CDC1/CDC2 are structurally more closely related to each other than to any other protein (Fig. 1c); they may thus have evolved by duplication. Close structural homologues could not be found for the CDN or the CDC domains. A regulatory loop mediates interdomain interactions To define the functional state of insect-cell-expressed ACC variants, we employed mass spectrometry (MS) for phosphorylation site detection. In insect-cell-expressed full-length SceACC, the highly conserved Ser1157 is the only fully occupied phosphorylation site with functional relevance in S. cerevisiae. Additional phosphorylation was detected for Ser2101 and Tyr2179; however, these sites are neither conserved across fungal ACC nor natively phosphorylated in yeast. MS analysis of dissolved crystals confirmed the phosphorylated state of Ser1157 also in SceCD crystals. The SceCD structure thus authentically represents the state of SceACC, where the enzyme is inhibited by SNF1-dependent phosphorylation. In the SceCD crystal structure, the phosphorylated Ser1157 resides in a regulatory 36-amino-acid loop between strands β2 and β3 of CDC1 (Fig. 1b,d), which contains two additional less-conserved phosphorylation sites (Ser1148 and Ser1162) confirmed in yeast, but not occupied here. This regulatory loop wedges between the CDC1 and CDC2 domains and provides the largest contribution to the interdomain interface. The N-terminal region of the regulatory loop also directly contacts the C-terminal region of CDC2 leading into CT. Phosphoserine 1157 is tightly bound by two highly conserved arginines (Arg1173 and Arg1260) of CDC1 (Fig. 1d). Already the binding of phosphorylated Ser1157 apparently stabilizes the regulatory loop conformation; the accessory phosphorylation sites Ser1148 and Ser1162 in the same loop may further modulate the strength of interaction between the regulatory loop and the CDC1 and CDC2 domains. Phosphorylation of the regulatory loop thus determines interdomain interactions of CDC1 and CDC2, suggesting that it may exert its regulatory function by modifying the overall structure and dynamics of the CD. The functional role of Ser1157 was confirmed by an activity assay based on the incorporation of radioactive carbonate into acid non-volatile material. Phosphorylated SceACC shows only residual activity (kcat=0.4±0.2 s−1, s.d. based on five replicate measurements), which increases 16-fold (kcat=6.5±0.3 s−1) after dephosphorylation with λ protein phosphatase. The values obtained for dephosphorylated SceACC are comparable to earlier measurements of non-phosphorylated yeast ACC expressed in E. coli. The variable CD is conserved between yeast and human To compare the organization of fungal and human ACC CD, we determined the structure of a human ACC1 fragment that comprises the BT and CD domains (HsaBT-CD), but lacks the mobile BCCP in between (Fig. 1a). An experimentally phased map was obtained at 3.7 Å resolution for a cadmium-derivatized crystal and was interpreted by a poly-alanine model (Fig. 1e and Table 1). Each of the four CD domains in HsaBT-CD individually resembles the corresponding SceCD domain; however, human and yeast CDs exhibit distinct overall structures. In agreement with their tight interaction in SceCD, the relative spatial arrangement of CDL and CDC1 is preserved in HsaBT-CD, but the human CDL/CDC1 didomain is tilted by 30° based on a superposition of human and yeast CDC2 (Supplementary Fig. 1c). As a result, the N terminus of CDL at helix Lα1, which connects to CDN, is shifted by 12 Å. Remarkably, CDN of HsaBT-CD adopts a completely different orientation compared with SceCD. With CDL/CDC1 superposed, CDN in HsaBT-CD is rotated by 160° around a hinge at the connection of CDN/CDL (Supplementary Fig. 1d). This rotation displaces the N terminus of CDN in HsaBT-CD by 51 Å compared with SceCD, resulting in a separation of the attachment points of the N-terminal linker to the BCCP domain and the C-terminal CT domain by 67 Å (the attachment points are indicated with spheres in Fig. 1e). The BT domain of HsaBT-CD consists of a helix that is surrounded at its N terminus by an antiparallel eight-stranded β-barrel. It resembles the BT of propionyl-CoA carboxylase; only the four C-terminal strands of the β-barrel are slightly tilted. On the basis of MS analysis of insect-cell-expressed human full-length ACC, Ser80 shows the highest degree of phosphorylation (90%). Ser29 and Ser1263, implicated in insulin-dependent phosphorylation and BRCA1 binding, respectively, are phosphorylated at intermediate levels (40%). The highly conserved Ser1216 (corresponding to S. cerevisiae Ser1157), as well as Ser1201, both in the regulatory loop discussed above, are not phosphorylated. However, residual phosphorylation levels were detected for Ser1204 (7%) and Ser1218 (7%) in the same loop. MS analysis of the HsaBT-CD crystallization sample reveals partial proteolytic digestion of the regulatory loop. Accordingly, most of this loop is not represented in the HsaBT-CD crystal structure. The absence of the regulatory loop might be linked to the less-restrained interface of CDL/CDC1 and CDC2 and altered relative orientations of these domains. Besides the regulatory loop, also the phosphopeptide target region for BRCA1 interaction is not resolved presumably because of pronounced flexibility. At the level of isolated yeast and human CD, the structural analysis indicates the presence of at least two hinges, one with large-scale flexibility at the CDN/CDL connection, and one with tunable plasticity between CDL/CDC1 and CDC2, plausibly affected by phosphorylation in the regulatory loop region. The integration of CD into the fungal ACC multienzyme To further obtain insights into the functional architecture of fungal ACC, we characterized larger multidomain fragments up to the intact enzymes. Using molecular replacement based on fungal ACC CD and CT models, we obtained structures of a variant comprising CthCT and CDC1/CDC2 in two crystal forms at resolutions of 3.6 and 4.5 Å (CthCD-CTCter1/2), respectively, as well as of a CthCT linked to the entire CD at 7.2 Å resolution (CthCD-CT; Figs 1a and 2, Table 1). No crystals diffracting to sufficient resolution were obtained for larger BC-containing fragments, or for full-length Cth or SceACC. To improve crystallizability, we generated ΔBCCP variants of full-length ACC, which, based on SAXS analysis, preserve properties of intact ACC (Supplementary Table 1 and Supplementary Fig. 2a–c). For CthΔBCCP, crystals diffracting to 8.4 Å resolution were obtained. However, molecular replacement did not reveal a unique positioning of the BC domain. Owing to the limited resolution the discussion of structures of CthCD-CT and CthΔBCCP is restricted to the analysis of domain localization. Still, these structures contribute considerably to the visualization of an intrinsically dynamic fungal ACC. In all these crystal structures, the CT domains build a canonical head-to-tail dimer, with active sites formed by contributions from both protomers (Fig. 2 and Supplementary Fig. 3a). The connection of CD and CT is provided by a 10-residue peptide stretch, which links the N terminus of CT to the irregular β-hairpin/β-strand extension of CDC2 (Supplementary Fig. 3b). The connecting region is remarkably similar in isolated CD and CthCD-CTCter structures, indicating inherent conformational stability. CD/CT contacts are only formed in direct vicinity of the covalent linkage and involve the β-hairpin extension of CDC2 as well as the loop between strands β2/β3 of the CT N-lobe, which contains a conserved RxxGxN motif. The neighbouring loop on the CT side (between CT β1/β2) is displaced by 2.5 Å compared to isolated CT structures (Supplementary Fig. 3c). On the basis of an interface area of ∼600 Å2 and its edge-to-edge connection characteristics, the interface between CT and CD might be classified as conformationally variable. Indeed, the comparison of the positioning of eight instances of the C-terminal part of CD relative to CT in crystal structures determined here, reveals flexible interdomain linking (Fig. 3a). The CDC2/CT interface acts as a true hinge with observed rotation up to 16°, which results in a translocation of the distal end of CDC2 by 8 Å. The interface between CDC2 and CDL/CDC1, which is mediated by the phosphorylated regulatory loop in the SceCD structure, is less variable than the CD–CT junction, and permits only limited rotation and tilting (Fig. 3b). Analysis of the impact of phosphorylation on the interface between CDC2 and CDL/CDC1 in CthACC variant structures is precluded by the limited crystallographic resolution. However, MS analysis of CthCD-CT and CthΔBCCP constructs revealed between 60 and 70% phosphorylation of Ser1170 (corresponding to SceACC Ser1157). The CDN domain positioning relative to CDL/CDC1 is highly variable with three main orientations observed in the structures of SceCD and the larger CthACC fragments: CDN tilts, resulting in a displacement of its N terminus by 23 Å (Fig. 4a, observed in both protomers of CthCD-CT and one protomer of CthΔBCCP, denoted as CthCD-CT1/2 and CthΔBCCP1, respectively). In addition, CDN can rotate around hinges in the connection between CDN/CDL by 70° (Fig. 4b, observed in the second protomer of CthΔBCCP, denoted as CthΔBCCP2) and 160° (Fig. 4c, observed in SceCD) leading to displacement of the anchor site for the BCCP linker by up to 33 and 40 Å, respectively. Conformational variability in the CD thus contributes considerably to variations in the spacing between the BC and CT domains, and may extend to distance variations beyond the mobility range of the flexibly tethered BCCP. On the basis of the occurrence of related conformational changes between fungal and human ACC fragments, the observed set of conformations may well represent general states present in all eukaryotic ACCs. Large-scale conformational variability of fungal ACC To obtain a comprehensive view of fungal ACC dynamics in solution, we employed SAXS and EM. SAXS analysis of CthACC agrees with a dimeric state and an elongated shape with a maximum extent of 350 Å (Supplementary Table 1). The smooth appearance of scattering curves and derived distance distributions might indicate substantial interdomain flexibility (Supplementary Fig. 2a–c). Direct observation of individual full-length CthACC particles, according to MS results predominantly in a phosphorylated low-activity state, in negative stain EM reveals a large set of conformations from rod-like extended to U-shaped particles. Class averages, obtained by maximum-likelihood-based two-dimensional (2D) classification, are focused on the dimeric CT domain and the full BC–BCCP–CD domain of only one protomer, due to the non-coordinated motions of the lateral BC/CD regions relative to the CT dimer. They identify the connections between CDN/CDL and between CDC2/CT as major contributors to conformational heterogeneity (Supplementary Fig. 4a,b). The flexibility in the CDC2/CT hinge appears substantially larger than the variations observed in the set of crystal structures. The BC domain is not completely disordered, but laterally attached to BT/CDN in a generally conserved position, albeit with increased flexibility. Surprisingly, in both the linear and U-shaped conformations, the approximate distances between the BC and CT active sites would remain larger than 110 Å. These observed distances are considerably larger than in static structures of any other related biotin-dependent carboxylase. Furthermore, based on an average length of the BCCP–CD linker in fungal ACC of 26 amino acids, mobility of the BCCP alone would not be sufficient to bridge the active sites of BC and CT. Consequently, increased flexibility or additional modes of conformational changes may be required for productive catalysis. The most relevant candidate site for mediating such additional flexibility and permitting an extended set of conformations is the CDC1/CDC2 interface, which is rigidified by the Ser1157-phosphorylated regulatory loop, as depicted in the SceCD crystal structure. Discussion Altogether, the architecture of fungal ACC is based on the central dimeric CT domain (Fig. 4d). The CD consists of four distinct subdomains and acts as a tether from the CT to the mobile BCCP and an oriented BC domain. The CD has no direct role in substrate recognition or catalysis but contributes to the regulation of all eukaryotic ACCs. In higher eukaryotic ACCs, regulation via phosphorylation is achieved by combining the effects of phosphorylation at Ser80, Ser1201 and Ser1263. In fungal ACC, however, Ser1157 in the regulatory loop of the CD is the only phosphorylation site that has been demonstrated to be both phosphorylated in vivo and involved in the regulation of ACC activity. In its phosphorylated state, the regulatory loop containing Ser1157 wedges between CDC1/CDC2 and presumably limits the conformational freedom at this interdomain interface. However, flexibility at this hinge may be required for full ACC activity, as the distances between the BCCP anchor points and the active sites of BC and CT observed here are such large that mobility of the BCCP alone is not sufficient for substrate transfer. The current data thus suggest that regulation of fungal ACC is mediated by controlling the dynamics of the unique CD, rather than directly affecting catalytic turnover at the active sites of BC and CT. A comparison between fungal and human ACC will help to further discriminate mechanistic differences that contribute to the extended control and polymerization of human ACC. Most recently, a crystal structure of near full-length non-phosphorylated ACC from S. cerevisae (lacking only 21 N-terminal amino acids, here denoted as flACC) was published by Wei and Tong. In flACC, the ACC dimer obeys twofold symmetry and assembles in a triangular architecture with dimeric BC domains (Supplementary Fig. 5a). In their study, mutational data indicate a requirement for BC dimerization for catalytic activity. The transition from the elongated open shape, observed in our experiments, towards a compact triangular shape is based on an intricate interplay of several hinge-bending motions in the CD (Fig. 4d). Comparison of flACC with our CthΔBCCP structure reveals the CDC2/CT hinge as a major contributor to conformational flexibility (Supplementary Fig. 5b,c). In flACC, CDC2 rotates ∼120° with respect to the CT domain. A second hinge can be identified between CDC1/CDC2. On the basis of a superposition of CDC2, CDC1 of the phosphorylated SceCD is rotated by 30° relative to CDC1 of the non-phosphorylated flACC (Supplementary Fig. 5d), similar to what we have observed for the non-phosphorylated HsaBT-CD (Supplementary Fig. 1d). When inspecting all individual protomer and fragment structures in their study, Wei and Tong also identify the CDN/CDC1 connection as a highly flexible hinge, in agreement with our observations. The only bona fide regulatory phophorylation site of fungal ACC in the regulatory loop is directly participating in CDC1/CDC2 domain interactions and thus stabilizes the hinge conformation. In flACC, the regulatory loop is mostly disordered, illustrating the increased flexibility due to the absence of the phosphoryl group. Only in three out of eight observed protomers a short peptide stretch (including Ser1157) was modelled. In those instances the Ser1157 residue is located at a distance of 14–20 Å away from the location of the phosphorylated serine observed here, based on superposition of either CDC1 or CDC2. Applying the conformation of the CDC1/CDC2 hinge observed in SceCD on flACC leads to CDN sterically clashing with CDC2 and BT/CDN clashing with CT (Supplementary Fig. 6a,b). Thus, in accordance with the results presented here, phosphorylation of Ser1157 in SceACC most likely limits flexibility in the CDC1/CDC2 hinge such that activation through BC dimerization is not possible (Fig. 4d), which however does not exclude intermolecular dimerization. In addition, EM micrographs of phosphorylated and dephosphorylated SceACC display for both samples mainly elongated and U-shaped conformations and reveal no apparent differences in particle shape distributions (Supplementary Fig. 7). This implicates that the triangular shape with dimeric BC domains has a low population also in the active form, even though a biasing influence of grid preparation cannot be excluded completely. Large-scale conformational variability has also been observed in most other carrier protein-based multienzymes, including polyketide and fatty-acid synthases (with the exception of fungal-type fatty-acid synthases), non-ribosomal peptide synthetases and the pyruvate dehydrogenase complexes, although based on completely different architectures. Together, this structural information suggests that variable carrier protein tethering is not sufficient for efficient substrate transfer and catalysis in any of these systems. The determination of a set of crystal structures of SceACC in two states, unphosphorylated and phosphorylated at the major regulatory site Ser1157, provides a unique depiction of multienzyme regulation by post-translational modification (Fig. 4d). The phosphorylated regulatory loop binds to an allosteric site at the interface of two non-catalytic domains and restricts conformational freedom at several hinges in the dynamic ACC. It disfavours the adoption of a rare, compact conformation, in which intramolecular dimerization of the BC domains results in catalytic turnover. The regulation of activity thus results from restrained large-scale conformational dynamics rather than a direct or indirect influence on active site structure. To our best knowledge, ACC is the first multienzyme for which such a phosphorylation-dependent mechanical control mechanism has been visualized. However, the example of ACC now demonstrates the possibility of regulating activity by controlled dynamics of non-enzymatic linker regions also in other families of carrier-dependent multienzymes. Understanding such structural and dynamic constraints imposed by scaffolding and linking in carrier protein-based multienzyme systems is a critical prerequisite for engineering of efficient biosynthetic assembly lines. Methods Protein expression and purification All proteins were expressed in the Baculovirus Expression Vector System. The MultiBac insect cell expression plasmid pACEBACI (Geneva Biotech) was modified to host a GATEWAY (LifeTechnologies) cassette with an N-terminal 10xHis-tag, named pAB1GW-NH10 hereafter. Full-length HsaACC (Genebank accession #Q13085), SceACC (#Q00955) and CthACC (#G0S3L5) were cloned into pAB1GW-NH10 using GATEWAY according to the manufacturer's manual. Truncated variants were constructed by PCR amplification, digestion of the template DNA with DpnI, phosphorylation of the PCR product and religation of the linear fragment to a circular plasmid. The following constructs were used for this study: SceACC (1–2,233), CthACC (1–2,297), CthΔBCCP (1–2,297, Δ700–765), CthCD-CT (788–2,297), CthCD-CTCter (1,114–2,297), SceCD (768–1,494) and HsaBT-CD (622–1,584, Δ753–818). Bacmid and virus production was carried out according to MultiBac instructions. Baculovirus generation and amplification as well as protein expression were performed in Sf21 cells (Expression Systems) in Insect-Xpress medium (Lonza). The cells were harvested 68–96 h post infection by centrifugation and stored at −80 °C until being processed. Cells were lysed by sonication and the lysate was cleared by ultracentrifugation. Soluble protein was purified using Ni-NTA (Genscript) and size exclusion chromatography (Superose 6, GE Healthcare). The affinity tag was removed by tobacco etch virus (TEV) protease cleavage overnight at 4 °C. TEV protease and uncleaved protein were removed by orthogonal Ni-NTA purification before size exclusion chromatography. SceACC, CthACC and CthΔBCCP were further purified by high-resolution anion exchange chromatography before size exclusion chromatography. Purified SceCD, CthCD-CTCter, CthCD-CT, CthΔBCCP, CthACC and SceACC were concentrated to 10 mg ml−1 in 30 mM 3-(N-morpholino) propanesulfonic acid (MOPS) pH 7, 200 mM ammonium sulfate, 5% glycerol and 10 mM dithiothreitol. Purified HsaBT-CD was concentrated to 20 mg ml−1 in 20 mM bicine pH 8.0, 200 mM NaCl, 5% glycerol and 5 mM tris(2-carboxyethyl) phosphine (TCEP). Proteins were used directly or were stored at −80 °C after flash-freezing in liquid nitrogen. Protein crystallization All crystallization experiments were conducted using sitting drop vapour diffusion. SceCD crystals were grown at 19 °C by mixing protein and reservoir solution (0.1 M BisTrisPropane pH 6.5, 0.05–0.2 M di-sodium malonate, 20–30% polyethylene glycol (PEG) 3350, 10 mM trimethylamine or 2% benzamidine) in a 1:1 or 2:1 ratio. Crystals appeared after several days and continued to grow for 20–200 days. Crystals were cryoprotected by short incubation in mother liquor supplemented with 22% ethylene glycol and flash-cooled in liquid nitrogen. For heavy metal derivatization the crystals were incubated in stabilization solution supplemented with 1 mM Thimerosal or 10 mM EuCl2, and then backsoaked for 15 s in stabilization solution without heavy metal. Initial crystals of HsaBT-CD grew in 0.1 M Tris pH 8.5, 0.35 M tri-potassium citrate and 2–3.5% PEG10000 at 19 °C. After several rounds of optimization, good-quality diffraction crystals were obtained at 19 °C in 0.1 M MES pH 6, 0.25–0.35 M tri-potassium citrate, 2–5% PEG10000 and 0.01–0.04 M cadmium chloride. The protein drop contained a 1:1 ratio of protein and reservoir solution. Crystals grew immediately and stopped growing after 3 days. They were dehydrated and cryoprotected in several steps in artificial mother liquor containing incrementally increasing concentrations of tri-potassium citrate, PEG10000 and ethylene glycol and then flash-cooled in liquid nitrogen. The final solution was composed of 0.1 M MES pH 6, 0.5 M tri-potassium citrate, 6.75% PEG10000, 0.01 M cadmium chloride and 22% ethylene glycol. CthCD-CTCter crystals were grown at 19 °C by mixing protein and reservoir solution (0.1 M HEPES pH 7.5, 2–7% Tacsimate pH 7, 7.5–15% PEG monomethyl ether 5000) in a 1:1 ratio. Crystals appeared after several days and continued to grow for up to 2 weeks. Crystals were cryoprotected by short incubation in mother liquor supplemented with 22% ethylene glycol. CthCD-CT ACC crystals were grown at 19 °C by mixing protein and reservoir solution (0.1 M Bicine pH 8.5–9.5, 4–8% PEG8000) in a 1:1 or 1:2 ratio. Crystals grew 8– 10 days and were cryoprotected by short incubation in mother liquor supplemented with 22% ethylene glycol before flash-cooling in liquid nitrogen. CthΔBCCP ACC crystals were grown at 19 °C by mixing protein and reservoir solution (0.1 M Morpheus buffer 3 (Molecular Dimensions, MD2-100-102), 7–12% Morpheus ethylene glycols mix (MD2-100-74), 8–12% PEG4000, 17–23% glycerol) in a 1:1 or 1:2 ratio. Crystals grew up to 3 weeks and were cryoprotected in reservoir solution before flash-cooling in liquid nitrogen. Structure determination and analysis of phosphorylation All X-ray diffraction data were collected at beamlines X06SA (PXI) or X06DA (PXIII) at the Swiss Light Source (SLS, Paul Scherrer Institute, Villigen, Switzerland) equipped with PILATUS detectors. The wavelength of data collection was 1.000 Å for native crystals, and 1.527 and 1.907 Å for crystals derivatized with europium and cadmium, respectively. Raw data were processed using XDS. Molecular replacement was carried out using Phaser 2.5.7 and 2.6.0, density modification was performed using Parrot and resolve, multicrystal averaging was carried out using phenix. All model building procedures were conducted using Coot and figures were prepared using PyMOL (Schrödinger LLC). Diffraction of initial SceCD crystals in space group P43212 with unit cell dimensions of a=b=110.3 Å and c=131.7 Å was limited to 3.5 Å. The resolution was improved to 3 Å by addition of trimethylamine or benzamidine to the reservoir solution without significant changes in unit cell dimensions. Crystals derivatized with thimerosal and europium were used for initial SAD phase determination using the SHELXC/D package. Two mercury and four europium sites were located, and an initial model was placed in the resulting maps. Since crystals derivatized with europium were slightly non-isomorphous with a c axis length of 127 Å, multicrystal averaging was used for density modification and provided directly interpretable maps. Iterative cycles of model building and refinement in Buster (version 2.10.2; Global Phasing Ltd) converged at Rwork/Rfree of 0.20/0.24. The final model lacks the disordered N terminus (amino acids 768–789), an extended loop in the CDC1 domain (1,203–1,215), a short stretch (1,147–1,149) preceding the regulatory loop and the two very C-terminal residues (1,493–1,494). On the basis of temperature factor analysis, the start and end of the regulatory loop show higher disorder than the region around the interacting phosphoserine 1157. MS analysis of dissolved crystals detected quantitative phosphorylation of the regulatory Ser1157, as also found for full-length SceACC, and additionally albeit with much lower occurrence, phosphorylation of Ser790, Ser1137, Ser1148 and Ser1159. A modelled phosphoryl position for Ser1159 could overlap with the one of Ser1157, and might be represented in the crystal. For all other phosphorylation sites no difference density could be observed, probably because of very low occupancy. PDBeFold was used to search for structural homologues. The thresholds for lowest acceptable percentage of matched secondary structure elements were 70% for the search query and 20% for the result. Initial HsaBT-CD crystals were obtained in space group I4122 with a=b=240.1 Å and c=768.9 Å and diffracted to 7.5 Å. Optimized and dehydrated crystals also belonged to space group I4122 but with unit cell parameters a=b=267.3 Å and c=210.6 Å and diffracted to a resolution of 3.7 Å. Phase information was obtained from SAD based on bound cadmium ions from the crystallization condition. Six cadmium positions were located in a 4.0-Å resolution data set at 1.9 Å wavelength using SHELXC/D via the HKL2MAP interface. Density modification and phasing based on this anomalous data set, a 3.7-Å resolution data set at 1.0 Å wavelength and additional non-isomorphous lower-resolution data sets led to a high-quality electron density map. At the intermediate resolution obtained, the map was interpreted by a poly-alanine model, which was guided by predicted secondary structure as well as sequence and structural alignment with SceCD. The final model contains five cadmium ions and refines using phenix against experimental data with Rwork/Rfree of 0.35/0.38, as expected for a poly-alanine model. Two HsaBT-CD monomers are packed in the asymmetric unit via the CDN and BT domains. Density on top of the β-barrel of one BT most likely representing parts of the BT–CD linker guided the assignment of this BT to its linked CD partner domain. This BT-to-CD assignment was further supported by the analysis of an additional lower-resolution crystal form. Cadmium ions were found to participate in crystal packing. In HsaACC, phosphorylation at regulatory sites was detected as provided in the main text. No phosphorylation was detected for other phosphosites previously identified in large-scale phosphoproteomics studies, namely serines 5, 23, 25, 48, 53, 78, 488, 786, 1273 (refs). Two different crystal forms were obtained for CthCD-CTCter (denoted as CthCD-CTCter1 and CthCD-CTCter2), diffracting to 3.6 and 4.5 Å. Both forms packed in space group P212121 with unit cell constants of a=97.7 Å, b=165.3 Å and c=219.2 Å or a=100.2 Å, b=153.5 Å and c=249.2 Å, respectively. Phases were determined by molecular replacement using a homology model based on SceCT (pdb 1od2) as search model in Phaser; multicrystal averaging was applied in density modification. The CT domain was rebuilt and an initial homology model based on the SceCD structure was fitted into difference density for CthCD-CTCter1. Iterative cycles of rebuilding and refinement in Buster converged at Rwork/Rfree of 0.20/0.24. The refined CD fragment served as a starting model for rebuilding CthCD-CTCter2 at lower resolution. Coordinate refinement in Buster was additionally guided by reference model restraints and converged at Rwork/Rfree of 0.24/0.24. Residues 1,114–1,185, 1,213–1,252, 1,380–1,385, 1,465–1,468 and 2,188–2,195 were disordered in both crystal forms and are not included in the models. Helical regions C terminal to Glu2264 of both protomers of CthCD-CTCter1 and C terminal to Leu2259 and Arg2261 of the two protomers of CthCD-CTCter2, respectively, could not be built unambiguously and were therefore interpreted by placing poly-alanine stretches. Conservation was mapped on the CthCD-CTCter1 crystal structure using al2co based on a sequence alignment of 367 fungal ACC sequences calculated by Clustal Omega. MS analysis of purified protein detected 7% phosphorylation at Ser1170 (corresponding to Ser1157 in SceCD). CthCD-CT crystallized in space group P31212 with unit cell constants of a=b=195.0 Å and c=189.5 Å and crystals diffracted to a resolution of 7.2 Å. The structure was solved by molecular replacement using a model composed of CthCT and CDC2 as search model in Phaser. CDC1 and CDN were placed manually into the resulting maps, and the model was refined using rigid-body, domain-wise TLS and B-factor refinement and NCS- and reference model-restrained coordinate refinement in Buster to Rwork/Rfree of 0.23/0.25. Owing to the low resolution, the maximum allowed B-factor in Buster refinement was increased from the default value of 300–500 Å2, minimizing B-factor clipping to 5% of all atoms. Residues 1,033–1,035, 1,134–1,152, 1,213–1,252, 1,380–1,385, 1,465–1,468 and 2,188–2,195 were not included in the models. Helical regions C terminal to Leu2259 and Arg2261 on the two protomers, respectively, were interpreted as described for CthCD-CTCter. Loop conformations, including the regulatory loop, were modelled as observed in SceCD. MS analysis of purified protein detected 60% phosphorylation at Ser1170 (corresponding to Ser1157 in SceCD). Conservation was mapped on the CthCD-CT crystal structure as for CthCD-CTCter. CthΔBCCP ACC crystallized in space group P6422 with unit cell constants of a=b=462.2 Å and c=204.6 Å, resolution was limited to 8.4 Å. Structure determination and refinement was performed as for CthCD-CT, with a maximum allowed B-factor of 500 Å2, minimizing B-factor clipping to 3% of all atoms. Although substantial difference density is observed, no defined positions of the BT and BC domains could be derived because of disorder or partial in situ proteolysis or combinations thereof. In addition, residues 1,032–1,039, 1,134–1,152, 1,213–1,252, 1,380–1,385, 1,465–1,468 and 2,188–2,195 were not included in the model. The MissingAtom macro implemented in Buster was employed to account for missing atoms, the final Rwork/Rfree were 0.30/0.32. A region C terminal to Leu2259 on one protomer was interpreted as poly-alanine. Loop conformations, including the regulatory loop, were modelled as observed in SceCD. MS analysis of purified protein detected 70% phosphorylation at Ser1170 (corresponding to Ser1157 in SceCD). Small-angle X-ray scattering Proteins were thawed on ice and dialysed overnight against 30 mM MOPS pH 7, 200 mM ammonium sulfate, 5% glycerol and 10 mM dithiothreitol. Raw scattering data were measured at SAXS beamline B21 at Diamond Light Source. The samples were measured at concentrations of 2.5, 5 and 10 mg ml−1. Data were processed using the ATSAS package according to standard procedures. A slight increase in scattering in the very low-resolution range was observed with increasing protein concentrations, which may be because of interparticle attraction or minor aggregation. Scattering intensities were thus extrapolated to zero concentration using point-wise extrapolation implemented in Primus. Direct comparison of raw scattering curves demonstrates the similarity of CthACC and CthΔBCCP, and the derived values such as Rg and Porod Volume match within expected error margins. Molecular mass estimations based on the SAXS–MOW method derive values of 534.7 and 534.0 kDa for CthACC and CthΔBCCP, respectively. The relative discrepancies to the theoretical weights of 516.8 kDa (CthACC) and 503.0 kDa (CthΔBCCP) are 3.5% and 6.2%, respectively, which is in a typical range for this method. Electron microscopy Full-length CthACC was diluted to 0.01 mg ml−1 in 30 mM MOPS pH 7.0, 200 mM ammonium sulfate, 5% glycerol and 10 mM dithiothreitol. Protein sample was adsorbed to a 200-μm copper grid and stained with 2% uranyl acetate. Grids of CthACC were imaged on a CM-200 microscope (Philips) equipped with a TVIPS F416 4k CMOS camera (Tietz Video and Image Processing Systems). The voltage used was 200 kV, and a magnification of × 50,000 results in a pixel size of 2.14 Å. Initial image processing and particle picking was carried out using Xmipp. Overall, 22,309 particles were picked semi-automatically from 236 micrographs with a box size of 300 × 300 pixels. After extraction, particles with a z-score of more than three were discarded and 22,257 particles were aligned and classified into 48 2D class averages using the maximum-likelihood target function in Fourier space (MLF2D). After 72 iterations, 4,226 additional particles were discarded and the remaining 18,031 particles were re-aligned and classified into 36 classes using MLF2D with a high-resolution cutoff of 30 Å. After 44 iterations the alignment converged and class averages were extracted. In vitro biotinylation and activity assay To ensure full functionality, SceACC was biotinylated in vitro using the E. coli biotin ligase BirA. The reaction mixture contained 10 μM ACC, 3.7 μM BirA, 50 mM Tris-HCl, pH 8, 5.5 mM MgCl2, 0.5 mM biotin, 60 mM NaCl, 3 mM ATP and 10% glycerol, and the reaction was allowed to proceed for 7 h at 30 °C. The catalytic activity of phosphorylated and dephosphorylated SceACC was measured by following the incorporation of radioactive 14C into acid-stable non-volatile material. Dephosphorylated ACC was prepared by overnight treatment with λ protein phosphatase (New England Biolabs) of partially purified ACC before the final gel filtration step. The removal of the phosphoryl group from Ser1157 was confirmed by MS. The reaction mixture contained 0.5 μg recombinant ACC in 100 mM potassium phosphate, pH 8, 3 mM ATP, 5 mM MgCl2, 50 mM NaH14CO3 (specific activity 7.4 MBq mmol−1) and 1 mM acetyl-CoA in a total reaction volume of 100 μl. The reaction mixture was incubated for 15 min at 30 °C, stopped by addition of 200 μl 6 M HCl and subsequently evaporated to dryness at 85 °C. The non-volatile residue was redissolved in 100 μl of water, 1 ml Ultima Gold XR scintillation medium (Perkin Elmer) was added and the 14C radioactivity was measured in a Packard Tricarb 2000CA liquid scintillation analyser. Measurements were carried out in five replicates and catalytic activities were calculated using a standard curve derived from measurements of varying concentrations of NaH14CO3 in reaction buffer. Additional information Accession codes: Atomic coordinates and structure factors have been deposited in the Protein Data Bank with accession codes 5I6E (SceCD), 5I87 (HsaBT-CD), 5I6F/5I6G (CthCD-CTCter1/2), 5I6H (CthCD-CT) and 5I6I (CthΔBCCP). How to cite this article: Hunkeler, M. et al. The dynamic organization of fungal acetyl-CoA carboxylase. Nat. Commun. 7:11196 doi: 10.1038/ncomms11196 (2016). Supplementary Material Evidence for the participation of biotin in the enzymic synthesis of fatty acids Fatty acid synthesis and its regulation Continuous fatty acid oxidation and reduced fat storage in mice lacking acetyl-CoA carboxylase 2 Acetyl-CoA carboxylase 2 mutant mice are protected against obesity and diabetes induced by high-fat/high-carbohydrate diets Acetyl-CoA carboxylase inhibition for the treatment of metabolic syndrome Enzymes of the fatty acid synthesis pathway are highly expressed in in situ breast carcinoma Increased lipogenesis in cancer cells: new players, novel targets Structural basis of phosphopeptide recognition by the BRCT domain of BRCA1 Structural evidence for direct interactions between the BRCT domains of human BRCA1 and a phospho-peptide from human ACC1 Acetyl-coenzyme A carboxylase: crucial metabolic enzyme and attractive target for drug discovery Fatty acid metabolism: target for metabolic syndrome A mechanism for the potent inhibition of eukaryotic acetyl-coenzyme A carboxylase by soraphen A, a macrocyclic polyketide natural product Bacterial fatty acid biosynthesis: targets for antibacterial drug discovery Expression and characterization of recombinant fungal acetyl-CoA carboxylase and isolation of a soraphen-binding domain Acetyl CoA carboxylase. I. Requirement for two protein fractions Acetyl CoA carboxylase, II. Deomonstration of biotin-protein and biotin carboxylase subunits Multi-subunit acetyl-CoA carboxylases Identification of an isozymic form of acetyl-CoA carboxylase The subcellular localization of acetyl-CoA carboxylase 2 Regulation of acetyl-CoA carboxylase Regulation of mammalian acetyl-CoA carboxylase Improving production of malonyl coenzyme A-derived metabolites by abolishing Snf1-dependent regulation of Acc1 Crystal structure of the alpha(6)beta(6) holoenzyme of propionyl-coenzyme A carboxylase An unanticipated architecture of the 750-kDa alpha6beta6 holoenzyme of 3-methylcrotonyl-CoA carboxylase Three-dimensional structure of the biotin carboxylase subunit of acetyl-CoA carboxylase Structure of the biotinyl domain of acetyl-coenzyme A carboxylase determined by MAD phasing Crystal structure of the carboxyltransferase domain of acetyl-coenzyme A carboxylase The structure of the carboxyltransferase component of acetyl-coA carboxylase reveals a zinc-binding motif unique to the bacterial enzyme Structure and function of biotin-dependent carboxylases Domain architecture of pyruvate carboxylase, a biotin-dependent multifunctional enzyme Structure and function of a single-chain, multi-domain long-chain acyl-CoA carboxylase Induced polymerization of mammalian acetyl-CoA carboxylase by MIG12 provides a tertiary level of regulation of fatty acid synthesis ACCA phosphopeptide recognition by the BRCT repeats of BRCA1 Location and function of three sites phosphorylated on rat acetyl-CoA carboxylase by the AMP-activated protein kinase Critical phosphorylation sites for acetyl-CoA carboxylase activity Molecular mechanism for the regulation of human ACC2 through phosphorylation by AMPK Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae Yeast SNF1 is functionally related to mammalian AMP-activated protein kinase and regulates acetyl-CoA carboxylase in vivo Global analysis of Cdk1 substrate phosphorylation sites provides insights into evolution Kinetic and structural analysis of a new group of Acyl-CoA carboxylases found in Streptomyces coelicolor A3(2) Crystal structure of the 500-kDa yeast acetyl-CoA carboxylase holoenzyme dimer Effect of interdomain dynamics on the structure determination of modular proteins by small-angle scattering Architecture of the polyketide synthase module: surprises from electron cryo-microscopy The crystal structure of a mammalian fatty acid synthase Asturias FJ. Conformational flexibility of metazoan fatty acid synthase enables catalysis Nonribosomal peptide synthetases: structures and dynamics The pyruvate dehydrogenase complexes: structure-based function and regulation Protein complex expression by using multigene baculoviral vectors XDS Combining constraints for electron-density modification Overview of the CCP4 suite and current developments PHENIX: a comprehensive Python-based system for macromolecular structure solution Features and development of Coot A short history of SHELX Secondary-structure matching (SSM), a new tool for fast protein structure alignment in three dimensions HKL2MAP: a graphical user interface for macromolecular phasing with SHELX programs Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during mitosis An enzyme assisted RP-RPLC approach for in-depth analysis of human liver phosphoproteome Combining protein-based IMAC, peptide-based IMAC, and MudPIT for efficient phosphoproteomic analysis Phaser crystallographic software SWISS-MODEL: modelling protein tertiary and quaternary structure using evolutionary information AL2CO: calculation of positional conservation in a protein sequence alignment Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega New developments in the program package for small-angle scattering data analysis Synchrotron-based small-angle X-ray scattering of proteins in solution Small-angle scattering for structural biology-Expanding the frontier while avoiding the pitfalls PRIMUS: a Windows PC-based system for small-angle scattering data analysis Determination of the molecular weight of proteins in solution from a single small-angle X-ray scattering measurement on a relative scale A Bayesian view on cryo-EM structure determination Xmipp: an image processing package for electron microscopy Author contributions M.H. cloned, expressed, purified and crystallized fungal ACC constructs, determined their structure and carried out SAXS analysis. E.S. cloned, expressed and crystallized human ACC CD and determined its structure. EM analysis was carried out by E.S., M.H. and A.H. S.I. contributed to structural analysis and figure preparation. T.M. designed and supervised work and analysed crystallographic data; all authors contributed to manuscript preparation. The phosphorylated central domain of yeast ACC. (a) Schematic overview of the domain organization of eukaryotic ACCs. Crystallized constructs are indicated. (b) Cartoon representation of the SceCD crystal structure. CDN is linked by a four-helix bundle (CDL) to two α–β-fold domains (CDC1 and CDC2). The regulatory loop is shown as bold cartoon, and the phosphorylated Ser1157 is marked by a red triangle. The N- and C termini are indicated by spheres. (c) Superposition of CDC1 and CDC2 reveals highly conserved folds. (d) The regulatory loop with the phosphorylated Ser1157 is bound into a crevice between CDC1 and CDC2, the conserved residues Arg1173 and Arg1260 coordinate the phosphoryl-group. (e) Structural overview of HsaBT-CD. The attachment points to the N-terminal BCCP domain and the C-terminal CT domain are indicated with spheres. All colourings are according to scheme a. Architecture of the CD–CT core of fungal ACC. Cartoon representation of crystal structures of multidomain constructs of CthACC. One protomer is shown in colour and one in grey. Individual domains are labelled; the active site of CT and the position of the conserved regulatory phosphoserine site based on SceCD are indicated by an asterisk and a triangle, respectively. Variability of the connections of CDC2 to CT and CDC1 in fungal ACC. (a) Hinge properties of the CDC2–CT connection analysed by a CT-based superposition of eight instances of the CDC2-CT segment. For clarity, only one protomer of CthCD-CTCter1 is shown in full colour as reference. For other instances, CDC2 domains are shown in transparent tube representation with only one helix each highlighted. The range of hinge bending is indicated and the connection points between CDC2 and CT (blue) as well as between CDC1 and CDC2 (green and grey) are marked as spheres. (b) The interdomain interface of CDC1 and CDC2 exhibits only limited plasticity. Representation as in a, but the CDC1 and CDC2 are superposed based on CDC2. One protomer of CthΔBCCP is shown in colour, the CDL domains are omitted for clarity and the position of the phosphorylated serine based on SceCD is indicated with a red triangle. The connection points from CDC1 to CDC2 and to CDL are represented by green spheres. The conformational dynamics of fungal ACC. (a–c) Large-scale conformational variability of the CDN domain relative to the CDL/CDC1 domain. CthCD-CT1 (in colour) serves as reference, the compared structures (as indicated, numbers after construct name differentiate between individual protomers) are shown in grey. Domains other than CDN and CDL/CDC1 are omitted for clarity. The domains are labelled and the distances between the N termini of CDN (spheres) in the compared structures are indicated. (d) Schematic model of fungal ACC showing the intrinsic, regulated flexibility of CD in the phosphorylated inhibited or the non-phosphorylated activated state. Flexibility of the CDC2/CT and CDN/CDL hinges is illustrated by arrows. The Ser1157 phosphorylation site and the regulatory loop are schematically indicated in magenta. Crystallographic data collection and refinement statistics.  \tSceCD\tSceCD Thimerosal\tSceCD Eu\tHsaBT-CD\tHsaBT-CD Cd2+\tCthCD-CTCter1\tCthCD-CTCter2\tCthCD-CT\tCthΔBCCP\t \tData collection\t \t Space group\tP43212\tP43212\tP43212\tI4122\tI4122\tP212121\tP212121\tP31212\tP6422\t \t Cell dimensions\t \t \t \t \t \t \t \t \t \t \t  a, b, c (Å)\t110.86, 110.86, 131.12\t111.22, 111.22, 131.49\t108.65, 108.65, 127.36\t267.27, 267.27, 210.61\t267.67, 267.67, 210.46\t97.66, 165.34, 219.23\t100.17, 153.45, 249,24\t295.02, 295.02, 189.52\t462.20, 462.20, 204.64\t \t  α, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 120\t90, 90, 120\t \t Resolution* (Å)\t3.0\t3.4\t4.0\t3.7\t4.1\t3.6\t4.5\t7.2\t8.4\t \t RMerge†\t18.2 (389.6)\t20.5 (306.1)\t40.6 (327.0)\t7.5 (400.9)\t15 (730.5)\t14.5 (384.5)\t27.4 (225.6)\t5.6 (302.6)\t29.4 (381.7)\t \t CC ½*†\t100 (58.3)\t99.9 (42.6)\t99.9 (48.5)\t100 (59.4)\t99.8 (73.2)\t99.9 (50.9)\t99.5 (46.7)\t100 (33.3)\t99.7 (35)\t \t I/σI†\t24.68 (1.46)\t7.99 (0.89)\t17.92 (1.85)\t21.24 (1.07)\t16.53 (1.41)\t10.61 (0.97)\t6.35 (1.00)\t18.95 (0.92)\t9.05 (0.9)\t \t Completeness†\t99.9 (99.9)\t99.6 (100)\t99.7 (96.8)\t99.8 (99.1)\t99.8 (99.7)\t99.7 (99.9)\t99.4 (98.6)\t99.6 (100)\t99.1 (99.9)\t \t Redundancy†\t39.1 (39.8)\t12.1 (14.3)\t81.6 (65.2)\t13.7 (13.7)\t20.9 (19.1)\t12.7 (13.5)\t6.1 (6.5)\t9.9 (10.4)\t18.5 (18.2)\t \t \t \t \t \t \t \t \t \t \t \t \tRefinement\t \t Resolution* (Å)\t46.4–3.0\t \t \t84.5–3.7\t \t49.2–3.6\t49.1–4.5\t49.9–7.2\t50.0–8.4\t \t Reflections\t16,928\t—\t—\t40,647\t—\t41,799\t23,340\t14,046\t12,111\t \t Rwork/Rfree\t0.20/0.24\t—\t—\t0.35/0.38‡\t—\t0.20/0.24\t0.24/0.24\t0.23/0.25\t0.30/0.32\t \t Number of atoms\t \t \t \t \t \t \t \t \t \t \t  Protein\t5,465\t \t \t6,925\t \t16,592\t16,405\t22,543\t22,445\t \t  Waters\t43\t—\t—\t—\t—\t—\t—\t—\t—\t \t  Ligand/ion\t7\t—\t—\t5\t—\t—\t—\t—\t—\t \t B-factors\t \t \t \t \t \t \t \t \t \t \t  Protein\t130\t \t \t158\t \t226\t275\t272\t250\t \t  Waters\t84\t—\t—\t—\t—\t—\t—\t—\t—\t \t  Ligand/ion\t90\t—\t—\t189\t—\t—\t—\t—\t—\t \t R.m.s.d.\t \t \t \t \t \t \t \t \t \t \t  RMS (angles, °)\t0.97\t—\t—\t0.83\t—\t1.07\t1.11\t1.15\t1.01\t \t  RMS (bonds, Å)\t0.01\t—\t—\t0.01\t—\t0.01\t0.01\t0.01\t0.01\t \t *Resolution cutoffs determined based on internal correlation significant at the 0.1% level as calculated by XDS. †Highest-resolution shell is shown in parentheses. ‡Modelled only as 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diff --git a/annotated_BioC_JSON/PMC4841544_ann.json b/annotated_BioC_JSON/PMC4841544_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..5bb21064e5f0e13e4b7f7a95a71f94fa088a0169
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4841544_ann.json
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+[{"sourceid":"4841544","sourcedb":"","project":"","target":"","text":"Molecular Basis of Ligand-Dependent Regulation of NadR, the Transcriptional Repressor of Meningococcal Virulence Factor NadA  Neisseria adhesin A (NadA) is present on the meningococcal surface and contributes to adhesion to and invasion of human cells. NadA is also one of three recombinant antigens in the recently-approved Bexsero vaccine, which protects against serogroup B meningococcus. The amount of NadA on the bacterial surface is of direct relevance in the constant battle of host-pathogen interactions: it influences the ability of the pathogen to engage human cell surface-exposed receptors and, conversely, the bacterial susceptibility to the antibody-mediated immune response. It is therefore important to understand the mechanisms which regulate nadA expression levels, which are predominantly controlled by the transcriptional regulator NadR (Neisseria adhesin A Regulator) both in vitro and in vivo. NadR binds the nadA promoter and represses gene transcription. In the presence of 4-hydroxyphenylacetate (4-HPA), a catabolite present in human saliva both under physiological conditions and during bacterial infection, the binding of NadR to the nadA promoter is attenuated and nadA expression is induced. NadR also mediates ligand-dependent regulation of many other meningococcal genes, for example the highly-conserved multiple adhesin family (maf) genes, which encode proteins emerging with important roles in host-pathogen interactions, immune evasion and niche adaptation. To gain insights into the regulation of NadR mediated by 4-HPA, we combined structural, biochemical, and mutagenesis studies. In particular, two new crystal structures of ligand-free and ligand-bound NadR revealed (i) the molecular basis of ‘conformational selection’ by which a single molecule of 4-HPA binds and stabilizes dimeric NadR in a conformation unsuitable for DNA-binding, (ii) molecular explanations for the binding specificities of different hydroxyphenylacetate ligands, including 3Cl,4-HPA which is produced during inflammation, (iii) the presence of a leucine residue essential for dimerization and conserved in many MarR family proteins, and (iv) four residues (His7, Ser9, Asn11 and Phe25), which are involved in binding 4-HPA, and were confirmed in vitro to have key roles in the regulatory mechanism in bacteria. Overall, this study deepens our molecular understanding of the sophisticated regulatory mechanisms of the expression of nadA and other genes governed by NadR, dependent on interactions with niche-specific signal molecules that may play important roles during meningococcal pathogenesis. Author Summary Serogroup B meningococcus (MenB) causes fatal sepsis and invasive meningococcal disease, particularly in young children and adolescents, as highlighted by recent MenB outbreaks in universities of the United States and Canada. The Bexsero vaccine protects against MenB and has recently been approved in \u003e 35 countries worldwide. Neisseria adhesin A (NadA) present on the meningococcal surface can mediate binding to human cells and is one of the three MenB vaccine protein antigens. The amount of NadA exposed on the meningococcal surface also influences the antibody-mediated serum bactericidal response measured in vitro. A deep understanding of nadA expression is therefore important, otherwise the contribution of NadA to vaccine-induced protection against meningococcal meningitis may be underestimated. The abundance of surface-exposed NadA is regulated by the ligand-responsive transcriptional repressor NadR. Here, we present functional, biochemical and high-resolution structural data on NadR. Our studies provide detailed insights into how small molecule ligands, such as hydroxyphenylacetate derivatives, found in relevant host niches, modulate the structure and activity of NadR, by ‘conformational selection’ of inactive forms. These findings shed light on the regulation of NadR, a key MarR-family virulence factor of this important human pathogen. Introduction The ‘Reverse Vaccinology’ approach was pioneered to identify antigens for a protein-based vaccine against serogroup B Neisseria meningitidis (MenB), a human pathogen causing potentially-fatal sepsis and invasive meningococcal disease. Indeed, Reverse Vaccinology identified Neisseria adhesin A (NadA), a surface-exposed protein involved in epithelial cell invasion and found in ~30% of clinical isolates. Recently, we reported the crystal structure of NadA, providing insights into its biological and immunological functions. Recombinant NadA elicits a strong bactericidal immune response and is therefore included in the Bexsero vaccine that protects against MenB and which was recently approved in over 35 countries worldwide. Previous studies revealed that nadA expression levels are mainly regulated by the Neisseria adhesin A Regulator (NadR). Although additional factors influence nadA expression, we focused on its regulation by NadR, the major mediator of NadA phase variable expression. Studies of NadR also have broader implications, since a genome-wide analysis of MenB wild-type and nadR knock-out strains revealed that NadR influences the regulation of \u003e 30 genes, including maf genes, from the multiple adhesin family. These genes encode a wide variety of proteins connected to many biological processes contributing to bacterial survival, adaptation in the host niche, colonization and invasion. NadR belongs to the MarR (Multiple Antibiotic Resistance Regulator) family, a group of ligand-responsive transcriptional regulators ubiquitous in bacteria and archaea. MarR family proteins can promote bacterial survival in the presence of antibiotics, toxic chemicals, organic solvents or reactive oxygen species and can regulate virulence factor expression. MarR homologues can act either as transcriptional repressors or as activators. Although \u003e 50 MarR family structures are known, a molecular understanding of their ligand-dependent regulatory mechanisms is still limited, often hampered by lack of identification of their ligands and/or DNA targets. A potentially interesting exception comes from the ligand-free and salicylate-bound forms of the Methanobacterium thermoautotrophicum protein MTH313 which revealed that two salicylate molecules bind to one MTH313 dimer and induce large conformational changes, apparently sufficient to prevent DNA binding. However, the homologous archeal Sulfolobus tokodaii protein ST1710 presented essentially the same structure in ligand-free and salicylate-bound forms, apparently contrasting the mechanism proposed for MTH313. Despite these apparent differences, MTH313 and ST1710 bind salicylate in approximately the same site, between their dimerization and DNA-binding domains. However, it is unknown whether salicylate is a relevant in vivo ligand of either of these two proteins, which share ~20% sequence identity with NadR, rendering unclear the interpretation of these findings in relation to the regulatory mechanisms of NadR or other MarR family proteins. NadR binds the nadA promoter and represses gene transcription. NadR binds nadA on three different operators (OpI, OpII and OpIII). The DNA-binding activity of NadR is attenuated in vitro upon addition of various hydroxyphenylacetate (HPA) derivatives, including 4-HPA. 4-HPA is a small molecule derived from mammalian aromatic amino acid catabolism and is released in human saliva, where it has been detected at micromolar concentration. In the presence of 4-HPA, NadR is unable to bind the nadA promoter and nadA gene expression is induced. In vivo, the presence of 4-HPA in the host niche of N. meningitidis serves as an inducer of NadA production, thereby promoting bacterial adhesion to host cells. Further, we recently reported that 3Cl,4-HPA, produced during inflammation, is another inducer of nadA expression. Extending our previous studies based on hydrogen-deuterium exchange mass spectrometry (HDX-MS), here we sought to reveal the molecular mechanisms and effects of NadR/HPA interactions via X-ray crystallography, NMR spectroscopy and complementary biochemical and in vivo mutagenesis studies. We obtained detailed new insights into ligand specificity, how the ligand allosterically influences the DNA-binding ability of NadR, and the regulation of nadA expression, thus also providing a deeper structural understanding of the ligand-responsive MarR super-family. Moreover, these findings are important because the activity of NadR impacts the potential coverage provided by anti-NadA antibodies elicited by the Bexsero vaccine and influences host-bacteria interactions that contribute to meningococcal pathogenesis. Results NadR is dimeric and is stabilized by specific hydroxyphenylacetate ligands Recombinant NadR was produced in E. coli using an expression construct prepared from N. meningitidis serogroup B strain MC58. Standard chromatographic techniques were used to obtain a highly purified sample of NadR (see Materials and Methods). In analytical size-exclusion high-performance liquid chromatography (SE-HPLC) experiments coupled with multi-angle laser light scattering (MALLS), NadR presented a single species with an absolute molecular mass of 35 kDa (S1 Fig). These data showed that NadR was dimeric in solution, since the theoretical molecular mass of the NadR dimer is 33.73 kDa; and, there was no change in oligomeric state on addition of 4-HPA. The thermal stability of NadR was examined using differential scanning calorimetry (DSC). Since ligand-binding often increases protein stability, we also investigated the effect of various HPAs (Fig 1A) on the melting temperature (Tm) of NadR. As a control of specificity, we also tested salicylate, a known ligand of some MarR proteins previously reported to increase the Tm of ST1710 and MTH313. The Tm of NadR was 67.4 ± 0.1°C in the absence of ligand, and was unaffected by salicylate. However, an increased thermal stability was induced by 4-HPA and, to a lesser extent, by 3-HPA. Interestingly, NadR displayed the greatest Tm increase upon addition of 3Cl,4-HPA (Table 1 and Fig 1B). Stability of NadR is increased by small molecule ligands. \n(A) Molecular structures of 3-HPA (MW 152.2), 4-HPA (MW 152.2), 3Cl,4-HPA (MW 186.6) and salicylic acid (MW 160.1). (B) DSC profiles, colored as follows: apo-NadR (violet), NadR+salicylate (red), NadR+3-HPA (green), NadR+4-HPA (blue), NadR+3Cl,4-HPA (pink). All DSC profiles are representative of triplicate experiments. Melting-point (Tm) and its ligand-induced increase (ΔTm) derived from DSC thermostability experiments. Dissociation constants (KD) of the NadR/ligand interactions from SPR steady-state binding experiments. Ligand\tTm (°C)\tΔTm (°C)\tKD (mM)\t \tNo ligand\t67.4 ± 0.1\tn.a.\tn.a.\t \tSalicylate\t67.5 ± 0.1\t0\tn.d.\t \t3-HPA\t70.0 ± 0.1\t2.7\t2.7 ± 0.1\t \t4-HPA\t70.7 ± 0.1\t3.3\t1.5 ± 0.1\t \t3Cl,4-HPA\t71.3 ± 0.2\t3.9\t1.1 ± 0.1\t \t n.a.: not applicable; n.d.: not determinable NadR displays distinct binding affinities for hydroxyphenylacetate ligands To further investigate the binding of HPAs to NadR, we used surface plasmon resonance (SPR). The SPR sensorgrams revealed very fast association and dissociation events, typical of small molecule ligands, thus prohibiting a detailed study of binding kinetics. However, steady-state SPR analyses of the NadR-HPA interactions allowed determination of the equilibrium dissociation constants (KD) (Table 1 and S2 Fig). The interactions of 4-HPA and 3Cl,4-HPA with NadR exhibited KD values of 1.5 mM and 1.1 mM, respectively. 3-HPA showed a weaker interaction, with a KD of 2.7 mM, while salicylate showed only a very weak response that did not reach saturation, indicating a non-specific interaction with NadR. A ranking of these KD values showed that 3Cl,4-HPA was the tightest binder, and thus matched the ranking of ligand-induced Tm increases observed in the DSC experiments. Although these KD values indicate rather weak interactions, they are similar to the values reported previously for the MarR/salicylate interaction (KD ~1 mM) and the MTH313/salicylate interaction (KD 2–3 mM), and approximately 20-fold tighter than the ST1710/salicylate interaction (KD ~20 mM). Crystal structures of holo-NadR and apo-NadR To fully characterize the NadR/HPA interactions, we sought to determine crystal structures of NadR in ligand-bound (holo) and ligand-free (apo) forms. First, we crystallized NadR (a selenomethionine-labelled derivative) in the presence of a 200-fold molar excess of 4-HPA. The structure of the NadR/4-HPA complex was determined at 2.3 Å resolution using a combination of the single-wavelength anomalous dispersion (SAD) and molecular replacement (MR) methods, and was refined to R work/R free values of 20.9/26.0% (Table 2). Despite numerous attempts, we were unable to obtain high-quality crystals of NadR complexed with 3Cl,4-HPA, 3,4-HPA, 3-HPA or DNA targets. However, it was eventually possible to crystallize apo-NadR, and the structure was determined at 2.7 Å resolution by MR methods using the NadR/4-HPA complex as the search model. The apo-NadR structure was refined to R work/R free values of 19.1/26.8% (Table 2). Data collection and refinement statistics for NadR structures. \tNadR SeMet + 4-HPA (SAD peak) (PDB code 5aip)\tNadR apo-form (PDB code 5aiq)\t \tData collection\t\t\t \tWavelength (Å)\t0.9792\t1.0\t \tBeamline\tSLS (PXII-X10SA)\tSLS (PXII-X10SA)\t \tResolution range (Å)\t39.2–2.3\t48.2–2.7\t \tSpace group\tP 43 21 2\tP 43 21 2\t \tUnit cell dimensions (Å)\t75.3, 75.3, 91.8\t69.4, 69.4, 253.8\t \tTotal reflections\t291132 (41090)\t225521 (35809)\t \tUnique reflections\t12320 (1773)\t17700 (2780)\t \tMultiplicity\t23.6 (23.2)\t12.7 (12.8)\t \tCompleteness (%)\t100.0 (100.00)\t99.9 (99.7)\t \tMean I/sigma(I)\t25.5 (9.0)\t22.6 (3.8)\t \tWilson B-factor\t23.9\t49.1\t \tRsym*\t10.9 (39.4)\t11.4 (77.6)\t \tRmeas**\t11.3\t11.8\t \tRefinement\t\t\t \tRwork♯\t20.9\t21.7\t \tRfree♯♯\t26.0\t27.2\t \tNumber of atoms\t\t\t \t  Non-hydrogen atoms\t2263\t4163\t \t  Macromolecules\t2207\t4144\t \t  Ligands\t11\t0\t \t  Water\t45\t19\t \tProtein residues\t275\t521\t \tRMS(bonds)\t0.008\t0.003\t \tRMS(angles)\t1.09\t0.823\t \tRamachandran (%)§\t\t\t \t  Favored\t100\t98.4\t \t  Outliers\t0\t0\t \tClash score\t5.0\t3.9\t \tAverage B-factor\t\t\t \t  Macromolecules\t34.8\t53.3\t \t  Ligands\t32.9\t-\t \t  Solvent\t37.3 (H2O)\t29.0 (H2O)\t \t Statistics for the highest-resolution shell are shown in parentheses. *R\nsym = Σhkl Σi |Ii(hkl)—\u003cI(hkl)\u003e| / Σhkl Σi Ii(hkl) ** R\nmeas = redundancy-independent (multiplicity-weighted) R\nmerge as reported from AIMLESS. \n♯\nR\nwork = Σ||F(obs)|- |F(calc)||/Σ|F(obs)| \n♯♯\nR\nfree = as for R\nwork, calculated for 5.0% of the total reflections, chosen at random, and omitted from refinement. \n§ Values obtained using Molprobity. The asymmetric unit of the NadR/4-HPA crystals (holo-NadR) contained one NadR homodimer, while the apo-NadR crystals contained two homodimers. In the apo-NadR crystals, the two homodimers were related by a rotation of ~90°; the observed association of the two dimers was presumably merely an effect of crystal packing, since the interface between the two homodimers is small (\u003c 550 Å2 of buried surface area), and is not predicted to be physiologically relevant by the PISA software. Moreover, our SE-HPLC/MALLS analyses (see above) revealed that in solution NadR is dimeric, and previous studies using native mass spectrometry (MS) revealed dimers, not tetramers. The NadR homodimer bound to 4-HPA has a dimerization interface mostly involving the top of its ‘triangular’ form, while the two DNA-binding domains are located at the base (Fig 2A). High-quality electron density maps allowed clear identification of the bound ligand, 4-HPA (Fig 2B). The overall structure of NadR shows dimensions of ~50 × 65 × 50 Å and a large homodimer interface that buries a total surface area of ~ 4800 Å2. Each NadR monomer consists of six α-helices and two short β-strands, with helices α1, α5, and α6 forming the dimer interface. Helices α3 and α4 form a helix-turn-helix motif, followed by the “wing motif” comprised of two short antiparallel β-strands (β1-β2) linked by a relatively long and flexible loop. Interestingly, in the α4-β2 region, the stretch of residues from R64-R91 presents seven positively-charged side chains, all available for potential interactions with DNA. Together, these structural elements constitute the winged helix-turn-helix (wHTH) DNA-binding domain and, together with the dimeric organization, are the hallmarks of MarR family structures. The crystal structure of NadR in complex with 4-HPA. \n(A) The holo-NadR homodimer is depicted in green and blue for chains A and B respectively, while yellow sticks depict the 4-HPA ligand (labelled). For simplicity, secondary structure elements are labelled for chain B only. Red dashes show hypothetical positions of chain B residues 88–90 that were not modeled due to lack of electron density. (B) A zoom into the pocket occupied by 4-HPA shows that the ligand contacts both chains A and B; blue mesh shows electron density around 4-HPA calculated from a composite omit map (omitting 4-HPA), using phenix. The map is contoured at 1σ and the figure was prepared with a density mesh carve factor of 1.7, using Pymol (www.pymol.org). A single conserved leucine residue (L130) is crucial for dimerization The NadR dimer interface is formed by at least 32 residues, which establish numerous inter-chain salt bridges or hydrogen bonds, and many hydrophobic packing interactions (Fig 3A and 3B). To determine which residues were most important for dimerization, we studied the interface in silico and identified several residues as potential mediators of key stabilizing interactions. Using site-directed mutagenesis, a panel of eight mutant NadR proteins was prepared (including mutations H7A, S9A, N11A, D112A, R114A, Y115A, K126A, L130K and L133K), sufficient to explore the entire dimer interface. Each mutant NadR protein was purified, and then its oligomeric state was examined by analytical SE-HPLC. Almost all the mutants showed the same elution profile as the wild-type (WT) NadR protein. Only the L130K mutation induced a notable change in the oligomeric state of NadR (Fig 3C). Further, in SE-MALLS analyses, the L130K mutant displayed two distinct species in solution, approximately 80% being monomeric (a 19 kDa species), and only 20% retaining the typical native dimeric state (a 35 kDa species) (Fig 3D), demonstrating that Leu130 is crucial for stable dimerization. It is notable that L130 is usually present as Leu, or an alternative bulky hydrophobic amino acid (e.g. Phe, Val), in many MarR family proteins, suggesting a conserved role in stabilizing the dimer interface. In contrast, most of the other residues identified in the NadR dimer interface were poorly conserved in the MarR family. Analysis of the NadR dimer interface. \n(A) Both orientations show chain A, green backbone ribbon, colored red to highlight all locations involved in dimerization; namely, inter-chain salt bridges or hydrogen bonds involving Q4, S5, K6, H7, S9, I10, N11, I15, Q16, R18, D36, R43, A46, Q59, C61, Y104, D112, R114, Y115, D116, E119, K126, E136, E141, N145, and the hydrophobic packing interactions involving I10, I12, L14, I15, R18, Y115, I118, L130, L133, L134 and L137. Chain B, grey surface, is marked blue to highlight residues probed by site-directed mutagenesis (E136 only makes a salt bridge with K126, therefore it was sufficient to make the K126A mutation to assess the importance of this ionic interaction; the H7 position is labelled for monomer A, since electron density was lacking for monomer B). (B) A zoom into the environment of helix α6 to show how residue L130 chain B (blue side chain) is a focus of hydrophobic packing interactions with L130, L133, L134 and L137 of chain A (red side chains). (C) SE-HPLC analyses of all mutant forms of NadR are compared with the wild-type (WT) protein. The WT and most of the mutants show a single elution peak with an absorbance maximum at 17.5 min. Only the mutation L130K has a noteworthy effect on the oligomeric state, inducing a second peak with a longer retention time and a second peak maximum at 18.6 min. To a much lesser extent, the L133K mutation also appears to induce a ‘shoulder’ to the main peak, suggesting very weak ability to disrupt the dimer. (D) SE-HPLC/MALLS analyses of the L130K mutant, shows 20% dimer and 80% monomer. The curves plotted correspond to Absorbance Units (mAU) at 280nm wavelength (green), light scattering (red), and refractive index (blue). The holo-NadR structure presents only one occupied ligand-binding pocket The NadR/4-HPA structure revealed the ligand-binding site nestled between the dimerization and DNA-binding domains (Fig 2). The ligand showed a different position and orientation compared to salicylate complexed with MTH313 and ST1710 (see Discussion). The binding pocket was almost entirely filled by 4-HPA and one water molecule, although there also remained a small tunnel 2-4Å in diameter and 5-6Å long leading from the pocket (proximal to the 4-hydroxyl position) to the protein surface. The tunnel was lined with rather hydrophobic amino acids, and did not contain water molecules. Unexpectedly, only one monomer of the holo-NadR homodimer contained 4-HPA in the binding pocket, whereas the corresponding pocket of the other monomer was unoccupied by ligand, despite the large excess of 4-HPA used in the crystallization conditions. Inspection of the protein-ligand interaction network revealed no bonds from NadR backbone groups to the ligand, but several key side chain mediated hydrogen (H)-bonds and ionic interactions, most notably between the carboxylate group of 4-HPA and Ser9 of chain A (SerA9), and chain B residues TrpB39, ArgB43 and TyrB115 (Fig 4A). At the other ‘end’ of the ligand, the 4-hydroxyl group was proximal to AspB36, with which it may establish an H-bond (see bond distances in Table 3). The water molecule observed in the pocket was bound by the carboxylate group and the side chains of SerA9 and AsnA11. Atomic details of NadR/HPA interactions. \nA) A stereo-view zoom into the binding pocket showing side chain sticks for all interactions between NadR and 4-HPA. Green and blue ribbons depict NadR chains A and B, respectively. 4-HPA is shown in yellow sticks, with oxygen atoms in red. A water molecule is shown by the red sphere. H-bonds up to 3.6Å are shown as dashed lines. The entire set of residues making H-bonds or non-bonded contacts with 4-HPA is as follows: SerA9, AsnA11, LeuB21, MetB22, PheB25, LeuB29, AspB36, TrpB39, ArgB43, ValB111 and TyrB115 (automated analysis performed using PDBsum and verified manually). Residues AsnA11 and ArgB18 likely make indirect yet local contributions to ligand binding, mainly by stabilizing the position of AspB36. Bond distances for interacting polar atoms are provided in Table 3. Side chains mediating hydrophobic interactions are shown in orange. (B) A model was prepared to visualize putative interactions of 3Cl,4-HPA (pink) with NadR, revealing the potential for additional contacts (dashed lines) of the chloro moiety (green stick) with LeuB29 and AspB36. List of 4-HPA atoms bound to NadR via ionic interactions and/or H-bonds. 4-HPA atom\tNadR residue/atom\tDistance (Å)\t \tO2\tTrpB39/NE1\t2.83\t \tO2\tArgB43/NH1\t2.76\t \tO1\tArgB43/NH1\t3.84\t \tO1\tSerA9/OG\t2.75\t \tO1\tTyrB115/OH\t2.50\t \tO2\tWater (*Ser9/Asn11)\t2.88\t \tOH\tAspB36/OD1/OD2\t3.6/3.7\t \t * Bond distance between the ligand carboxylate group and the water molecule, which in turn makes H-bond to the SerA9 and AsnA11 side chains. In addition to the H-bonds involving the carboxylate and hydroxyl groups of 4-HPA, binding of the phenyl moiety appeared to be stabilized by several van der Waals’ contacts, particularly those involving the hydrophobic side chain atoms of LeuB21, MetB22, PheB25, LeuB29 and ValB111 (Fig 4A). Notably, the phenyl ring of PheB25 was positioned parallel to the phenyl ring of 4-HPA, potentially forming π-π parallel-displaced stacking interactions. Consequently, residues in the 4-HPA binding pocket are mostly contributed by NadR chain B, and effectively created a polar ‘floor’ and a hydrophobic ‘ceiling’, which house the ligand. Collectively, this mixed network of polar and hydrophobic interactions endows NadR with a strong recognition pattern for HPAs, with additional medium-range interactions potentially established with the hydroxyl group at the 4-position. Structure-activity relationships: molecular basis of enhanced stabilization by 3Cl,4-HPA We modelled the binding of other HPAs by in silico superposition onto 4-HPA in the holo-NadR structure, and thereby obtained molecular explanations for the binding specificities of diverse ligands. For example, similar to 4-HPA, the binding of 3Cl,4-HPA could involve multiple bonds towards the carboxylate group of the ligand and some to the 4-hydroxyl group. Additionally, the side chains of LeuB29 and AspB36 would be only 2.6–3.5 Å from the chlorine atom, thus providing van der Waals’ interactions or H-bonds to generate the additional binding affinity observed for 3Cl,4-HPA (Fig 4B). The presence of a single hydroxyl group at position 2, as in 2-HPA, rather than at position 4, would eliminate the possibility of favorable interactions with AspB36, resulting in the lack of NadR regulation by 2-HPA described previously. Finally, salicylate is presumably unable to specifically bind NadR due to the 2-hydroxyl substitution and the shorter aliphatic chain connecting its carboxylate group (Fig 1A): the compound simply seems too small to simultaneously establish the network of beneficial bonds observed in the NadR/HPA interactions. Analysis of the pockets reveals the molecular basis for asymmetric binding and stoichiometry We attempted to investigate further the binding stoichiometry using solution-based techniques. However, studies based on tryptophan fluorescence were confounded by the fluorescence of the HPA ligands, and isothermal titration calorimetry (ITC) was unfeasible due to the need for very high concentrations of NadR in the ITC chamber (due to the relatively low affinity), which exceeded the solubility limits of the protein. However, it was possible to calculate the binding stoichiometry of the NadR-HPA interactions using an SPR-based approach. In SPR, the signal measured is proportional to the total molecular mass proximal to the sensor surface; consequently, if the molecular weights of the interactors are known, then the stoichiometry of the resulting complex can be determined. This approach relies on the assumption that the captured protein (‘the ligand’, according to SPR conventions) is 100% active and freely-accessible to potential interactors (‘the analytes’). This assumption is likely valid for this pair of interactors, for two main reasons. Firstly, NadR is expected to be covalently immobilized on the sensor chip as a dimer in random orientations, since it is a stable dimer in solution and has sixteen lysines well-distributed around its surface, all able to act as potential sites for amine coupling to the chip, and none of which are close to the ligand-binding pocket. Secondly, the HPA analytes are all very small (MW 150–170, Fig 1A) and therefore are expected to be able to diffuse readily into all potential binding sites, irrespective of the random orientations of the immobilized NadR dimers on the chip.The stoichiometry of the NadR-HPA interactions was determined using Eq 1 (see Materials and Methods), and revealed stoichiometries of 1.13 for 4-HPA, 1.02 for 3-HPA, and 1.21 for 3Cl,4-HPA, strongly suggesting that one NadR dimer bound to 1 HPA analyte molecule. The crystallographic data, supported by the SPR studies of binding stoichiometry, revealed the lack of a second 4-HPA molecule in the homodimer, suggesting negative co-operativity, a phenomenon previously described for the MTH313/salicylate interaction and for other MarR family proteins. To explore the molecular basis of asymmetry in holo-NadR, we superposed its ligand-free monomer (chain A) onto the ligand-occupied monomer (chain B). Overall, the superposition revealed a high degree of structural similarity (Cα root mean square deviation (rmsd) of 1.5Å), though on closer inspection a rotational difference of ~9 degrees along the long axis of helix α6 was observed, suggesting that 4-HPA induced a slight conformational change (Fig 5A). However, since residues of helix α6 were not directly involved in ligand binding, an explanation for the lack of 4-HPA in monomer A did not emerge by analyzing only these backbone atom positions, suggesting that a more complex series of allosteric events may occur. Indeed, we noted interesting differences in the side chains of Met22, Phe25 and Arg43, which in monomer B are used to contact the ligand while in monomer A they partially occupied the pocket and collectively reduced its volume significantly. Specifically, upon analysis with the CASTp software, the pocket in chain B containing the 4-HPA exhibited a total volume of approximately 370 Å3, while the pocket in chain A was occupied by these three side chains that adopted ‘inward’ positions and thereby divided the space into a few much smaller pockets, each with volume \u003c 50 Å3, evidently rendering chain A unfavorable for ligand binding. Most notably, atomic clashes between the ligand and the side chains of MetA22, PheA25 and ArgA43 would occur if 4-HPA were present in the monomer A pocket (Fig 5B). Subsequently, analyses of the pockets in apo-NadR revealed that in the absence of ligand the long Arg43 side chain was always in the open ‘outward’ position compatible with binding to the 4-HPA carboxylate group. In contrast, the apo-form Met22 and Phe25 residues were still encroaching the spaces of the 4-hydroxyl group and the phenyl ring of the ligand, respectively (Fig 5C). The ‘outward’ position of Arg43 generated an open apo-form pocket with volume approximately 380Å3. Taken together, these observations suggest that Arg43 is a major determinant of ligand binding, and that its ‘inward’ position inhibits the binding of 4-HPA to the empty pocket of holo-NadR. Structural differences of NadR in ligand-bound or free forms. \n(A) Aligned monomers of holo-NadR (chain A: green; chain B: blue), reveal major overall differences by the shift of helix α6. (B) Comparison of the two binding pockets in holo-NadR shows that in the ligand-free monomer A (green) residues Met22, Phe25 and Arg43 adopt ‘inward’ positions (highlighted by arrows) compared to the ligand-occupied pocket (blue residues); these ‘inward’ conformations appear unfavorable for binding of 4-HPA due to clashes with the 4-hydroxyl group, the phenyl ring and the carboxylate group, respectively. In these crystals, the ArgA43 side chain showed two alternate conformations, modelled with 50% occupancy in each state, as indicated by the two ‘mirrored’ arrows. The inner conformer is the one that would display major clashes if 4-HPA were present. (C) Comparison of the empty pocket from holo-NadR (green residues) with the four empty pockets of apo-NadR (grey residues), shows that in the absence of 4-HPA the Arg43 side chain is always observed in the ‘outward’ conformation. Finally, we applied 15N heteronuclear solution NMR spectroscopy to examine the interaction of 4-HPA with apo NadR. We collected NMR spectra on NadR in the presence and absence of 4-HPA (see Materials and Methods). The 1H-15N TROSY-HSQC spectrum of apo-NadR, acquired at 25°C, displayed approximately 140 distinct peaks (Fig 6A), most of which correspond to backbone amide N-H groups. The broad spectral dispersion and the number of peaks observed, which is close to the number of expected backbone amide N-H groups for this polypeptide, confirmed that apo-NadR is well-folded under these conditions and exhibits one conformation appreciable on the NMR timescale, i.e. in the NMR experiments at 25°C, two or more distinct conformations of apo-NadR monomers were not readily apparent. Upon the addition of 4-HPA, over 45 peaks showed chemical shift perturbations, i.e. changed position in the spectrum or disappeared, while the remaining peaks remained unchanged. This observation showed that 4-HPA was able to bind NadR and induce notable changes in specific regions of the protein. NMR spectra of NadR in the presence and absence of 4-HPA. \n(A) Superposition of two 1H-15N TROSY-HSQC spectra recorded at 25°C on apo-NadR (cyan) and on NadR in the presence of 4-HPA (red). (B,C) Overlay of selected regions of the 1H-15N TROSY-HSQC spectra acquired at 25°C of apo-NadR (cyan) and NadR/4-HPA (red) superimposed with the spectra acquired at 10°C of apo-NadR (blue) and NadR/4-HPA (green). The spectra acquired at 10°C are excluded from panel A for simplicity. However, in the presence of 4-HPA, the 1H-15N TROSY-HSQC spectrum of NadR displayed approximately 140 peaks, as for apo-NadR, i.e. two distinct stable conformations (that might have potentially revealed the molecular asymmetry observed crystallographically) were not notable. Considering the small size, fast diffusion and relatively low binding affinity of 4-HPA, it would not be surprising if the ligand associates and dissociates rapidly on the NMR time scale, resulting in only one set of peaks whose chemical shifts represent the average environment of the bound and unbound states. Interestingly, by cooling the samples to 10°C, we observed that a number of those peaks strongly affected by 4-HPA (and therefore likely to be in the ligand-binding site) demonstrated evidence of peak splitting, i.e. a tendency to become two distinct peaks rather than one single peak (Fig 6B and 6C). These doubled peaks may therefore reveal that the cooler temperature partially trapped the existence in solution of two distinct states, in presence or absence of 4-HPA, with minor conformational differences occurring at least in proximity to the binding pocket. Although more comprehensive NMR experiments and full chemical shift assignment of the spectra would be required to precisely define this multi-state behavior, the NMR data clearly demonstrate that NadR exhibits conformational flexibility which is modulated by 4-HPA in solution. Apo-NadR structures reveal intrinsic conformational flexibility The apo-NadR crystal structure contained two homodimers in the asymmetric unit (chains A+B and chains C+D). Upon overall structural superposition, these dimers revealed a few minor differences in the α6 helix (a major component of the dimer interface) and the helices α4-α5 (the DNA binding region), and an rmsd of 1.55Å (Fig 7A). Similarly, the entire holo-homodimer could be closely superposed onto each of the apo-homodimers, showing rmsd values of 1.29Å and 1.31Å, and with more notable differences in the α6 helix positions (Fig 7B). The slightly larger rmsd between the two apo-homodimers, rather than between apo- and holo-homodimers, further indicate that apo-NadR possesses a notable degree of intrinsic conformational flexibility. Overall apo- and holo-NadR structures are similar. \n(A) Pairwise alignment of the two distinct apo-NadR homodimers (AB and CD) present in the apo-NadR crystals. (B) Alignment of the holo-NadR homodimer (green and blue chains) onto the apo-NadR homodimers. Here, larger differences are observed in the α6 helices (top). 4-HPA stabilizes concerted conformational changes in NadR that prevent DNA-binding To further investigate the conformational rearrangements of NadR, we performed local structural alignments using only a subset of residues in the DNA-binding helix (α4). By selecting and aligning residues Arg64-Ala77 of one α4 helix per dimer, superposition of the holo-homodimer onto the two apo-homodimers revealed differences in the monomer conformations of each structure. While one monomer from each structure was closely superimposable (Fig 8A, left side), the second monomer displayed quite large differences (Fig 8A, right side). Most notably, the position of the DNA-binding helix α4 shifted by as much as 6 Å (Fig 8B). Accordingly, helix α4 was also found to be one of the most dynamic regions in previous HDX-MS analyses of apo-NadR in solution. Structural comparisons of NadR and modelling of interactions with DNA. \n(A) The holo-homodimer structure is shown as green and blue cartoons, for chain A and B, respectively, while the two homodimers of apo-NadR are both cyan and pale blue for chains A/C and B/D, respectively. The three homodimers (chains AB holo, AB apo, and CD apo) were overlaid by structural alignment exclusively of all heavy atoms in residues R64-A77 (shown in red, with side chain sticks) of chains A holo, A apo, and C apo, belonging to helix α4 (left). The α4 helices aligned closely, Cα rmsd 0.2Å for 14 residues. (B) The relative positions of the α4 helices of the 4-HPA-bound holo homodimer chain B (blue), and of apo homodimers AB and CD (showing chains B and D) in pale blue. Dashes indicate the Ala77 Cα atoms, in the most highly shifted region of the ‘non-fixed’ α4 helix. (C) The double-stranded DNA molecule (grey cartoon) from the OhrR-ohrA complex is shown after superposition with NadR, to highlight the expected positions of the NadR α4 helices in the DNA major grooves. The proteins share ~30% amino acid sequence identity. For clarity, only the α4 helices are shown in panels (B) and (C). (D) Upon comparison with the experimentally-determined OhrR:ohrA structure (grey), the α4 helix of holo-NadR (blue) is shifted ~8Å out of the major groove. However, structural comparisons revealed that the shift of holo-NadR helix α4 induced by the presence of 4-HPA was also accompanied by several changes at the holo dimer interface, while such extensive structural differences were not observed in the apo dimer interfaces, particularly notable when comparing the α6 helices (S3 Fig). In summary, compared to ligand-stabilized holo-NadR, apo-NadR displayed an intrinsic flexibility focused in the DNA-binding region. This was also evident in the greater disorder (i.e. less well-defined electron density) in the β1-β2 loops of the apo dimers (density for 16 residues per dimer was missing) compared to the holo dimer (density for only 3 residues was missing). In holo-NadR, the distance separating the two DNA-binding α4 helices was 32 Å, while in apo-NadR it was 29 Å for homodimer AB, and 34 Å for homodimer CD (Fig 8C). Thus, the apo-homodimer AB presented the DNA-binding helices in a conformation similar to that observed in the protein:DNA complex of OhrR:ohrA from Bacillus subtilis (Fig 8C). Interestingly, OhrR contacts ohrA across 22 base pairs (bp), and similarly the main NadR target sites identified in the nadA promoter (the operators Op I and Op II) both span 22 bp. Pairwise superpositions showed that the NadR apo-homodimer AB was the most similar to OhrR (rmsd 2.6 Å), while the holo-homodimer was the most divergent (rmsd 3.3 Å) (Fig 8C). Assuming the same DNA-binding mechanism is used by OhrR and NadR, the apo-homodimer AB seems ideally pre-configured for DNA binding, while 4-HPA appeared to stabilize holo-NadR in a conformation poorly suited for DNA binding. Specifically, in addition to the different inter-helical translational distances, the α4 helices in the holo-NadR homodimer were also reoriented, resulting in movement of α4 out of the major groove, by up to 8Å, and presumably preventing efficient DNA binding in the presence of 4-HPA (Fig 8D). When aligned with OhrR, the apo-homodimer CD presented yet another different intermediate conformation (rmsd 2.9Å), apparently not ideally pre-configured for DNA binding, but which in solution can presumably readily adopt the AB conformation due to the intrinsic flexibility described above. NadR residues His7, Ser9, Asn11 and Phe25 are essential for regulation of NadA expression in vivo  While previous studies had correctly suggested the involvement of several NadR residues in ligand binding, the crystal structures presented here revealed additional residues with previously unknown roles in dimerization and/or binding to 4-HPA. To explore the functional involvement of these residues, we characterized the behavior of four new NadR mutants (H7A, S9A, N11A and F25A) in an in vivo assay using the previously described MC58-Δ1843 nadR-null mutant strain, which was complemented either by wild-type nadR or by the nadR mutants. NadA protein abundance levels were assessed by Western blotting to evaluate the ability of the NadR mutants to repress the nadA promoter, in the presence or absence of 4-HPA. The nadR H7A, S9A and F25A complemented strains showed hyper-repression of nadA expression in vivo, i.e. these mutants repressed nadA more efficiently than the NadR WT protein, either in the presence or absence of 4-HPA, while complementation with wild-type nadR resulted in high production of NadA only in the presence of 4-HPA (Fig 9). Interestingly, and on the contrary, the nadR N11A complemented strain showed hypo-repression (i.e. exhibited high expression of nadA both in absence and presence of 4-HPA). This mutagenesis data revealed that NadR residues His7, Ser9, Asn11 and Phe25 play key roles in the ligand-mediated regulation of NadR; they are each involved in the controlled de-repression of the nadA promoter and synthesis of NadA in response to 4-HPA in vivo. Structure-based point mutations shed light on ligand-induced regulation of NadR. Western blot analyses of wild-type (WT) strain (lanes 1–2) or isogenic nadR knockout strains (ΔNadR) complemented to express the indicated NadR WT or mutant proteins (lanes 3–12) or not complemented (lanes 13–14), grown in the presence (even lanes) or absence (odd lanes) of 5mM 4-HPA, showing NadA and NadR expression. Complementation of ΔNadR with WT NadR enables induction of nadA expression by 4-HPA. The H7A, S9A and F25A mutants efficiently repress nadA expression but are less ligand-responsive than WT NadR. The N11A mutant does not efficiently repress nadA expression either in presence or absence of 4-HPA. (The protein abundance levels of the meningococcal factor H binding protein (fHbp) were used as a gel loading control). Discussion NadA is a surface-exposed meningococcal protein contributing to pathogenesis, and is one of three main antigens present in the vaccine Bexsero. A detailed understanding of the in vitro repression of nadA expression by the transcriptional regulator NadR is important, both because it is a relevant disease-related model of how small-molecule ligands can regulate MarR family proteins and thereby impact bacterial virulence, and because nadA expression levels are linked to the prediction of vaccine coverage. The repressive activity of NadR can be relieved by hydroxyphenylacetate (HPA) ligands, and HDX-MS studies previously indicated that 4-HPA stabilizes dimeric NadR in a configuration incompatible with DNA binding. Despite these and other studies, the molecular mechanisms by which ligands regulate MarR family proteins are relatively poorly understood and likely differ depending on the specific ligand. Given the importance of NadR-mediated regulation of NadA levels in the contexts of meningococcal pathogenesis, we sought to characterize NadR, and its interaction with ligands, at atomic resolution. Firstly, we confirmed that NadR is dimeric in solution and demonstrated that it retains its dimeric state in the presence of 4-HPA, indicating that induction of a monomeric status is not the manner by which 4-HPA regulates NadR. These observations were in agreement with (i) a previous study of NadR performed using SEC and mass spectrometry, and (ii) crystallographic studies showing that several MarR homologues are dimeric. We also used structure-guided site-directed mutagenesis to identify an important conserved residue, Leu130, which stabilizes the NadR dimer interface, knowledge of which may also inform future studies to explore the regulatory mechanisms of other MarR family proteins. Secondly, we assessed the thermal stability and unfolding of NadR in the presence or absence of ligands. All DSC profiles showed a single peak, suggesting that a single unfolding event simultaneously disrupted the dimer and the monomer. HPA ligands specifically increased the stability of NadR. The largest effects were induced by the naturally-occurring compounds 4-HPA and 3Cl,4-HPA, which, in SPR assays, were found to bind NadR with KD values of 1.5 mM and 1.1 mM, respectively. Although these NadR/HPA interactions appeared rather weak, their distinct affinities and specificities matched their in vitro effects and their biological relevance appears similar to previous proposals that certain small molecules, including some antibiotics, in the millimolar concentration range may be broad inhibitors of MarR family proteins. Indeed, 4-HPA is found in human saliva and 3Cl,4-HPA is produced during inflammatory processes, suggesting that these natural ligands are encountered by N. meningitidis in the mucosa of the oropharynx during infections. It is also possible that NadR responds to currently unidentified HPA analogues. Indeed, in the NadR/4-HPA complex there was a water molecule close to the carboxylate group and also a small unfilled tunnel ~5Å long, both factors suggesting that alternative larger ligands could occupy the pocket. It is conceivable that such putative ligands may establish different bonding networks, potentially binding in a 2:2 ratio, rather than the 1:2 ratio observed herein. The ability to respond to various ligands might enable NadR in vivo to orchestrate multiple response mechanisms and modulate expression of genes other than nadA. Ultimately, confirmation of the relevance of each ligand will require a deeper understanding of the available concentration in vivo in the host niche during bacterial colonization and inflammation. Here, we determined the first crystal structures of apo-NadR and holo-NadR. These experimentally-determined structures enabled a new detailed characterization of the ligand-binding pocket. In holo-NadR, 4-HPA interacted directly with at least 11 polar and hydrophobic residues. Several, but not all, of these interactions were predicted previously by homology modelling combined with ligand docking in silico. Subsequently, we established the functional importance of His7, Ser9, Asn11 and Phe25 in the in vitro response of meningococcus to 4-HPA, via site-directed mutagenesis. More unexpectedly, the crystal structure revealed that only one molecule of 4-HPA was bound per NadR dimer. We confirmed this stoichiometry in solution using SPR methods. We also used heteronuclear NMR spectroscopy to detect substantial conformational changes of NadR occurring in solution upon addition of 4-HPA. Moreover, NMR spectra at 10°C suggested the existence of two distinct conformations of NadR in the vicinity of the ligand-binding pocket. More powerfully, our unique crystallographic observation of this ‘occupied vs unoccupied site’ asymmetry in the NadR/4-HPA interaction is, to our knowledge, the first example reported for a MarR family protein. Structural analyses suggested that ‘inward’ side chain positions of Met22, Phe25 and especially Arg43 precluded binding of a second ligand molecule. Such a mechanism indicates negative cooperativity, which may enhance the ligand-responsiveness of NadR. Comparisons of the NadR/4-HPA complex with available MarR family/salicylate complexes revealed that 4-HPA has a previously unobserved binding mode. Briefly, in the M. thermoautotrophicum MTH313 dimer, one molecule of salicylate binds in the pocket of each monomer, though with two rather different positions and orientations, only one of which (site-1) is thought to be biologically relevant (Fig 10A). In the S. tokodaii protein ST1710, salicylate binds to the same position in each monomer of the dimer, in a site equivalent to the putative biologically relevant site of MTH313 (Fig 10B). Unlike other MarR family proteins which revealed multiple ligand binding interactions, we observed only 1 molecule of 4-HPA bound to NadR, suggesting a more specific and less promiscuous interaction. In NadR, the single molecule of 4-HPA binds in a position distinctly different from the salicylate binding site: translated by \u003e 10 Å and with a 180° inverted orientation (Fig 10C). NadR shows a ligand binding site distinct from other MarR homologues. \n(A) A structural alignment of MTH313 chains A and B shows that salicylate is bound in distinct locations in each monomer; site-1 (thought to be the biologically relevant site) and site-2 differ by ~7Å (indicated by black dotted line) and also by ligand orientation. (B) A structural alignment of MTH313 chain A and ST1710 (pink) (Cα rmsd 2.3Å), shows that they bind salicylate in equivalent sites (differing by only ~3Å) and with the same orientation. (C) Addition of holo-NadR (chain B, blue) to the alignment reveals that bound 4-HPA differs in position by \u003e 10 Å compared to salicylate, and adopts a novel orientation. Interestingly, a crystal structure was previously reported for a functionally-uncharacterized meningococcal homologue of NadR, termed NMB1585, which shares 16% sequence identity with NadR. The two structures can be closely aligned (rmsd 2.3 Å), but NMB1585 appears unsuited for binding HPAs, since its corresponding ‘pocket’ region is occupied by several bulky hydrophobic side chains. It can be speculated that MarR family members have evolved separately to engage distinct signaling molecules, thus enabling bacteria to use the overall conserved MarR scaffold to adapt and respond to diverse changing environmental stimuli experienced in their natural niches. Alternatively, it is possible that other MarR homologues (e.g. NMB1585) may have no extant functional binding pocket and thus may have lost the ability to respond to a ligand, acting instead as constitutive DNA-binding regulatory proteins. The apo-NadR crystal structures revealed two dimers with slightly different conformations, most divergent in the DNA-binding domain. It is not unusual for a crystal structure to reveal multiple copies of the same protein in very slightly different conformations, which are likely representative of the lowest-energy conformations sampled by the dynamic ensemble of molecular states occurring in solution, and which likely have only small energetic differences, as described previously for MexR (a MarR protein) or more recently for the solute-binding protein FhuD2. Further, the holo-NadR structure was overall more different from the two apo-NadR structures (rmsd values ~1.3Å), suggesting that the ligand selected and stabilized yet another conformation of NadR. These observations suggest that 4-HPA, and potentially other similar ligands, can shift the molecular equilibrium, changing the energy barriers that separate active and inactive states, and stabilizing the specific conformation of NadR poorly suited to bind DNA. Comparisons of the apo- and holo-NadR structures revealed that the largest differences occurred in the DNA-binding helix α4. The shift of helix α4 in holo-NadR was also accompanied by rearrangements at the dimer interface, involving helices α1, α5, and α6, and this holo-form appeared poorly suited for DNA-binding when compared with the known OhrR:ohrA complex. While some flexibility of helix α4 was also observed in the two apo-structures, concomitant changes in the dimer interfaces were not observed, possibly due to the absence of ligand. One of the two conformations of apo-NadR appeared ideally suited for DNA-binding. Overall, these analyses suggest that the apo-NadR dimer has a pre-existing equilibrium that samples a variety of conformations, some of which are compatible with DNA binding. This intrinsically dynamic nature underlies the possibility for different conformations to inter-convert or to be preferentially selected by a regulatory ligand, as generally described in the ‘conformational selection’ model for protein-ligand interactions (the Monod-Wyman-Changeux model), rather than an ‘induced fit’ model (Koshland-Nemethy-Filmer). The noted flexibility may also explain how NadR can adapt to bind various DNA target sequences with slightly different structural features. Subsequently, upon ligand binding, holo-NadR adopts a structure less suited for DNA-binding and this conformation is selected and stabilized by a network of protein-ligand interactions and concomitant rearrangements at the NadR holo dimer interface. In an alternative and less extensive manner, the binding of two salicylate molecules to the M. thermoautotrophicum protein MTH313 appeared to induce large changes in the wHTH domain, which was associated with reduced DNA-binding activity. Here we have presented two new crystal structures of the transcription factor, NadR, which regulates expression of the meningococcal surface protein, virulence factor and vaccine antigen NadA. Detailed structural analyses provided a molecular explanation for the ligand-responsive regulation by NadR on the majority of the promoters of meningococcal genes regulated by NadR, including nadA. Intriguingly, NadR exhibits a reversed regulatory mechanism on a second class of promoters, including mafA of the multiple adhesin family–i.e. NadR represses these genes in the presence but not absence of 4-HPA. The latter may influence the surface abundance or secretion of maf proteins, an emerging class of highly conserved meningococcal putative adhesins and toxins with many important roles. Further work is required to investigate how the two different promoter types influence the ligand-responsiveness of NadR during bacterial infection and may provide insights into the regulatory mechanisms occurring during these host-pathogen interactions. Ultimately, knowledge of the ligand-dependent activity of NadR will continue to deepen our understanding of nadA expression levels, which influence meningococcal pathogenesis. Materials and Methods Bacterial strains, culture conditions and mutant generation In this study we used N. meningitidis MC58 wild type strain and related mutant derivatives. The MC58 isolate was kindly provided to us by Professor E. Richard Moxon, University of Oxford, UK, and was previously submitted to the Meningococcal Reference Laboratory, Manchester, UK. Strains were routinely cultured, stocked, and transformed as described previously. To generate N. meningitidis MC58 mutant strains expressing only the amino acid substituted forms of NadR, plasmids containing the sequence of nadR mutated to insert alanine codons to replace His7, Ser9, Asn11 or Phe25 were constructed using the QuikChange II XL Site-Directed Mutagenesis Kit (Stratagene). The nadR gene (also termed NMB1843) was mutated in the pComEry-1843 plasmid using couples of mutagenic primers (forward and reverse). The resulting plasmids were named pComEry-1843H7A, -1843S9A, -1843N11A or -1843F25A, and contained a site-directed mutant allele of the nadR gene, in which the selected codons were respectively substituted by a GCG alanine codon, and were used for transformation of the MC-Δ1843 strain. Total lysates from single colonies of all transformants were used as a template for PCR analysis to confirm the correct insertion by double homologous recombinant event. When indicated, bacterial strains were grown in presence of 5 mM 4-HPA (MW 152, Sigma-Aldrich). Molecular cloning The preparation of the expression construct enabling production of soluble NadR with an N-terminal His-tag followed by a thrombin cleavage site (MGSSHHHHHHSSGLVPR↓GSH-) (where the arrow indicates the cleavage site) and then NadR residues M1-S146 (Uniprot code Q7DD70), and methods to generate site-directed mutants, were described previously. Briefly, site-directed mutagenesis was performed using two overlapping primers containing the desired mutation to amplify pET15b containing several NadR variants. (Full oligonucleotide sequences of primers are available upon request). 1–10 ng of plasmid DNA template were amplified using Kapa HiFi DNA polymerase (Kapa Biosystems) and with the following cycling conditions: 98°C for 5 min, 15 cycles of (98°C for 30 s, 60°C for 30 s, 72°C for 6 min) followed by a final extension of 10 min at 72°C. Residual template DNA was digested by 30 min incubation with FastDigest DpnI (Thermo Scientific) at 37°C and 1 μl of this reaction was used to transform E. coli DH5α competent cells. The full recombinant tagged NadR protein generated contained 166 residues, with a theoretical MW of 18746, while after thrombin-cleavage the untagged protein contained 149 residues, with a theoretical MW of 16864. Protein production and purification The NadR expression constructs (wild-type or mutant clones) were transformed into E. coli BL21 (DE3) cells and were grown at 37°C in Luria-Bertani (LB) medium supplemented with 100 μg/mL ampicillin, until an OD600 of 0.5 was reached. Target protein production was induced by the addition of 1 mM IPTG followed by incubation with shaking overnight at 21°C. For production of the selenomethionine (SeMet) derivative form of NadR for crystallization studies, essentially the same procedure was followed, but using the E. coli B834 strain grown in a modified M9 minimal medium supplemented with 40 mg/L L-SeMet. For production of 15N-labeled NadR for NMR analyses, the EnPresso B Defined Nitrogen-free medium (Sigma-Aldrich) was used; in brief, BL21 (DE3) cells were grown in BioSilta medium at 30°C for 30 h, and production of the 15N-labeled NadR was enabled by the addition of 2.5 g/L 15NH4Cl and further incubation for 2 days. Cells were harvested by centrifugation (6400 g, 30 min, 4°C), resuspended in 20 mM HEPES pH 8.0, 300 mM NaCl, 20 mM imidazole, and were lysed by sonication (Qsonica Q700). Cell lysates were clarified by centrifugation at 2800 g for 30 min, and the supernatant was filtered using a 0.22 μm membrane (Corning filter system) prior to protein purification. NadR was purified by affinity chromatography using an AKTA purifier (GE Healthcare). All steps were performed at room temperature (18–26°C), unless stated otherwise. The filtered supernatant was loaded onto an Ni-NTA resin (5 mL column, GE Healthcare), and NadR was eluted using 4 steps of imidazole at 20, 30, 50 and 250 mM concentration, at a flow rate of 5 mL/min. Eluted fractions were examined by reducing and denaturing SDS-PAGE analysis. Fractions containing NadR were identified by a band migrating at ~17 kDa, and were pooled. The N-terminal 6-His tag was removed enzymatically using the Thrombin CleanCleave Kit (Sigma-Aldrich). Subsequently, the sample was reloaded on the Ni-NTA resin to capture the free His tag (or unprocessed tagged protein), thus allowing elution in the column flow-through of tagless NadR protein, which was used in all subsequent studies. The NadR sample was concentrated and loaded onto a HiLoad Superdex 75 (16/60) preparative size-exclusion chromatography (SEC) column equilibrated in buffer containing 20 mM HEPES pH 8.0, 150 mM NaCl, at a flow-rate of 1 mL/min. NadR protein was collected and the final yield of purified protein obtained from 0.5 L LB growth medium was approximately 8 mg (~2 mg protein per g wet biomass). Samples were used immediately for crystallization or analytical experiments, or were frozen for storage at -20°C. SE-HPLC/MALLS analyses MALLS analyses were performed online with SE-HPLC using a Dawn TREOS MALLS detector (Wyatt Corp., Santa Barbara, CA, USA) and an incident laser wavelength of 658 nm. The intensity of the scattered light was measured at 3 angles simultaneously. Data analysis was performed using the Astra V software (Wyatt) to determine the weighted-average absolute molecular mass (MW), the polydispersity index (MW/Mn) and homogeneity (Mz/Mn) for each oligomer present in solution. Normalization of the MALLS detectors was performed in each analytical session by use of bovine serum albumin. Differential scanning calorimetry The thermal stability of NadR proteins was assessed by differential scanning calorimetry (DSC) using a MicroCal VP-Capillary DSC instrument (GE Healthcare). NadR samples were prepared at a protein concentration of 0.5 mg/mL (~30 μM) in buffer containing 20 mM HEPES, 300 mM NaCl, pH 7.4, with or without 6 mM HPA or salicylate. The DSC temperature scan ranged from 10°C to 110°C, with a thermal ramping rate of 200°C per hour and a 4 second filter period. Data were analyzed by subtraction of the reference data for a sample containing buffer only, using the Origin 7 software. All experiments were performed in triplicate, and mean values of the melting temperature (Tm) were determined. Surface plasmon resonance (SPR) Determination of equilibrium dissociation constant, KD  Surface plasmon resonance binding analyses were performed using a Biacore T200 instrument (GE Healthcare) equilibrated at 25°C. The ligand (NadR) was covalently immobilized by amine-coupling on a CM-5 sensor chip (GE Healthcare), using 20 μg/mL purified protein in 10 mM sodium acetate buffer pH 5, injected at 10 μl/min for 120 s until ~9000 response units (RU) were captured. A high level of ligand immobilization was required due to the small size of the analytes. An unmodified surface was used as the reference channel. Titrations with analytes (HPAs or salicylate) were performed with a flow-rate of 30 μl/min, injecting the compounds in a concentration range of 10 μM to 20 mM, using filtered running buffer containing Phosphate Buffered Saline (PBS) with 0.05% Tween-20, pH 7.4. Following each injection, sensor chip surfaces were regenerated with a 30-second injection of 10 mM Glycine pH 2.5. Each titration series contained 20 analyte injections and was performed in triplicate. Titration experiments with long injection phases (\u003e 15 mins) were used to enable steady-state analyses. Data were analyzed using the BIAcore T200 evaluation software and the steady-state affinity model. A buffer injection was subtracted from each curve, and reference sensorgrams were subtracted from experimental sensorgrams to yield curves representing specific binding. The equilibrium dissociation constant, KD, was determined from the plot of RUeq against analyte concentration (S2 Fig), as described previously. Determination of binding stoichiometry: From each plot of RUeq against analyte concentration, obtained from triplicate experiments, the Rmax value (maximum analyte binding capacity of the surface) was extrapolated from the experimental data (S2 Fig). Stoichiometry was calculated using the molecular weight of dimeric NadR as ligand molecule (MWligand) and the molecular weights of the HPA analyte molecules (MWanalyte), and the following equation:  where Rligand is recorded directly from the sensorgram during ligand immobilization prior to the titration series, as described previously. The stoichiometry derived therefore represented the number of HPA molecules bound to one dimeric NadR protein. Crystallization of NadR in the presence or absence of 4-HPA Purified NadR was concentrated to 2.7 mg/mL (~160 μM) using a centrifugal concentration device (Amicon Ultra-15 Centrifugal Filter Unit with Ultracel-10 membrane with cut-off size 10 kDa; Millipore) running at 600 g in a bench top centrifuge (Thermo Scientific IEC CL40R) refrigerated at 2–8°C. To prepare holo-NadR samples, HPA ligands were added at a 200-fold molar excess prior to the centrifugal concentration step. The concentrated holo- or apo-NadR was subjected to crystallization trials performed in 96-well low-profile Intelli-Plates (Art Robbins) or 96-well low-profile Greiner crystallization plates, using a nanodroplet sitting-drop vapour-diffusion format and mixing equal volumes (200 nL) of protein samples and crystallization buffers using a Gryphon robot (Art Robbins). Crystallization trays were incubated at 20°C. Crystals of apo-NadR were obtained in 50% PEG 3350 and 0.13 M di-Ammonium hydrogen citrate, whereas crystals of SeMet–NadR in complex with 4-HPA grew in condition H4 of the Morpheus screen (Molecular Dimensions), which contains 37.5% of the pre-mixed precipitant stock MPD_P1K_PEG 3350, buffer system 1 and 0.1 M amino acids, at pH 6.5. All crystals were mounted in cryo-loops using 10% ethylene glycol or 10% glycerol as cryo-protectant before cooling to 100 K for data collection. X-ray diffraction data collection and structure determination X-ray diffraction data from crystals of apo-NadR and SeMet–NadR/4-HPA were collected on beamline PXII-X10SA of the Swiss Light Source (SLS) at the Paul Scherrer Institut (PSI), Villigen, Switzerland. All diffraction data were processed with XDS and programs from the CCP4 suite. Crystals of apo-NadR and 4-HPA-bound SeMet-NadR belonged to space group P43 21 2 (see Table 2). Apo-NadR crystals contained four molecules (two dimers) in the asymmetric unit (Matthews coefficient 2.25 Å3 Da−1, for a solvent content of 45%), while crystals of SeMet–NadR/4-HPA contained two molecules (one dimer) in the asymmetric unit (Matthews coefficient 1.98 Å3 Da−1, for a solvent content of 38%). In solving the holo-NadR structure, an initial and marginal molecular replacement (MR) solution was obtained using as template search model the crystal structure of the transcriptional regulator PA4135 (PBD entry 2FBI), with which NadR shares ~54% sequence identity. This solution was combined with SAD data to aid identification of two selenium sites in NadR, using autosol in phenix and this allowed generation of high-quality electron density maps that were used to build and refine the structure of the complex. Electron densities were clearly observed for almost the entire dimeric holo-NadR protein, except for a new N-terminal residues and residues 88–90 of chain B. The crystal structure of apo-NadR was subsequently solved by MR in Phaser at 2.7 Å, using the final refined model of SeMet-NadR/4-HPA as the search model. For apo-NadR, electron densities were clearly observed for almost the entire protein, although residues 84–91 of chains A, C, and D, and residues 84–90 of chain B lacked densities suggesting local disorder. Both structures were refined and rebuilt using phenix and Coot, and structural validation was performed using Molprobity. Data collection and refinement statistics are reported in Table 2. Atomic coordinates of the two NadR structures have been deposited in the Protein Data Bank, with entry codes 5aip (NadR bound to 4-HPA) and 5aiq (apo-NadR). All crystallographic software was compiled, installed and maintained by SBGrid. NMR spectroscopy For heteronuclear NMR experiments, the NadR protein concentration used was 85 μM (~ 1.4 mg/mL) in a solution containing 100 mM sodium phosphate buffer (90% H2O and 10% D2O) and 200 mM NaCl, prepared in the apo-form or in the presence of a 200-fold molar excess of 4-HPA, at pH 6.5. The stability, integrity and dimeric state of the protein in the NMR buffer was confirmed by analytical SEC (Superdex 75, 10/300 column) prior to NMR studies. 1H-15N transverse relaxation-optimized spectroscopy (TROSY)-heteronuclear single quantum coherence (HSQC) experiments on apo-NadR and NadR in the presence of 4-HPA were acquired using an Avance 950 Bruker spectrometer, operating at a proton frequency of 949.2 MHz and equipped with triple resonance cryogenically-cooled probe at two different temperatures (298 K and 283 K). 1H-15N TROSY-HSQC experiments were recorded for 12 h, with a data size of 1024 x 232 points. Spectra were processed using the Bruker TopSpin 2.1 and 3.1 software packages. Western blot Western blot analysis was performed as described previously. Supporting Information References Identification of vaccine candidates against serogroup B meningococcus by whole-genome sequencing  Neisseria adhesin A variation and revised nomenclature scheme  Neisseria meningitidis NadA is a new invasin which promotes bacterial adhesion to and penetration into human epithelial cells NadA, a novel vaccine candidate of Neisseria meningitidis  Structure of the meningococcal vaccine antigen NadA and epitope mapping of a bactericidal antibody A multi-component meningococcal serogroup B vaccine (4CMenB): the clinical development program Expression of the meningococcal adhesin NadA is controlled by a transcriptional regulator of the MarR family . MtrR control of a transcriptional regulatory pathway in Neisseria meningitidis that influences expression of a gene (nadA) encoding a vaccine candidate A novel phase variation mechanism in the meningococcus driven by a ligand-responsive repressor and differential spacing of distal promoter elements In the NadR Regulon, Adhesins and Diverse Meningococcal Functions Are Regulated in Response to Signals in Human Saliva A new family of secreted toxins in pathogenic Neisseria species Emergence of a new epidemic Neisseria meningitidis serogroup A Clone in the African meningitis belt: high-resolution picture of genomic changes that mediate immune evasion The mar regulon: multiple resistance to antibiotics and other toxic chemicals MarR family transcription factors Regulation of virulence by members of the MarR/SlyA family Molecular mechanisms of ligand-mediated attenuation of DNA binding by MarR family transcriptional regulators Structural insight on the mechanism of regulation of the MarR family of proteins: high-resolution crystal structure of a transcriptional repressor from Methanobacterium thermoautotrophicum  ST1710-DNA complex crystal structure reveals the DNA binding mechanism of the MarR family of regulators The presence of 4-hydroxyphenylacetic acid in human saliva and the possibility of its nitration by salivary nitrite in the stomach Transcriptional regulation of the nadA gene in Neisseria meningitidis impacts the prediction of coverage of a multicomponent meningococcal serogroup B vaccine Structural Insight into the Mechanism of DNA-Binding Attenuation of the Neisserial Adhesin Repressor NadR by the Small Natural Ligand 4-Hydroxyphenylacetic Acid Binding of purified multiple antibiotic-resistance repressor protein (MarR) to mar operator sequences Inference of macromolecular assemblies from crystalline state BiaCore analysis of leptin-leptin receptor interaction: evidence for 1:1 stoichiometry Analytical shape computation of macromolecules: I. Molecular area and volume through alpha shape Structure of an OhrR-ohrA operator complex reveals the DNA binding mechanism of the MarR family Qualitative and quantitative assessment of meningococcal antigens to evaluate the potential strain coverage of protein-based vaccines The metabolism and dechlorination of chlorotyrosine in vivo The structure of NMB1585, a MarR-family regulator from Neisseria meningitidis  Crystal structure of the MexR repressor of the mexRAB-oprM multidrug efflux operon of Pseudomonas aeruginosa  Structural and functional characterization of the Staphylococcus aureus virulence factor and vaccine candidate FhuD2 Crystal and solution structure analysis of FhuD2 from Staphylococcus aureus in multiple unliganded conformations and bound to ferrioxamine-B Conformational selection or induced fit? 50 years of debate resolved Point mutation in meningococcal por A gene associated with increased endemic disease Mechanistic implications for LDL receptor degradation from the PCSK9/LDLR structure at neutral pH Xds The CCP4 suite: programs for protein crystallography PHENIX: a comprehensive Python-based system for macromolecular structure solution Phaser crystallographic software Features and development of Coot MolProbity: all-atom structure validation for macromolecular crystallography Collaboration gets the most out of software Scaling and assessment of data quality PDBsum 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+[{"sourceid":"4848090","sourcedb":"","project":"","target":"","text":"Mechanistic insight into a peptide hormone signaling complex mediating floral organ abscission Plants constantly renew during their life cycle and thus require to shed senescent and damaged organs. Floral abscission is controlled by the leucine-rich repeat receptor kinase (LRR-RK) HAESA and the peptide hormone IDA. It is unknown how expression of IDA in the abscission zone leads to HAESA activation. Here we show that IDA is sensed directly by the HAESA ectodomain. Crystal structures of HAESA in complex with IDA reveal a hormone binding pocket that accommodates an active dodecamer peptide. A central hydroxyproline residue anchors IDA to the receptor. The HAESA co-receptor SERK1, a positive regulator of the floral abscission pathway, allows for high-affinity sensing of the peptide hormone by binding to an Arg-His-Asn motif in IDA. This sequence pattern is conserved among diverse plant peptides, suggesting that plant peptide hormone receptors may share a common ligand binding mode and activation mechanism. DOI: http://dx.doi.org/10.7554/eLife.15075.001 eLife digest Plants can shed their leaves, flowers or other organs when they no longer need them. But how does a leaf or a flower know when to let go? A receptor protein called HAESA is found on the surface of the cells that surround a future break point on the plant. When its time to shed an organ, a hormone called IDA instructs HAESA to trigger the shedding process. However, the molecular details of how IDA triggers organ shedding are not clear. The shedding of floral organs (or leaves) can be easily studied in a model plant called Arabidopsis. Santiago et al. used protein biochemistry, structural biology and genetics to uncover how the IDA hormone activates HAESA. The experiments show that IDA binds directly to a canyon shaped pocket in HAESA that extends out from the surface of the cell. IDA binding to HAESA allows another receptor protein called SERK1 to bind to HAESA, which results in the release of signals inside the cell that trigger the shedding of organs. The next step following on from this work is to understand what signals are produced when IDA activates HAESA. Another challenge will be to find out where IDA is produced in the plant and what causes it to accumulate in specific places in preparation for organ shedding. DOI: http://dx.doi.org/10.7554/eLife.15075.002 Introduction The HAESA ectodomain folds into a superhelical assembly of 21 leucine-rich repeats. (A) SDS PAGE analysis of the purified Arabidopsis thaliana HAESA ectodomain (residues 20–620) obtained by secreted expression in insect cells. The calculated molecular mass is 65.7 kDa, the actual molecular mass obtained by mass spectrometry is 74,896 Da, accounting for the N-glycans. (B) Ribbon diagrams showing front (left panel) and side views (right panel) of the isolated HAESA LRR domain. The N- (residues 20–88) and C-terminal (residues 593–615) capping domains are shown in yellow, the central 21 LRR motifs are in blue and disulphide bonds are highlighted in green (in bonds representation). (C) Structure based sequence alignment of the 21 leucine-rich repeats in HAESA with the plant LRR consensus sequence shown for comparison. Conserved hydrophobic residues are shaded in gray, N-glycosylation sites visible in our structures are highlighted in blue, cysteine residues involved in disulphide bridge formation in green. (D) Asn-linked glycans mask the N-terminal portion of the HAESA ectodomain. Oligomannose core structures (containing two N-actylglucosamines and three terminal mannose units) as found in Trichoplusia ni cells and in plants were modeled onto the seven glycosylation sites observed in our HAESA structures, to visualize the surface areas potentially not masked by carbohydrate. The HAESA ectodomain is shown in blue (in surface representation), the glycan structures are shown in yellow. Molecular surfaces were calculated with the program MSMS, with a probe radius of 1.5 Å. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.004 Hydrophobic contacts and a hydrogen-bond network mediate the interaction between HAESA and the peptide hormone IDA. (A) Details of the IDA binding pocket. HAESA is shown in blue (ribbon diagram), the C-terminal Arg-His-Asn motif (left panel), the central Hyp anchor (center) and the N-terminal Pro-rich motif in IDA (right panel) are shown in yellow (in bonds representation). HAESA interface residues are shown as sticks, selected hydrogen bond interactions are denoted as dotted lines (in magenta). (B) View of the complete IDA (in bonds representation, in yellow) binding pocket in HAESA (surface view, in blue). Orientation as in (A). (C) Structure based sequence alignment of leucine-rich repeats in HAESA with the plant LRR consensus sequence shown for comparison. Residues mediating hydrophobic interactions with the IDA peptide are highlighted in blue, residues contributing to hydrogen bond interactions and/or salt bridges are shown in red. The IDA binding pocket covers LRRs 2–14 and all residues originate from the inner surface of the HAESA superhelix. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.005 The IDA-HAESA and SERK1-HAESA complex interfaces are conserved among HAESA and HAESA-like proteins from different plant species. Structure-based sequence alignment of the HAESA family members: Arabidopsis thaliana HAESA (Uniprot (http://www.uniprot.org) ID P47735), Arabidopsis thaliana HSL2 (Uniprot ID C0LGX3), Capsella rubella HAESA (Uniprot ID R0F2U6), Citrus clementina HSL2 (Uniprot ID V4U227), Vitis vinifera HAESA (Uniprot ID F6HM39). The alignment includes a secondary structure assignment calculated with the program DSSP and colored according to Figure 1, with the N- and C-terminal caps and the 21 LRR motifs indicated in orange and blue, respectively. Cysteine residues engaged in disulphide bonds are depicted in green. HAESA residues interacting with the IDA peptide and/or the SERK1 co-receptor kinase ectodomain are highlighted in blue and orange, respectively. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.006 The peptide hormone IDA binds to the HAESA LRR ectodomain. (A) Multiple sequence alignment of selected IDA family members. The conserved PIP motif is highlighted in yellow, the central Hyp in blue. The PKGV motif present in our N-terminally extended IDA peptide is highlighted in red. (B) Isothermal titration calorimetry of the HAESA ectodomain vs. IDA and including the synthetic peptide sequence. (C) Structure of the HAESA – IDA complex with HAESA shown in blue (ribbon diagram). IDA (in bonds representation, surface view included) is depicted in yellow. The peptide binding pocket covers HAESA LRRs 2–14. (D) Close-up view of the entire IDA (in yellow) peptide binding site in HAESA (in blue). Details of the interactions between the central Hyp anchor in IDA and the C-terminal Arg-His-Asn motif with HAESA are highlighted in (E) and (F), respectively. Hydrogren bonds are depicted as dotted lines (in magenta), a water molecule is shown as a red sphere. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.003 During their growth, development and reproduction plants use cell separation processes to detach no-longer required, damaged or senescent organs. Abscission of floral organs in Arabidopsis is a model system to study these cell separation processes in molecular detail. The LRR-RKs HAESA (greek: to adhere to) and HAESA-LIKE 2 (HSL2) redundantly control floral abscission. Loss-of-function of the secreted small protein INFLORESCENCE DEFICIENT IN ABSCISSION (IDA) causes floral organs to remain attached while its over-expression leads to premature shedding. Full-length IDA is proteolytically processed and a conserved stretch of 20 amino-acids (termed EPIP) can rescue the IDA loss-of-function phenotype (Figure 1A). It has been demonstrated that a dodecamer peptide within EPIP is able to activate HAESA and HSL2 in transient assays in tobacco cells. This sequence motif is highly conserved among IDA family members (IDA-LIKE PROTEINS, IDLs) and contains a central Pro residue, presumed to be post-translationally modified to hydroxyproline (Hyp; Figure 1A). The available genetic and biochemical evidence suggests that IDA and HAESA together control floral abscission, but it is poorly understood if IDA is directly sensed by the receptor kinase HAESA and how IDA binding at the cell surface would activate the receptor. Results IDA directly binds to the LRR domain of HAESA Active IDA-family peptide hormones are hydroxyprolinated dodecamers. Close-up views of (A) IDA, (B) the N-terminally extended PKGV-IDA and (C) IDL1 bound to the HAESA hormone binding pocket (in bonds representation, in yellow) and including simulated annealing 2Fo–Fc omit electron density maps contoured at 1.0 σ. Note that Pro58IDA and Leu67IDA are the first residues defined by electron density when bound to the HAESA ectodomain. (D) Table summaries for equilibrium dissociation constants (Kd), binding enthalpies (ΔH), binding entropies (ΔS) and stoichoimetries (N) for different IDA peptides binding to the HAESA ectodomain ( ± fitting errors; n.d. no detectable binding). (E) Structural superposition of the active IDA (in bonds representation, in gray) and IDL1 peptide (in yellow) hormones bound to the HAESA ectodomain. Root mean square deviation (r.m.s.d.) is 1.0 Å comparing 100 corresponding atoms. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.007 The receptor kinase SERK1 acts as a HAESA co-receptor and promotes high-affinity IDA sensing. (A) Petal break-strength assays measure the force (expressed in gram equivalents) required to remove the petals from the flower of serk mutant plants compared to haesa/hsl2 mutant and Col-0 wild-type flowers. Petal break-strength is measured from positions 1 to 8 along the primary inflorescence where positions 1 is defined as the flower at anthesis (n=15, bars=SD). This treatment-by-position balanced two-way layout was analyzed separately per position, because of the serious interaction, by means of a Dunnett-type comparison against the Col-0 control, allowing for heterogeneous variances. Petal break-strength was found significantly increased in almost all positions (indicated with a *) for haesa/hsl2 and serk1-1 mutant plants with respect to the Col-0 control. Calculations were performed in R (version 3.2.3). (B) Analytical size-exclusion chromatography. The HAESA LRR domain elutes as a monomer (black dotted line), as does the isolated SERK1 ectodomain (blue dotted line). A HAESA – IDA – SERK1 complex elutes as an apparent heterodimer (red line), while a mixture of HAESA and SERK1 yields two isolated peaks that correspond to monomeric HAESA and SERK1, respectively (black line). Void (V0) volume and total volume (Vt) are shown, together with elution volumes for molecular mass standards (A, Thyroglobulin, 669,000 Da; B, Ferritin, 440,00 Da, C, Aldolase, 158,000 Da; D, Conalbumin, 75,000 Da; E, Ovalbumin, 44,000 Da; F, Carbonic anhydrase, 29,000 Da). A SDS PAGE of the peak fractions is shown alongside. Purified HAESA and SERK1 are ~75 and ~28 kDa, respectively. (C) Isothermal titration calorimetry of wild-type and Hyp64→Pro IDA versus the HAESA and SERK1 ectodomains. The titration of IDA wild-type versus the isolated HAESA ectodomain from Figure 1B is shown for comparison (red line; n.d. no detectable binding) (D) Analytical size-exclusion chromatography in the presence of the IDA Hyp64→Pro mutant peptide reveals no complex formation between HAESA and SERK1 ectodomains. A SDS PAGE of the peak fractions is shown alongside. (E) In vitro kinase assays of the HAESA and SERK1 kinase domains. Wild-type HAESA and SERK1 kinase domains (KDs) exhibit auto-phosphorylation activities (lanes 1 + 3). Mutant (m) versions, which carry point mutations in their active sites (Asp837HAESA→Asn, Asp447SERK1→Asn) possess no autophosphorylation activity (lanes 2+4). Transphosphorylation activity from the active kinase to the mutated form can be observed in both directions (lanes 5+6). A coomassie-stained gel loading control is shown below. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.008 Crystallographic data collection, phasing and refinement statistics for the isolated A. thaliana HAESA ectodomain. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.009 \tHAESA NaI shortsoak\tHAESA apo\t \tPDB-ID\t\t5IXO\t \tData collection\t\t\t \tSpace group\tP31 21\tP31 21\t \tCell dimensions\t\t\t \ta, b, c (Å)\t148.55, 148.55, 58.30\t149.87, 149.87, 58.48\t \tα, β, γ (°)\t90, 90, 120\t90, 90, 120\t \tResolution (Å)\t48.63–2.39 (2.45–2.39)\t45.75–1.74 (1.85–1.74)\t \tRmeas#\t0.096 (0.866)\t0.038 (1.02)\t \tCC(1/2)#\t100/86.6\t100/75.6\t \tI/σ I#\t27.9 (4.9)\t18.7 (1.8)\t \tCompleteness (%)#\t99.9 (98.6)\t99.6 (97.4)\t \tRedundancy#\t53.1 (29.9)\t14.4 (14.0)\t \tWilson B-factor (Å2)#\t84.45\t81.10\t \tRefinement\t\t\t \tResolution (Å)\t\t45.75 – 1.74\t \tNo. reflections\t\t71,213\t \tRwork/Rfree$\t\t0.188/0.218\t \tNo. atoms\t\t\t \tProtein/glycan\t\t4,533/126\t \tWater\t\t71\t \tRes. B-factors (Å2)$\t\t\t \tProtein\t\t77.54\t \tGlycan\t\t95.98\t \tWater\t\t73.20\t \tR.m.s deviations$\t\t\t \tBond lengths (Å)\t\t0.0095\t \tBond angles (°)\t\t1.51\t \t Highest resolution shell is shown in parenthesis. #As defined in XDS. $As defined in Refmac5. Crystallographic data collection and refinement statistics for the HAESA – IDA, – PKGV-IDA, – IDL1 and – IDA – SERK1 complexes. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.010 \tHAESA – IDA\tHAESA – PKGV-IDA\tHAESA – IDL1\tHAESA – IDA – SERK1\t \tPDB-ID\t5IXQ\t5IXT\t5IYN\t5IYX\t \tData collection\t\t\t\t\t \tSpace group\tP31 21\tP31 21\tP31 21\tP212121\t \tCell dimensions\t\t\t\t\t \ta, b, c (Å)\t148.55, 148.55, 58.30\t148.92, 148.92, 58.02\t150.18, 150.18, 60.07\t74.51, 100.46, 142.76\t \tα, β, γ (°)\t90, 90, 120\t90, 90, 120\t90, 90, 120\t90, 90, 90\t \tResolution (Å)\t48.54–1.86 (1.97–1.86)\t48.75–1.94 (2,06–1.94)\t49.16–2.56 (2.72–2.56)\t47.59–2.43 (2.57–2.43)\t \tRmeas#\t0.057 (1.35)\t0.037 (0.97)\t0.056 (1.27)\t0.113 (1.37)\t \tCC(1/2)#\t100/77.9\t100/80.3\t100/89.5\t100/77.6\t \tI/σI#\t16.7 (2.0)\t20.9 (2.4)\t26.0 (1.9)\t16.12 (2.0)\t \tCompleteness# (%)\t99.8 (98.6)\t99.4 (97.9)\t99.5 (98.8)\t99.4 (96.4s\t \tRedundancy#\t20.3 (19.1)\t11.2 (11.1)\t14.7 (14.7)\t9.7 (9.3)\t \tWilson B-factor (Å2)#\t80.0\t81.7\t89.5\t59.3\t \tRefinement\t\t\t\t\t \tResolution (Å)\t48.54–1.86\t48.75–1.94\t49.16–2.56\t47.59–2.43\t \tNo. reflections\t58,551\t51,557\t23,835\t38,969\t \tRwork/Rfree$\t0.190/0.209\t0.183/0.208\t0.199/0.236\t0.199/0.235\t \tNo. atoms\t\t\t\t\t \tProtein/Glycan\t4,541/176\t4,545/176\t4,499/176\t5,965/168\t \tPeptide\t93\t93\t90\t112\t \tWater\t39\t40\t9\t136\t \tRes. B-factors (Å2)$\t\t\t\t\t \tProtein/Glycan\t79.48/109.02\t79.63/113.24\t102.12/132.49\t60.05/73.48\t \tPeptide\t87.19\t89.50\t125.74\t51.06\t \tWater\t75.32\t71.92\t74.65\t51.47\t \tR.m.s deviations$\t\t\t\t\t \tBond lengths (Å)\t0.0087\t0.0091\t0.0081\t0.0074\t \tBond angles (°)\t1.48\t1.47\t1.36\t1.34\t \t Highest resolution shell is shown in parenthesis. #As defined in XDS. $As defined in Refmac5. We purified the HAESA ectodomain (residues 20–620) from baculovirus-infected insect cells (Figure 1—figure supplement 1A, see Materials and methods) and quantified the interaction of the ~75 kDa glycoprotein with synthetic IDA peptides using isothermal titration calorimetry (ITC). A Hyp-modified dodecamer comprising the highly conserved PIP motif in IDA (Figure 1A) interacts with HAESA with 1:1 stoichiometry (N) and with a dissociation constant (Kd) of ~20 μM (Figure 1B). We next determined crystal structures of the apo HAESA ectodomain and of a HAESA-IDA complex, at 1.74 and 1.86 Å resolution, respectively (Figure 1C; Figure 1—figure supplement 1B–D; Tables 1,2). IDA binds in a completely extended conformation along the inner surface of the HAESA ectodomain, covering LRRs 2–14 (Figure 1C,D, Figure 1—figure supplement 2). The central Hyp64IDA is buried in a specific pocket formed by HAESA LRRs 8–10, with its hydroxyl group establishing hydrogen bonds with the strictly conserved Glu266HAESA and with a water molecule, which in turn is coordinated by the main chain oxygens of Phe289HAESA and Ser311HAESA (Figure 1E; Figure 1—figure supplement 3). The restricted size of the Hyp pocket suggests that IDA does not require arabinosylation of Hyp64IDA for activity in vivo, a modification that has been reported for Hyp residues in plant CLE peptide hormones. The C-terminal Arg-His-Asn motif in IDA maps to a cavity formed by HAESA LRRs 11–14 (Figure 1D,F). The COO- group of Asn69IDA is in direct contact with Arg407HAESA and Arg409HAESA and HAESA cannot bind a C-terminally extended IDA-SFVN peptide (Figures 1D,F, 2D). This suggests that the conserved Asn69IDA may constitute the very C-terminus of the mature IDA peptide in planta and that active IDA is generated by proteolytic processing from a longer pre-protein. Mutation of Arg417HSL2 (which corresponds to Arg409HAESA) causes a loss-of-function phenotype in HSL2, which indicates that the peptide binding pockets in different HAESA receptors have common structural and sequence features. Indeed, we find many of the residues contributing to the formation of the IDA binding surface in HAESA to be conserved in HSL2 and in other HAESA-type receptors in different plant species (Figure 1—figure supplement 3). A N-terminal Pro-rich motif in IDA makes contacts with LRRs 2–6 of the receptor (Figure 1D, Figure 1—figure supplement 2A–C). Other hydrophobic and polar interactions are mediated by Ser62IDA, Ser65IDA and by backbone atoms along the IDA peptide (Figure 1D, Figure 1—figure supplement 2A–C). HAESA specifically senses IDA-family dodecamer peptides We next investigated whether HAESA binds N-terminally extended versions of IDA. We obtained a structure of HAESA in complex with a PKGV-IDA peptide at 1.94 Å resolution (Table 2). In this structure, no additional electron density accounts for the PKGV motif at the IDA N-terminus (Figure 2A,B). Consistently, PKGV-IDA and IDA have similar binding affinities in our ITC assays, further indicating that HAESA senses a dodecamer peptide comprising residues 58-69IDA (Figure 2D). We next tested if HAESA binds other IDA peptide family members. IDL1, which can rescue IDA loss-of-function mutants when introduced in abscission zone cells, can also be sensed by HAESA, albeit with lower affinity (Figure 2D). A 2.56 Å co-crystal structure with IDL1 reveals that different IDA family members use a common binding mode to interact with HAESA-type receptors (Figure 2A–C,E, Table 2). We do not detect interaction between HAESA and a synthetic peptide missing the C-terminal Asn69IDA (ΔN69), highlighting the importance of the polar interactions between the IDA carboxy-terminus and Arg407HAESA/Arg409HAESA (Figures 1F, 2D). Replacing Hyp64IDA, which is common to all IDLs, with proline impairs the interaction with the receptor, as does the Lys66IDA/Arg67IDA → Ala double-mutant discussed below (Figure 1A, 2D). Notably, HAESA can discriminate between IDLs and functionally unrelated dodecamer peptides with Hyp modifications, such as CLV3 (Figures 2D, 7). The co-receptor kinase SERK1 allows for high-affinity IDA sensing Our binding assays reveal that IDA family peptides are sensed by the isolated HAESA ectodomain with relatively weak binding affinities (Figures 1B, 2A–D). It has been recently reported that SOMATIC EMBRYOGENESIS RECEPTOR KINASES (SERKs) are positive regulators of floral abscission and can interact with HAESA and HSL2 in an IDA-dependent manner. As all five SERK family members appear to be expressed in the Arabidopsis abscission zone, we quantified their relative contribution to floral abscission in Arabidopsis using a petal break-strength assay. Our experiments suggest that among the SERK family members, SERK1 is a positive regulator of floral abscission. We found that the force required to remove the petals of serk1-1 mutants is significantly higher than that needed for wild-type plants, as previously observed for haesa/hsl2 mutants, and that floral abscission is delayed in serk1-1 (Figure 3A). The serk2-2, serk3-1, serk4-1 and serk5-1 mutant lines showed a petal break-strength profile not significantly different from wild-type plants. Possibly because SERKs have additional roles in plant development such as in pollen formation and brassinosteroid signaling, we found that higher-order SERK mutants exhibit pleiotropic phenotypes in the flower, rendering their analysis and comparison by quantitative petal break-strength assays difficult. We thus focused on analyzing the contribution of SERK1 to HAESA ligand sensing and receptor activation. In vitro, the LRR ectodomain of SERK1 (residues 24–213) forms stable, IDA-dependent heterodimeric complexes with HAESA in size exclusion chromatography experiments (Figure 3B). We next quantified the contribution of SERK1 to IDA recognition by HAESA. We found that HAESA senses IDA with a ~60 fold higher binding affinity in the presence of SERK1, suggesting that SERK1 is involved in the specific recognition of the peptide hormone (Figure 3C). We next titrated SERK1 into a solution containing only the HAESA ectodomain. In this case, there was no detectable interaction between receptor and co-receptor, while in the presence of IDA, SERK1 strongly binds HAESA with a dissociation constant in the mid-nanomolar range (Figure 3C). This suggests that IDA itself promotes receptor – co-receptor association, as previously described for the steroid hormone brassinolide and for other LRR-RK complexes. Importantly, hydroxyprolination of IDA is critical for HAESA-IDA-SERK1 complex formation (Figure 3C,D). Our calorimetry experiments now reveal that SERKs may render HAESA, and potentially other receptor kinases, competent for high-affinity sensing of their cognate ligands. Upon IDA binding at the cell surface, the kinase domains of HAESA and SERK1, which have been shown to be active protein kinases, may interact in the cytoplasm to activate each other. Consistently, the HAESA kinase domain can transphosphorylate SERK1 and vice versa in in vitro transphosphorylation assays (Figure 3E). Together, our genetic and biochemical experiments implicate SERK1 as a HAESA co-receptor in the Arabidopsis abscission zone. SERK1 senses a conserved motif in IDA family peptides Crystal structure of a HAESA – IDA – SERK1 signaling complex. (A) Overview of the ternary complex with HAESA in blue (surface representation), IDA in yellow (bonds representation) and SERK1 in orange (surface view). (B) The HAESA ectodomain undergoes a conformational change upon SERK1 co-receptor binding. Shown are Cα traces of a structural superposition of the unbound (yellow) and SERK1-bound (blue) HAESA ectodomains (r.m.s.d. is 1.5 Å between 572 corresponding Cα atoms). SERK1 (in orange) and IDA (in red) are shown alongside. The conformational change in the C-terminal LRRs and capping domain is indicated by an arrow. (C) SERK1 forms an integral part of the receptor's peptide binding pocket. The N-terminal capping domain of SERK1 (in orange) directly contacts the C-terminal part of IDA (in yellow, in bonds representation) and the receptor HAESA (in blue). Polar contacts of SERK1 with IDA are shown in magenta, with the HAESA LRR domain in gray. (D) Details of the zipper-like SERK1-HAESA interface. Ribbon diagrams of HAESA (in blue) and SERK1 (in orange) are shown with selected interface residues (in bonds representation). Polar interactions are highlighted as dotted lines (in magenta). DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.011 To understand in molecular terms how SERK1 contributes to high-affinity IDA recognition, we solved a 2.43 Å crystal structure of the ternary HAESA – IDA – SERK1 complex (Figure 4A, Table 2). HAESA LRRs 16–21 and its C-terminal capping domain undergo a conformational change upon SERK1 binding (Figure 4B). The SERK1 ectodomain interacts with the IDA peptide binding site using a loop region (residues 51-59SERK1) from its N-terminal cap (Figure 4A,C). SERK1 loop residues establish multiple hydrophobic and polar contacts with Lys66IDA and the C-terminal Arg-His-Asn motif in IDA (Figure 4C). SERK1 LRRs 1–5 and its C-terminal capping domain form an additional zipper-like interface with residues originating from HAESA LRRs 15–21 and from the HAESA C-terminal cap (Figure 4D). SERK1 binds HAESA using these two distinct interaction surfaces (Figure 1—figure supplement 3), with the N-cap of the SERK1 LRR domain partially covering the IDA peptide binding cleft. The IDA C-terminal motif is required for HAESA-SERK1 complex formation and for IDA bioactivity. (A) Size exclusion chromatography experiments similar to Figure 3B,D reveal that IDA mutant peptides targeting the C-terminal motif do not form biochemically stable HAESA-IDA-SERK1 complexes. Deletion of the C-terminal Asn69IDA completely inhibits complex formation. Void (V0) volume and total volume (Vt) are shown, together with elution volumes for molecular mass standards (A, Thyroglobulin, 669,000 Da; B, Ferritin, 440,00 Da, C, Aldolase, 158,000 Da; D, Conalbumin, 75,000 Da; E, Ovalbumin, 44,000 Da; F, Carbonic anhydrase, 29,000 Da). Purified HAESA and SERK1 are ~75 and ~28 kDa, respectively. Left panel: IDA K66A/R67A; center: IDA ΔN69, right panel: SDS-PAGE of peak fractions. Note that the HAESA and SERK1 input lanes have already been shown in Figure 3D. (B) Isothermal titration thermographs of wild-type and mutant IDA peptides titrated into a HAESA - SERK1 mixture in the cell. Table summaries for calorimetric binding constants and stoichoimetries for different IDA peptides binding to the HAESA – SERK1 ectodomain mixture ( ± fitting errors; n.d. no detectable binding) are shown alongside. (C) Quantitative petal break-strength assay for Col-0 wild-type flowers and 35S::IDA wild-type and 35S::IDA K66A/R67A mutant flowers. Petal break is measured from positions 1 to 8 along the primary inflorescence where positions 1 is defined as the flower at anthesis (n=15, bars=SD). The three treatment groups in this unbalanced one-way layout were compared by Tukey’s all-pairs comparison procedure using the package multcomp in R (version 3.2.3). 35S::IDA plants showed significantly increased abscission compared to Col-0 controls in inflorescence positions 2 and 3 (a). Up to inflorescence position 4, petal break in 35S::IDA K66A/R67A mutant plants was significantly increased compared to both Col-0 control plants (b) and 35S::IDA plants (c) (D) Normalized expression levels (relative expression ± standard error; ida: -0.02 ± 0.001; Col-0: 1 ± 0.11; 35S::IDA 124 ± 0.75; 35S::IDA K66A/R67A: 159 ± 0.58) of IDA wild-type and mutant transcripts in the 35S promoter over-expression lines analyzed in (C). (E) Magnified view of representative abscission zones from 35S::IDA, Col-0 wild-type and 35S::IDA K66A/R67A double-mutant T3 transgenic lines. 15 out of 15 35S::IDA plants, 0 out of 15 Col-0 plants and 0 out of 15 35S::IDA K66A/R67A double-mutant plants, showed an enlarged abscission zone, respectively (3 independent lines were analyzed). DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.012 The four C-terminal residues in IDA (Lys66IDA-Asn69IDA) are conserved among IDA family members and are in direct contact with SERK1 (Figures 1A, 4C). We thus assessed their contribution to HAESA – SERK1 complex formation. Deletion of the buried Asn69IDA completely inhibits receptor – co-receptor complex formation and HSL2 activation (Figure 5A,B). A synthetic Lys66IDA/Arg67IDA → Ala mutant peptide (IDA K66A/R66A) showed a 10 fold reduced binding affinity when titrated in a HAESA/SERK1 protein solution (Figures 5A,B, 2D). We over-expressed full-length wild-type IDA or this Lys66IDA/Arg67IDA → Ala double-mutant to similar levels in Col-0 Arabidopsis plants (Figure 5D). We found that over-expression of wild-type IDA leads to early floral abscission and an enlargement of the abscission zone (Figure 5C–E). In contrast, over-expression of the IDA Lys66IDA/Arg67IDA → Ala double mutant significantly delays floral abscission when compared to wild-type control plants, suggesting that the mutant IDA peptide has reduced activity in planta (Figure 5C–E). Comparison of 35S::IDA wild-type and mutant plants further indicates that mutation of Lys66IDA/Arg67IDA → Ala may cause a weak dominant negative effect (Figure 5C–E). In agreement with our structures and biochemical assays, this experiment suggests a role of the conserved IDA C-terminus in the control of floral abscission. Discussion In contrast to animal LRR receptors, plant LRR-RKs harbor spiral-shaped ectodomains and thus they require shape-complementary co-receptor proteins for receptor activation. For a rapidly growing number of plant signaling pathways, SERK proteins act as these essential co-receptors (; ). SERK1 has been previously reported as a positive regulator in plant embryogenesis, male sporogenesis, brassinosteroid signaling and in phytosulfokine perception. Recent findings by and our mechanistic studies now also support a positive role for SERK1 in floral abscission. As serk1-1 mutant plants show intermediate abscission phenotypes when compared to haesa/hsl2 mutants, SERK1 likely acts redundantly with other SERKs in the abscission zone (Figure 3A). It has been previously suggested that SERK1 can inhibit cell separation. However our results show that SERK1 also can activate this process upon IDA sensing, indicating that SERKs may fulfill several different functions in the course of the abscission process. While the sequence of the mature IDA peptide has not been experimentally determined in planta, our HAESA-IDA complex structures and calorimetry assays suggest that active IDLs are hydroxyprolinated dodecamers. It will be thus interesting to see if proteolytic processing of full-length IDA in vivo is regulated in a cell-type or tissue-specific manner. The central Hyp residue in IDA is found buried in the HAESA peptide binding surface and thus this post-translational modification may regulate IDA bioactivity. Our comparative structural and biochemical analysis further suggests that IDLs share a common receptor binding mode, but may preferably bind to HAESA, HSL1 or HSL2 in different plant tissues and organs. In our quantitative biochemical assays, the presence of SERK1 dramatically increases the HAESA binding specificity and affinity for IDA. This observation is consistent with our complex structure in which receptor and co-receptor together form the IDA binding pocket. The fact that SERK1 specifically interacts with the very C-terminus of IDLs may allow for the rational design of peptide hormone antagonists, as previously demonstrated for the brassinosteroid pathway. Importantly, our calorimetry assays reveal that the SERK1 ectodomain binds HAESA with nanomolar affinity, but only in the presence of IDA (Figure 3C). This ligand-induced formation of a receptor – co-receptor complex may allow the HAESA and SERK1 kinase domains to efficiently trans-phosphorylate and activate each other in the cytoplasm. It is of note that our reported binding affinities for IDA and SERK1 have been measured using synthetic peptides and the isolated HAESA and SERK1 ectodomains, and thus might differ in the context of the full-length, membrane-embedded signaling complex. SERK1 uses partially overlapping surface areas to activate different plant signaling receptors. (A) Structural comparison of plant steroid and peptide hormone membrane signaling complexes. Left panel: Ribbon diagram of HAESA (in blue), SERK1 (in orange) and IDA (in bonds and surface represention). Right panel: Ribbon diagram of the plant steroid receptor BRI1 (in blue) bound to brassinolide (in gray, in bonds representation) and to SERK1, shown in the same orientation (PDB-ID. 4lsx). (B) View of the inner surface of the SERK1 LRR domain (PDB-ID 4lsc, surface representation, in gray). A ribbon diagram of SERK1 in the same orientation is shown alongside. Residues interacting with the HAESA or BRI1 LRR domains are shown in orange or magenta, respectively. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.013 Comparison of our HAESA – IDA – SERK1 structure with the brassinosteroid receptor signaling complex, where SERK1 also acts as co-receptor, reveals an overall conserved mode of SERK1 binding, while the ligand binding pockets map to very different areas in the corresponding receptors (LRRs 2 – 14; HAESA; LRRs 21 – 25, BRI1) and may involve an island domain (BRI1) or not (HAESA) (Figure 6A). Several residues in the SERK1 N-terminal capping domain (Thr59SERK1, Phe61SERK1) and the LRR inner surface (Asp75SERK1, Tyr101SERK1, SER121SERK1, Phe145SERK1) contribute to the formation of both complexes (Figures 4C,D, 6B). In addition, residues 53-55SERK1 from the SERK1 N-terminal cap mediate specific interactions with the IDA peptide (Figures 4C, 6B). These residues are not involved in the sensing of the steroid hormone brassinolide. In both cases however, the co-receptor completes the hormone binding pocket. This fact together with the largely overlapping SERK1 binding surfaces in HAESA and BRI1 allows us to speculate that SERK1 may promote high-affinity peptide hormone and brassinosteroid sensing by simply slowing down dissociation of the ligand from its cognate receptor. Different plant peptide hormone families contain a C-terminal (Arg)-His-Asn motif, which in IDA represents the co-receptor recognition site. Structure-guided multiple sequence alignment of IDA and IDA-like peptides with other plant peptide hormone families, including CLAVATA3 – EMBRYO SURROUNDING REGION-RELATED (CLV3/CLE), ROOT GROWTH FACTOR – GOLVEN (RGF/GLV), PRECURSOR GENE PROPEP1 (PEP1) from Arabidopsis thaliana. The conserved (Arg)-His-Asn motif is highlighted in red, the central Hyp residue in IDLs and CLEs is marked in blue. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.014 Our experiments reveal that SERK1 recognizes a C-terminal Arg-His-Asn motif in IDA. Importantly, this motif can also be found in other peptide hormone families (Figure 7). Among these are the CLE peptides regulating stem cell maintenance in the shoot and the root. It is interesting to note, that CLEs in their mature form are also hydroxyprolinated dodecamers, which bind to a surface area in the BARELY ANY MERISTEM 1 receptor that would correspond to part of the IDA binding cleft in HAESA. Diverse plant peptide hormones may thus also bind their LRR-RK receptors in an extended conformation along the inner surface of the LRR domain and may also use small, shape-complementary co-receptors for high-affinity ligand binding and receptor activation. Materials and methods Protein expression and purification Synthetic genes coding for the Arabidopsis thaliana HAESA (residues 20–620) and SERK1 ectodomains (residues 24–213, carrying Asn115→Asp and Asn163→Gln mutations), codon optimized for expression in Trichoplusia ni (Geneart, Germany), were cloned into a modified pBAC-6 transfer vector (Novagen, Billerica, MA), providing an azurocidin signal peptide and a C-terminal TEV (tobacco etch virus protease) cleavable Strep-9xHis tandem affinity tag. Recombinant baculoviruses were generated by co-transfecting transfer vectors with linearised baculovirus DNA (ProFold-ER1, AB vector, San Diego, CA) followed by viral amplification in Spodoptera frugiperda Sf9 cells. The HAESA and SERK1 ectodomains were individually expressed in Trichoplusia ni Tnao38 cells using a multiplicity of infection of 3, and harvested from the medium 2 days post infection by tangential flow filtration using 30 kDa MWCO and 10 kDa MWCO (molecular weight cut-off) filter membranes (GE Healthcare Life Sciences, Pittsburgh, PA), respectively. Proteins were purified separately by sequential Ni2+ (HisTrap HP, GE Healthcare) and Strep (Strep-Tactin Superflow high-capacity, IBA, Germany) affinity chromatography. Next, affinity tags were removed by incubating the purified proteins with recombinant Strep-tagged TEV protease in 1:100 molar ratio. The cleaved tag and the protease were separated from HAESA and SERK1 by a second Strep affinity step. The purified HAESA ectodomain was incubated with a synthetic IDA peptide (YVPIPPSA-Hyp-SKRHN, the N-terminal Tyr residue was added to allow for peptide quantification by UV absorbance) and the SERK1 ectodomain in 1:1:1.5 molar ratio. The HAESA-IDA-SERK1 complex was purified by size exclusion chromatography on a Superdex 200 HR10/30 column (GE Healthcare) equilibrated in 20 mM citric acid pH 5.0, 100 mM NaCl). Peak fractions containing the complex were concentrated to ~10 mg/mL and immediately used for crystallization. About 0.2 mg of purified HAESA and 0.1 mg of purified SERK1 protein were obtained from 1 L of insect cell culture, respectively. Crystallization and data collection Hexagonal crystals of the isolated HAESA ectodomain developed at room-temperature in hanging drops composed of 1.0 μL of protein solution (5.5 mg/mL) and 1.0 μL of crystallization buffer (21% PEG 3,350, 0.2 M MgCl2 · 6 H2O, 0.1 M citric acid pH 4.0), suspended above 1.0 mL of crystallization buffer. For structure solution crystals were derivatized and cryo-protected by serial transfer into crystallization buffer supplemented with 0.5 M NaI and 15% ethylene glycol and cryo-cooled in liquid nitrogen. Redundant single-wavelength anomalous diffraction (SAD) data to 2.39 Å resolution were collected at beam-line PXII at the Swiss Light Source (SLS), Villigen, CH with λ=1.7 Å. A native data set to 1.74 Å resolution was collected on a crystal from the same drop cryo-protected by serial transfer into crystallization buffer supplemented with 15% (v/v) ethylene glycol only (λ=1.0 Å; Table 1). HAESA complexes with IDA (PIPPSA-Hyp-SKRHN), PKGV-IDA (YPKGVPIPPSA-Hyp-SKRHN) and IDL1 (LVPPSG-Hyp-SMRHN) peptide hormones were obtained by soaking apo crystals in crystallization buffer containing the respective synthetic peptide at a final concentration of 15 mM. Soaked crystals diffracted to 1.86 Å (HAESA – IDA), 1.94 Å (HAESA-PKGV-IDA) and 2.56 Å resolution (HAESA – IDL1), respectively (Table 2). Orthorhombic crystals of the HAESA-IDA-SERK1 complex developed in 18% PEG 8000, MgCl2 · 6 H2O, 0.1 M citric acid and diffracted to 2.43 Å resolution (Table 2). Data processing and scaling was done in XDS (version: Nov 2014). Structure solution and refinement The SAD method was used to determine the structure of the isolated HAESA ectodomain. SHELXD located 32 iodine sites (CC All/Weak 37.7/14.9). 20 consistent sites were input into the program SHARP for phasing and identification of 8 additional sites at 2.39 Å resolution. Refined heavy atom sites and phases were provided to PHENIX.AUTOBUILD for density modification and automated model building. The structure was completed in alternating cycles of model building in COOT and restrained TLS refinement in REFMAC5 (version 5.8.0107) against an isomorphous high resolution native data set. Crystals contain one HAESA monomer per asymmetric unit with a solvent content of ~55%, the final model comprises residues 20 – 615. The refined structure has excellent stereochemistry, with 93.8% of all residues in the favored region of the Ramachandran plot, no outliers and a PHENIX.MOLPROBITY score of 1.34 (Table 1). The HAESA – IDA – SERK1 complex structure was determined by molecular replacement with the program PHASER, using the isolated HAESA and SERK1 (PDB-ID: 4LSC) LRR domain structures as search models. The solution comprises one HASEA-IDA-SERK1 complex in the asymmetric unit. The structure was completed in iterative cycles of manual model-building in COOT and restrained TLS refinement in REFMAC5. Amino acids whose side-chain position could not be modeled with confidence were truncated to alanine (0.6 – 1% of total residues), the stereochemistry of N-linked glycan structures was assessed with the CCP4 program PRIVATEER-VALIDATE. The refined model has 94.44% of all residues in the favored region of the Ramachandran plot, no outliers and a PHENIX.MOLPROBITY score of 1.17 (Table 2). Structural visualization was done with POVScript+ and POV-Ray (http://www.povray.org). Size-exclusion chromatography Gel filtration experiments were performed using a Superdex 200 HR 10/30 column (GE Healthcare) pre-equilibrated in 20 mM citric acid (pH 5) and 100 mM NaCl. 100 μL of the isolated HAESA ectodomain (5.5 mg/mL), of the purified SERK1 LRR domain (3 mg/mL) or of mixtures of HAESA and SERK1 (either in the presence or absence of synthetic wild-type IDA, wild-type IDL1 or mutant IDA peptides at a concentration of 25 μM; 10 mg/mL; samples contained HAESA and SERK1 in 1:1 molar ratio) were loaded sequentially onto the column and elution at 0.5 mL/min was monitored by ultraviolet absorbance at 280 nm. Isothermal titration calorimetry ITC experiments were performed using a Nano ITC (TA Instruments, New Castle, DE) with a 1.0 mL standard cell and a 250 μL titration syringe. Proteins were dialyzed extensively against ITC buffer (20 mM citric acid pH 5.0, 100 mM NaCl) and synthetic wild-type or point-mutant peptides (with wild-type IDA sequence YVPIPPSA-Hyp-SKRHN, PKGV-IDA YPKGVPIPPSA-Hyp-SKRHN, IDA-SFVN YPIPPSA-Hyp-SKRHNSFVN, IDL1 YLVPPSG-Hyp-SMRHN and CLV3 sequence YRTV-Hyp-SG-Hyp-DPLHH) were dissolved in ITC buffer prior to all titrations. Molar protein concentrations for SERK1 and HAESA were calculated using their molar extinction coefficient and a molecular weight of 27,551 and 74,896 Da, respectively (determined by MALDI-TOF mass spectrometry). Experiments were performed at 25°C. A typical experiment consisted of injecting 10 μL aliquots of peptide solution (250 μM) into 20 μM HAESA. The concentrations for the complex titrations were 150 μM of ligand (either wild-type or point-mutant IDA peptides) in the syringe and 10 μM of a 1:1 HAESA – SERK1 protein mixture in the cell at time intervals of 150 s to ensure that the titration peak returned to the baseline. Binding of SERK1 to HAESA was assessed by titrating SERK1 (100 μM) into a solution containing HAESA (10 μM) in the pre- or absence of 150 μM wild-type IDA peptide. ITC data were corrected for the heat of dilution by subtracting the mixing enthalpies for titrant solution injections into protein free ITC buffer. Data were analyzed using the NanoAnalyze program (version 2.3.6) as provided by the manufacturer. In vitro kinase trans-phosphorylation assay Coding sequences of SERK1 kinase domain (SERK1-KD) (residues 264–625) and HAESA-KD (residues 671–969) were cloned into a modified pET (Novagen) vector providing an TEV-cleavable N-terminal 8xHis-StrepII-Thioredoxin tag. Point mutations were introduced into the SERK1 (Asp447→Asn; mSERK1) and HAESA (Asp837→Asn; mHAESA) coding sequences by site directed mutagenesis, thereby rendering the kinases inactive. The plasmids were transformed into E.coli Rosetta 2 (DE3) (Novagen). Protein expression was induced by adding IPTG to final concentration of 0.5 mM to cell cultures grown to an OD600 = 0.6. Cells were then incubated at 16°C for 18 hr, pelleted by centrifugation at 5000 x g and 4°C for 15 min, and resuspended in buffer A (20 mM Tris-HCl pH 8, 500 mM NaCl, 4 mM MgCl2 and 2 mM β-Mercaptoethanol) supplemented with 15 mM Imidazole and 0.1% (v/v) Igepal. After cell lysis by sonication, cell debris was removed by centrifugation at 35,000 x g and 4°C for 30 min. The recombinant proteins were isolated by Co2+ metal affinity purification using a combination of batch and gravity flow approaches (HIS-Select Cobalt Affinity Gel, Sigma, St. Louis, MO). After washing the resin with the wash buffer (buffer A + 15 mM Imidazole) proteins were eluted in buffer A supplemented with 250 mM Imidazole. All elutions were then dialyzed against 20 mM Tris-HCl pH 8, 250 mM NaCl, 4 mM MgCl2 and 0.5 mM TCEP. For SERK1-KD and mSERK1-KD the 8xHis-StrepII-Thioredoxin tag was removed with 6xHis tagged TEV protease. TEV and the cleaved tag were removed by a second metal affinity purification step. Subsequently, all proteins were purified by gel filtration on a Superdex 200 10/300 GL column equilibrated in 20 mM Tris pH 8, 250 mM NaCl, 4 mM MgCl2 and 0.5 mM TCEP. Peak fractions were collected and concentrated using Amicon Ultra centrifugation devices (10,000 MWCO). For in vitro kinase assays, 1 μg of HAESA-KD, 0.25 μg of SERK1-KD and 2 μg of mSERK1 and mHAESA were used in a final reaction volume of 20 μl. The reaction buffer consisted of 20 mM Tris pH 8, 250 mM NaCl, 4 mM MgCl2 and 0.5 mM TCEP. The reactions were started by the addition of 4 μCi [γ-32P]-ATP (Perkin-Elmer, Waltham, MA), incubated at room temperature for 45 min and stopped by the addition of 6x SDS-loading dye immediately followed by incubating the samples at 95°C. Proteins of the whole reaction were subsequently separated via SDS-PAGE in 4–15% gradient gels (TGX, Biorad, Hercules, CA) and stained with Instant Blue (Expedeon, San Diego, CA). After pictures were taken of the stained gel, 32P-derived signals were visualized by exposing the gel to an X-ray film (Fuji, SuperRX, Valhalla, NY). Plant material and generation of transgenic lines 35S::IDA wild-type and 35S::IDA (R66 → Ala/K67 → Ala) over-expressing transgenic lines in Col-0 background were generated as follows: The constructs were introduced in the destination vector pB7m34GW2 and transferred to A. tumefaciens strain pGV2260. Plants were transformed using the floral dip method. Transformants were selected in medium supplemented with BASTA up to the T3 generation. For phenotyping, plants were grown at 21°C with 50% humidity and a 16h light: 8 hr dark cycle. RNA analyses Plants were grown on ½ Murashige and Skoog (MS) plates supplemented with 1% sucrose. After 7 d, ∼30 to 40 seedlings were collected and frozen in liquid nitrogen. Total RNA was extracted using a RNeasy plant mini kit (Qiagen, Valencia, CA), and 1 μg of the RNA solution obtained was reverse-transcribed using the SuperScritpVILO cDNA synthesis kit (Invitrogen, Grand Island, NY). RT-qPCR amplifications and measurements were performed using a 7900HT Fast Real Time PCR-System by Applied Biosystems (Carlsbad, CA). RT-qPCR amplifications were monitored using SYBR-Green fluorescent stain (Applied Biosystems). Relative quantification of gene expression data was performed using the 2−ΔΔCT (or comparative CT) method. Expression levels were normalized using the CT values obtained for the actin2 gene (forward: TGCCAATCTACGAGGGTTTC; reverse: TTCTCGATGGAAGAGCTGGT). For detection and amplification of IDA sequence we used specific primers (forward: TCGTACGATGATGGTTCTGC; reverse: GAATGGGAACGCCTTTAGGT). The presence of a single PCR product was further verified by dissociation analysis in all amplifications. All quantifications were made in quadruplicates on RNA samples obtained from three independent experiments. Petal break measurements serk1-1, serk2-2, serk3-1, serk4-1 and serk5-1 and Col-0 wild-type plants were grown in growth chambers at 22°C under long days (16 hr day/8 hr dark) at a light intensity of 100 µE·m-2·sec-1. Petal break-strength was quantified as the force in gram equivalents required for removal of a petal from a flower when the plants had a minimum of twenty flowers and siliques. Measurements were performed using a load transducer as described in. Break-strength was measured for 15 plants and a minimum of 15 measurements at each position. Funding Information This paper was supported by the following grants:   to Michael Hothorn.   to Michael Hothorn.   to Michael Hothorn.   to Melinka A Butenko.   to Benjamin Brandt.   to Julia Santiago. Additional information Competing interests The authors declare that no competing interests exist. Author contributions JS, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article. BB, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article. MW, Acquisition of data, Analysis and interpretation of data. UH, Acquisition of data, Analysis and interpretation of data. LAH, Analysis and interpretation of data, Drafting or revising the article. MAB, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article. MH, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article. References Ida: A peptide ligand regulating cell separation processes in Arabidopsis The Arabidopsis thaliana SOMATIC EMBRYOGENESIS RECEPTOR-LIKE KINASES1 and 2 control male sporogenesis Arabidopsis SOMATIC EMBRYOGENESIS RECEPTOR KINASE proteins serve brassinosteroid-dependent and -independent signaling pathways Crystal structures of the phosphorylated BRI1 kinase domain and implications for brassinosteroid signal initiation SERK co-receptor kinases Generation, representation and flow of phase information in structure determination: Recent developments in and around SHARP 2.0 Inflorescence deficient in abscission controls floral organ abscission in Arabidopsis and identifies a novel family of putative ligands in plants Plant peptides in signalling: Looking for new partners Tools and strategies to match peptide-ligand receptor pairs Regulation of floral organ abscission in Arabidopsis thaliana CLAVATA3 is a specific regulator of shoot and floral meristem development affecting the same processes as CLAVATA1 Floral dip: A simplified method for agrobacterium-mediated transformation of Arabidopsis thaliana Arabidopsis SOMATIC EMBRYOGENESIS RECEPTOR KINASES1 and 2 are essential for tapetum development and microspore maturation Molprobity: All-atom contacts and structure validation for proteins and nucleic acids Coot : Model-building tools for molecular graphics Povscript+ : A program for model and data visualization using persistence of vision ray-tracing Genetic evidence for an indispensable role of somatic embryogenesis receptor kinases in brassinosteroid signaling Ao38, a new cell line from eggs of the black witch moth, Ascalapha odorata (Lepidoptera: Noctuidae), is permissive for AcMNPV infection and produces high levels of recombinant proteins Multiple contrast tests in the presence of heteroscedasticity The Arabidopsis SOMATIC EMBRYOGENESIS RECEPTOR KINASE 1 gene is expressed in developing ovules and embryos and enhances embryogenic competence in culture Simultaneous inference in general parametric models Structural basis of steroid hormone perception by the receptor kinase BRI1 HAESA, an Arabidopsis leucine-rich repeat receptor kinase, controls floral organ abscission Dictionary of protein secondary structure: Pattern recognition of hydrogen-bonded and geometrical features Automatic processing of rotation diffraction data from crystals of initially unknown symmetry and cell constants A plant peptide encoded by CLV3 identified by in situ MALDI-TOF MS analysis A petal breakstrength meter for Arabidopsis abscission studies The SERK1 receptor-like kinase regulates organ separation in Arabidopsis flowers Analysis of relative gene expression data using real-time quantitative PCR and the 2(-delta delta C(T)) method Secreted peptide signals required for maintenance of root stem cell niche in Arabidopsis Phaser crystallographic software Differential function of Arabidopsis SERK family receptor-like kinases in stomatal patterning Ligand-induced receptor-like kinase complex regulates floral organ abscission in Arabidopsis Refinement of macromolecular structures by the maximum-likelihood method Brassinolide-2,3-acetonide: A brassinolide-induced rice lamina joint inclination antagonist Letting go is never easy: Abscission and receptor-like protein kinases Transcriptional profiling of the Arabidopsis abscission mutant hae hsl2 by RNA-Seq Arabidopsis CLV3 peptide directly binds CLV1 ectodomain A glycopeptide regulating stem cell fate in Arabidopsis thaliana  AtSERK1 expression precedes and coincides with early somatic embryogenesis in Arabidopsis thaliana Reduced surface: An efficient way to compute molecular surfaces Molecular mechanism for plant steroid receptor activation by somatic embryogenesis co-receptor kinases Role of threonines in the Arabidopsis thaliana somatic embryogenesis receptor kinase 1 activation loop in phosphorylation A short history of SHELX Biochemical mapping of a ligand-binding domain within Arabidopsis BAM1 reveals diversified ligand recognition mechanisms of plant LRR-RKs Overexpression of INFLORESCENCE DEFICIENT IN ABSCISSION activates cell separation in vestigial abscission zones in Arabidopsis The EPIP peptide of INFLORESCENCE DEFICIENT IN ABSCISSION is sufficient to induce abscission in Arabidopsis through the receptor-like kinases HAESA and HAESA-LIKE2 Structure reveals that BAK1 as a co-receptor recognizes the BRI1-bound brassinolide Structural basis for recognition of an endogenous peptide by the plant receptor kinase PEPR1 Analysis of phosphorylation of the receptor-like protein kinase HAESA during arabidopsis floral abscission Iterative model building, structure refinement and density modification with the PHENIX autobuild wizard Allosteric receptor activation by the plant peptide hormone phytosulfokine 10.7554/eLife.15075.017 Decision letter Zhang Mingjie In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included. Thank you for submitting your article \"Mechanistic insight into a peptide hormone signaling complex mediating floral organ abscission\" to eLife for consideration by eLife. Your article has been favorably evaluated by John Kuyiyan (Senior editor) and three reviewers, one of whom, (Mingjie Zhang) is a member of our Board of Reviewing Editors. The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. As you will see that the required revisions are essentially clarifications and some additional analysis of existing data in nature. Summary: In this work, Hothorn and colleagues investigated the structural basis governing the recognition of peptide hormone IDA during plant floral abscission process. Through an array of complex structures, supplemented with biochemical and genetic experiments, the authors uncovered the IDA recognition mechanism by a co-receptor (HAESA and SERK1) detection mechanism. The structures also reveal the specific recognition mechanism of the 12-residue IDA core peptide sequence by the co-receptors, and suggest that this 12-residue IDA sequence is likely to be the mature peptide hormone functioning in plants. The comparison of the structures of the HAESA/SERK1/IDA complex and the previously determined BRA1/SERK1/brassinolide complex by the same group also suggests a co-receptor pairing mechanism for various plant hormones. The story gives detailed and novel mechanistic insights in the perception of IDA during floral abscission, and is convincing and worthy to be considered for publication in eLife with the following revisions. Key issues which need to be addressed: 1) As the results reported here rest largely on the interpretation of the structural data, the following points need to be addressed by the authors. i) Temperature factors (Wilson B and residual) are unusually high for the reported resolution. Are there portions of the structure that exhibit more disorder or are these high temperature factors throughout the structure? Are there potential problems with radiation damage due to the high multiplicity? ii) A simulated annealing omit map figure should be provided as the peptides all exhibit very high temperature factors. iii) How many side chains were trimmed due to poor electron density? If this is a significant percentage, it should be noted. 2) A further discussion comparing the mechanisms of BRI1-BL-SERK1 and HAESA-IDA-SERK1 would be helpful as a structural comparison is presented. As BR1 binds its ligand with high affinity, the low affinity of HAESA for IDA could be further discussed. 3) Are the results here applicable to the related receptor HSL2? Are the residues that interact with SERK1 and IDA conserved in HSL2? 4) The authors show that N-terminal extension of the peptide does not impact on binding efficiency, but what would happen if the peptide was extended at the C-terminal end, at the suggested cleavage site? Would cleavage be required for recognition? A brief discussion on this point may help. 5) Figure 3C: Could the authors comment on the difference between the blue (SERK1 vs. HAESA-IDA) and the black (IDA vs. HAESA-SERK1) line? SERK1 vs. HAESA-IDA gives a Kd of 75 nM, yet IDA vs. SERK1-HAESA gives a Kd of 350 nM. In the text the authors keep referring to the 75 nM Kd, but not the 350 nM Kd. Is it really fair to say the Kd is 75 nM? 6) In the Discussion, can the authors comment further on the discrepancy between their study and the study of Lewis et al. (Plant J, 2010) concerning the role of SERK1 in floral abscission? Similarly, could the authors comment on the fact that the LRR-RLP EVD/SOBIR seems to be a negative regulator of the HAESA/HSL2 pathway (Leslie et al., Development, 2010), which seems puzzling given that EVD/SOBIR function is normally restricted to LRR-RLPs (Gust \u0026 Felix, Curr Op Plant Biol, 2014). 7) Given that the central hydroxyproline in IDA is of such crucial importance for binding, isn't it surprising that IDAΔ69N does not bind to HAESA at all? Wouldn't it be expected that the remaining part of the peptide still binds to HAESA? 8) Figure 1—figure supplement 2: How is it possible that charged amino acids are involved in hydrophobic interactions? 9) Are the distances shown in the graphical representation of the structures proportional? It would seem that some of the aromatic rings could cause steric hindrance. 10) Why did the authors decide to express HAESA and SERK1 without signal peptide? Would it make a difference for binding of IDA, if they leave the SP on? 11) Figure 7: Are the homologous regions also the active parts of these peptides? And could the authors display amino acids numbers on either side of the fragments? 12) Have the authors ever measured dissociation of the peptide from the complex? And in this regard, to what does \"highly stable receptor – co-receptor complex\" refer/compare to? 13) Figures 3A and 5C require statistical analyses. 10.7554/eLife.15075.018 Author response  1) As the results reported here rest largely on the interpretation of the structural data, the following points need to be addressed by the authors. i) Temperature factors (Wilson B and residual) are unusually high for the reported resolution.  We thank the reviewer(s) for pointing out this issue: Indeed our mean B-values deviate substantially from the expected mean B-values (resolution 1.75 – 2.00 A, B(Wilson) ~ 27.0 over 5,510 structures in the PDB; Pavel Afonine, personal communication). We would like to note that due to the many N-glycosylation sites dispersed over the HAESA LRR domain (shown in Figure 1—figure supplement 1D), we find relatively few crystal contacts in our P3121 crystal form, which may rationalize our high B-values. We have reanalyzed our space group assignment (using the CCP4 program ZANUDA) and checked for any signs of problems during data collection (ice rings, multiple crystal lattices, splitting, using the programs XDS and XDSSTAT), as well as for twinning and pseudosymmetry (using phenix.xtriage). No such problems appear to exist, our structures refine very well and our refined B-values are in good agreement with our Wilson B-factors (see Table 2). Thus, the high B-values appear to represent an intrinsic property of our crystals and are not the result of a poor data collection strategy or inappropriate crystallographic analysis. Are there portions of the structure that exhibit more disorder or are these high temperature factors throughout the structure?  Cα trace of the HAESA LRR domain and IDA peptide colored according to B-factor from low (60.9, in blue) to high (134.7, in red). Mean B-value is 79.5. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.015 Yes. As shown in revised Figure 4B, the Cterminal LRRs of HAESA in contact with SERK1 in our complex structure appear to be somewhat flexible. Author response image 1. illustrates that the B-values are significantly higher in the C-terminal part of the HAESA LRR domain (with the Cterminal capping domain being the most flexible), while both the N-terminal LRRs of HAESA (with exception of the LRR N-terminal capping domain) and the IDA peptide appear better ordered in our P3121 crystals form. Are there potential problems with radiation damage due to the high multiplicity?  No. Data were collected at SLS beamline PXIII equipped with a Dectris Pilatus 2M-F detector. We perform all our data collections at very low dose and high multiplicity of measurement, which at this beam-line produces similar results compared to exposing the crystal at higher dose for a smaller angular range. We collected 360 deg with 0.1 deg slices and obtained a Wilson B-value of 80, with no sign of radiation damage in our data processing (subroutine COLSPOT in XDS over all frames). To test the reviewer's hypothesis we cut the data after 90 deg (when completeness approaches 100%) and we obtained a Wilson B-value of 78 and a refined mean B-value of around 75. These value do not significantly differ from our presented 360 deg data set and thus it is unlikely that radiation damage produces these high B-values. Again, they rather appear to be an intrinsic property of our crystals. ii) A simulated annealing omit map figure should be provided as the peptides all exhibit very high temperature factors. Thank you for this suggestion. In our first submission, we presented 2Fo-Fc omit electron density maps for our HAESA-IDA/IDL complex structures. As suggested, we now present simulated annealing omit maps in revised Figure 2A, B, C. The maps were generated like this: phenix.composite_omit_map *.pdb *.mtz *.cif nproc=8 anneal=True We would like to note that our peptide are well ordered in our structures, and their B-values match the B-values of their interacting LRR surface (compare Author response image 1). iii) How many side chains were trimmed due to poor electron density? If this is a significant percentage, it should be noted.  Here are the requested numbers (trimmed residues out of total residues in asymmetric unit, percentage): HAESA apo: 7 out of 595 (1%) HAESA IDA: 6 out of 597 + 12 (1%) HAESA IDL1: 6 out of 597 + 12 (1%) HAESA – IDA – SERK1: 5 out of 594 + 12 + 185 (0.6%) We have included a statement in the Methods section that reads: “Amino acids whose side-chain position could not be modeled with confidence were truncated to alanine (0.6 – 1% of total residues)[…]”  2) A further discussion comparing the mechanisms of BRI1-BL-SERK1 and HAESA-IDA-SERK1 would be helpful as a structural comparison is presented.  We have expanded our discussion of the HAESA – SERK1 and BRI1 – SERK1 interfaces. We now specify the SERK1 residues in contact with both receptors and the SERK1 residues unique to HAESA/IDA sensing. We also comment on the very different ligand binding modes in HAESA and BRI1 and specify that different LRR segments contribute to the formation of the respective steroid and peptide hormone binding pockets. We feel however that an in-depth comparison of the interacting surfaces is beyond the scope of this report and partially redundant with our earlier work (Santiago et al., Science, 2013). In our opinion, such an analysis seems more appropriate for a review on the subject, which we are currently preparing. Our revised Discussion now reads: “Comparison of our HAESA – IDA – SERK1 structure with the brassinosteroid receptor signaling complex, where SERK1 also acts as co-receptor (Santiago, Henzler, and Hothorn 2013), reveals an overall conserved mode of SERK1 binding, while the ligand binding pockets map to very different areas in the corresponding receptors (LRRs 2 – 14; HAESA; LRRs 21 – 25, BRI1) and may involve an island domain (BRI1) or not (HAESA) (Figure 6A).[…] These residues are not involved in the sensing of the steroid hormone brassinolide (Santiago, Henzler, and Hothorn 2013). In both cases however, the co-receptor completes the hormone binding pocket.” As BR1 binds its ligand with high affinity, the low affinity of HAESA for IDA could be further discussed.  High affinity brassinosteroid binding to BRI1 was previously shown using BRI1-enriched plant extracts and radiolabeled brassinolide (Wang et al., Nature410:380-383, 2001). We now know that co-immunoprecipitations of BRI1 from Arabidopsis contain SERK proteins (compare for example Jaillais et al., PNAS, 2011) and thus the reported binding constants likely correspond to steroid binding to BRI1-SERK complexes, not to BRI1 alone. We would thus prefer not to compare the binding affinities for brassinosteroid and peptide hormone ligands at this point. 3) Are the results here applicable to the related receptor HSL2? Are the residues that interact with SERK1 and IDA conserved in HSL2?  Yes. We present a structure-based sequence alignment of AtHAESA and AtHSL2, as well as other HAESA-type receptors from different plant species in Figure 1—figure supplement 3. In the peptide binding surface, 17 out of 26 contributing amino-acids are conserved among AtHAESA and AtHSL2. 13 out of 19 interacting residues in the HAESA – SERK1 complex are also present in AtHSL2. We feel that this is strong conservation given that the AtHAESA and AtHSL2 ectodomains share 45% overall sequence identity. We have included a statement in our manuscript that reads: “Indeed, we find many of the residues contributing to the formation of the IDA binding surface in HAESA to be conserved in HSL2 and other HAESA-type receptors in different plant species (Figure 1—figure supplement 3).” 4) The authors show that N-terminal extension of the peptide does not impact on binding efficiency, but what would happen if the peptide was extended at the C-terminal end, at the suggested cleavage site?  Isothermal titration calorimetry thermograph of the C-terminally extended IDA-SFVN peptide (200 μM) titrated into a solution containing 20 μM of the purified HAESA ectodomain. No detectable binding is observed. DOI:\nhttp://dx.doi.org/10.7554/eLife.15075.016 Thank you for suggesting this experiment. We synthesized a C-terminally extended version of the IDA peptide (IDA-SFVN with sequence YPIPPSA-Hyp- SKRHN SFVN) and performed quantitative binding assays by ITC. As shown in Author response image 2, we cannot observe any detectable binding of this C-terminally extended peptide to the HAESA ectodomain, consistent with our crystallographic models that suggest that HAESA specifically senses an active IDA 12mer. We have incorporated this new result in Figure 2D. We have included a new statement in the manuscript that reads: “The COO- group of Asn69IDA is in direct contact with Arg407HAESA and Arg409HAESA and HAESA cannot bind a C-terminally extended IDA-SFVN peptide (Figures 1D, F, 2D).” Would cleavage be required for recognition? A brief discussion on this point may help.  Yes. We have modified our manuscript accordingly: “This suggests that the conserved Asn69 may constitute the very C-terminus of the mature IDA peptide in planta and that active IDA is generated by proteolytic processing from a longer pre-protein (Stenvik et al. 2008).“ 5) Figure 3C: Could the authors comment on the difference between the blue (SERK1 vs. HAESA-IDA) and the black (IDA vs. HAESA-SERK1) line? SERK1 vs. HAESA-IDA gives a Kd of 75 nM, yet IDA vs. SERK1-HAESA gives a Kd of 350 nM. In the text the authors keep referring to the 75 nM Kd, but not the 350 nM Kd. Is it really fair to say the Kd is 75 nM? Yes and No. These are two different experiments. We first measured (by ITC) the binding affinity of IDA (in the syringe) binding to a protein solution containing equimolar ratios of HAESA and SERK1 (Kdis 350 nM in this case). Next, we titrated a concentrated SERK1 solution (in the syringe) into a solution of HAESA containing IDA in 10fold molar excess (Kdin this case is 75 nM). Given that the experimental conditions (protein and peptide concentrations and molar ratios between the components) are very different, we feel that the Kd 's obtained by these experiments are in good agreement (4.5 fold difference vs. a 60-260 fold difference when compared to the isolated HAESA ectodomain). Nevertheless, we addressed the reviewer's concern by modifying our manuscript which now states: “In this case, there was no detectable interaction between receptor and co-receptor, while in the presence of IDA, SERK1 strongly binds HAESA with a dissociation constant in the mid-nanomolar range. (Figure 3C).” 6) In the Discussion, can the authors comment further on the discrepancy between their study and the study of Lewis et al. (Plant J, 2010) concerning the role of SERK1 in floral abscission?  The process of floral organ abscission in Arabidopsis is divided into distinct steps where a gradual loosening of the cell wall between abscising cells can be measured as a reduction in petal breakstrength (Bleecker and Patterson, 1997). During floral abscission in wild-type plants a significant drop in breakstrength occurs shortly before the petals drop (shown in Figure 3A in our manuscript). Previously reported negative regulators of abscission, such as the transcription factor KNAT1, have an earlier reduction in breakstrength, indicative of early cell wall remodeling (Shi et al. 2011). Our results show that serk1 mutant plants, contrary to knat1 mutants and wild type, have a delay in cell wall loosening and organ separation (Figure 3A) thus positively regulating organ separation during abscission. The weaker phenotype when compared to haesa/hsl2 mutants is likely due to the redundant nature of other SERKs inthe abscission zone (recent work of Meng et al. 2016, cited in the Results and Discussion sections of our manuscript). It has previously been reported that mutations in SERK1 can rescue the block in abscission in plants without the functional ADP-ribosylation factor GTPase-activating protein NEVERSHED (NEV) (Lewis et al. 2010). However, as a mutation in SERK1 is not capable of rescuing the ida mutant phenotype (Lewis et al. 2010) and revertant mutants capable of rescuing the abscission defect of ida do not complement nev, it has been suggested that NEV and IDA function in parallel pathways to promote cell separation (Liu 2013). Our work does not rule out a function for SERK1 in such a parallel pathway, we merely report SERK1 can ALSO act as a positive regulator of abscission by interacting with HAESA in an IDA-dependent manner. We do not observe negative regulation of floral abscission using our SERK1 mutant alleles. Based on the available evidence there is thus little to discuss and speculate about the different functions of SERK1 in abscission, as no molecular mechanism for the negative role of SERK1 in this pathway has been reported thus far. We feel that it is beyond the scope of our manuscript to clarify the different roles of SERK1 in the Arabidopsis abscission zone. Similarly, could the authors comment on the fact that the LRR-RLP EVD/SOBIR seems to be a negative regulator of the HAESA/HSL2 pathway (Leslie et al., Development, 2010), which seems puzzling given that EVD/SOBIR function is normally restricted to LRR-RLPs (Gust \u0026 Felix, Curr Op Plant Biol, 2014). We did attempt to express and purify the EVR/SOBIR extracellular domain, but in our hands the protein is not properly secreted and hence unfolded. We thus could not further investigate the potential mechanism of EVR/SOBIR in the HAESA pathway. 7) Given that the central hydroxyproline in IDA is of such crucial importance for binding, isn't it surprising that IDAΔ69N does not bind to HAESA at all? Wouldn't it be expected that the remaining part of the peptide still binds to HAESA?  We thank the reviewers for pointing this out to us. Indeed, we find several structural and sequence features in IDA peptide to be important determinants for HAESA binding, namely the correct size of the peptide, the presence of a central Hyp residue and an intact C-terminal Arg-His-Asn motif that is buried in the structure. In the revised manuscript we now provide new experiments (binding of a C-terminal extended IDA peptide to HAESA) that clarifies this point (summarized in revised Figure 2D). We have revised our statement in the Discussion accordingly: “The central Hyp residue in IDA is found buried in the HAESA peptide binding surfaceand thus this post-translational modification may regulateIDA bioactivity.” 8) Figure 1—figure supplement 2: How is it possible that charged amino acids are involved in hydrophobic interactions?  We apologize for this confusing statement. It now reads: “A N-terminal Pro-rich motif in IDA makes contacts LRRs 2-6 of the receptor(Figure 1D, Figure 1—figure supplement 2A-C).” 9) Are the distances shown in the graphical representation of the structures proportional? It would seem that some of the aromatic rings could cause steric hindrance.  No. The graphical representation are proportional and e.g. Trp218 in the back of the binding pocket is not producing steric clashes with the peptide with the closest distance being 4.5 A. 10) Why did the authors decide to express HAESA and SERK1 without signal peptide? Would it make a difference for binding of IDA, if they leave the SP on?  No. We did express both the HAESA and SERK1 ectodomains fused to the signal peptide of human azurocidin, which provides very efficient secretion of LRR proteins in insect cells (Olczak \u0026 Olczak, Anal. Biochem., 2006) (see Methods, subsection “Protein Expression and Purification“). Both the native signal peptides for SERK1 and HAESA as well as the azurocidin signal peptide are being recognized and cleaved by the Trichoplusia ni signal peptidase. This results, just like in planta, in a mature receptor/coreceptor ectodomain starting with the first α-helix of the N-terminal capping domain (residues 20 and 24, respectively). Thus, there is no reason to believe that the signal peptide would play a role in IDA sensing. Using our system, we cannot produce HAESA and/or SERK1 ectodomains with an intact signal peptide, as this would impair folding and proper secretion of the recombinant proteins. 11) Figure 7: Are the homologous regions also the active parts of these peptides?  Yes. We have included three additional references in the Discussion section of our manuscript, which report the bioactive regions of CLV3/CLE, RGF and PEP peptides shown in Figure 7. The revised section now reads: “Importantly, this motif can also be found in other peptide hormone families (Kondo et al. 2006; Matsuzaki et al. 2010; Tang et al. 2015)(Figure 7). Among these are the CLE peptides regulating stem cell maintenance in the shoot and the root (Clark, Running, and Meyerowitz 1995). It is interesting to note, that CLEs in their mature form are also hydroxyprolinated dodecamers, which bind to a surface area in the BARELY ANY MERISTEM 1 receptor that would correspond to part of the IDA binding cleft in HAESA (Kondo et al. 2006;Ogawa et al. 2008; Shinohara et al. 2012).” And could the authors display amino acids numbers on either side of the fragments?  Yes. We have now included the residues number of each peptide in Figure 7. 12) Have the authors ever measured dissociation of the peptide from the complex?  No. We have not performed any biochemical experiment that would allow us to quantify the dissociation of the peptide from the ternary complex. In qualitative terms it is however of note that HAESA-IDA-SERK1 complexes do not dissociate in size exclusion chromatography experiments, even when the peptide is not provided in excess or supplied in the running buffer. And in this regard, to what does \"highly stable receptor – co-receptor complex\" refer/compare to?  The reviewers are correct, we should not claim that the complex is 'highly stable' if we have not quantified the dissociation rate. The revised sentence reads: “This ligand-induced formation of a receptor – co-receptor complex may allow the HAESA and SERK1 kinase domains to efficiently trans-phosphorylate and activate each other in the cytoplasm.” 13) Figures 3A and 5C require statistical analyses. Thank you for pointing this out to us. The statistical analysis of the petal break-strength assays shown in Figures 3A and 5C has been carried out by Prof. Ludwig A. Hothorn, Institute for Biostatistics, University of Hannover, Germany, whom we have added as an author on our manuscript: Statistical analysis for Figure 3A: The statistical analysis is described in the figure legend of Figure 3A; statistical significant changes are indicated by a * in the Figure itself. The revised figure legend reads: “Petal break-strength assays measure the force (expressed in gram equivalents) required to remove the petals from the flower of serk mutant plants compared to haesa/hsl2 mutant and Col-0 wild-type flowers. […] Petal break was found significantly increased in almost all positions (indicated with a *) for haesa/hsl2 and serk1-1 mutant plants with respect to the Col-0 control. Calculations were performed in R (R Core Team 2014) (version 3.2.3).” The two new references have been added to the Reference section of the manuscript. We have changed our Results section accordingly: “Our experiments suggest that among the SERK family members, SERK1 is a positive regulator offloral abscission. We found that the force required to remove the petals of serk1-1 mutants is significantlyhigher than that needed for wild-type plants, as previously observed for haesa/hsl2 mutants (Stenvik et al. 2008), and that floral abscission is delayed in serk1-1 (Figure 3A). The serk2-2, serk3-1, serk4-1 and serk5-1 mutant lines (Albrecht et al. 2008) showed a petal break-strength profile not significantly differentfrom wild-type plants.” Statistical analysis for Figure 5C: The statistical analysis is described in the figure legend of Figure 5C; statistical significant changes are indicated by * and # symbols in the Figure itself. The revised figure legend reads:”Quantitative petal break assay for Col-0 wild-type flowers and 35S::IDA wild-type and 35S::IDA K66A/R67A mutant flowers. […] Up to inflorescence position 4, petal break in 35S::IDA K66A/R67A mutant plants was significantly increased compared to both Col-0 control plants (b) and 35S::IDA plants (c).” We have changed our Results section accordingly: “We overexpressed full-length wild-type IDA or this Lys66IDA/Arg67IDA → Ala double-mutant to similar levels in Col-0 Arabidopsis plants (Figure 5D). […] Comparison of 35S::IDA wild-type and mutant plants further indicates that mutation of Lys66IDA/Arg67IDA→ Ala may cause a weak dominant negative effect (Figure 5C-E).” Following the suggestions from for example Nuzzo (Nature506:150-152, 2014) and Trafirmow and Marks (Basic and Applied Social Psychology37:1-2, 2015), we decided not to report 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+[{"sourceid":"4848761","sourcedb":"","project":"","target":"","text":"Predictive features of ligand‐specific signaling through the estrogen receptor Abstract Some estrogen receptor‐α (ERα)‐targeted breast cancer therapies such as tamoxifen have tissue‐selective or cell‐specific activities, while others have similar activities in different cell types. To identify biophysical determinants of cell‐specific signaling and breast cancer cell proliferation, we synthesized 241 ERα ligands based on 19 chemical scaffolds, and compared ligand response using quantitative bioassays for canonical ERα activities and X‐ray crystallography. Ligands that regulate the dynamics and stability of the coactivator‐binding site in the C‐terminal ligand‐binding domain, called activation function‐2 (AF‐2), showed similar activity profiles in different cell types. Such ligands induced breast cancer cell proliferation in a manner that was predicted by the canonical recruitment of the coactivators NCOA1/2/3 and induction of the GREB1 proliferative gene. For some ligand series, a single inter‐atomic distance in the ligand‐binding domain predicted their proliferative effects. In contrast, the N‐terminal coactivator‐binding site, activation function‐1 (AF‐1), determined cell‐specific signaling induced by ligands that used alternate mechanisms to control cell proliferation. Thus, incorporating systems structural analyses with quantitative chemical biology reveals how ligands can achieve distinct allosteric signaling outcomes through ERα. Introduction Many drugs are small‐molecule ligands of allosteric signaling proteins, including G protein‐coupled receptors (GPCRs) and nuclear receptors such as ERα. These receptors regulate distinct phenotypic outcomes (i.e., observable characteristics of cells and tissues, such as cell proliferation or the inflammatory response) in a ligand‐dependent manner. Small‐molecule ligands control receptor activity by modulating recruitment of effector enzymes to distal regions of the receptor, relative to the ligand‐binding site. Some of these ligands achieve selectivity for a subset of tissue‐ or pathway‐specific signaling outcomes, which is called selective modulation, functional selectivity, or biased signaling, through structural mechanisms that are poorly understood (Frolik et al, 1996; Nettles \u0026 Greene, 2005; Overington et al, 2006; Katritch et al, 2012; Wisler et al, 2014). For example, selective estrogen receptor modulators (SERMs) such as tamoxifen (Nolvadex®; AstraZeneca) or raloxifene (Evista®; Eli Lilly) (Fig 1A) block the ERα‐mediated proliferative effects of the native estrogen, 17β‐estradiol (E2), on breast cancer cells, but promote beneficial estrogenic effects on bone mineral density and adverse estrogenic effects such as uterine proliferation, fatty liver, or stroke (Frolik et al, 1996; Fisher et al, 1998; McDonnell et al, 2002; Jordan, 2003). Allosteric control of ERα activity Chemical structures of some common ERα ligands. BSC, basic side chain. E2‐rings are numbered A‐D. The E‐ring is the common site of attachment for BSC found in many SERMS. ERα domain organization lettered, A‐F. DBD, DNA‐binding domain; LBD, ligand‐binding domain; AF, activation function Schematic illustration of the canonical ERα signaling pathway. Linear causality model for ERα‐mediated cell proliferation. Branched causality model for ERα‐mediated cell proliferation. ERα contains structurally conserved globular domains of the nuclear receptor superfamily, including a DNA‐binding domain (DBD) that is connected by a flexible hinge region to the ligand‐binding domain (LBD), as well as unstructured AB and F domains at its amino and carboxyl termini, respectively (Fig 1B). The LBD contains a ligand‐dependent coactivator‐binding site called activation function‐2 (AF‐2). However, the agonist activity of SERMs derives from activation function‐1 (AF‐1)—a coactivator recruitment site located in the AB domain (Berry et al, 1990; Shang \u0026 Brown, 2002; Abot et al, 2013). AF‐1 and AF‐2 bind distinct but overlapping sets of coregulators (Webb et al, 1998; Endoh et al, 1999; Delage‐Mourroux et al, 2000; Yi et al, 2015). AF‐2 binds the signature LxxLL motif peptides of coactivators such as NCOA1/2/3 (also known as SRC‐1/2/3). AF‐1 binds a separate surface on these coactivators (Webb et al, 1998; Yi et al, 2015). Yet, it is unknown how different ERα ligands control AF‐1 through the LBD, and whether this inter‐domain communication is required for cell‐specific signaling or anti‐proliferative responses. In the canonical model of the ERα signaling pathway (Fig 1C), E2‐bound ERα forms a homodimer that binds DNA at estrogen‐response elements (EREs), recruits NCOA1/2/3 (Metivier et al, 2003; Johnson \u0026 O'Malley, 2012), and activates the GREB1 gene, which is required for proliferation of ERα‐positive breast cancer cells (Ghosh et al, 2000; Rae et al, 2005; Deschenes et al, 2007; Liu et al, 2012; Srinivasan et al, 2013). However, ERα‐mediated proliferative responses vary in a ligand‐dependent manner (Srinivasan et al, 2013); thus, it is not known whether this canonical model is widely applicable across diverse ERα ligands. Our long‐term goal is to be able to predict proliferative or anti‐proliferative activity of a ligand in different tissues from its crystal structure by identifying different structural perturbations that lead to specific signaling outcomes. The simplest response model for ligand‐specific proliferative effects is a linear causality model, where the degree of NCOA1/2/3 recruitment determines GREB1 expression, which in turn drives ligand‐specific cell proliferation (Fig 1D). Alternatively, a more complicated branched causality model could explain ligand‐specific proliferative responses (Fig 1E). In this signaling model, multiple coregulator binding events and target genes (Won Jeong et al, 2012; Nwachukwu et al, 2014), LBD conformation, nucleocytoplasmic shuttling, the occupancy and dynamics of DNA binding, and other biophysical features could contribute independently to cell proliferation (Lickwar et al, 2012). To test these signaling models, we profiled a diverse library of ERα ligands using systems biology approaches to X‐ray crystallography and chemical biology (Srinivasan et al, 2013), including a series of quantitative bioassays for ERα function that were statistically robust and reproducible, based on the Z’‐statistic (Fig EV1A and B; see Materials and Methods). We also determined the structures of 76 distinct ERα LBD complexes bound to different ligand types, which allowed us to understand how diverse ligand scaffolds distort the active conformation of the ERα LBD. Our findings here indicate that specific structural perturbations can be tied to ligand‐selective domain usage and signaling patterns, thus providing a framework for structure‐based design of improved breast cancer therapeutics, and understanding the different phenotypic effects of environmental estrogens. High‐throughput screens for ERα ligand profiling Summary of ligand screening assays used to measure ER‐mediated activities. ERE, estrogen‐response element; Luc, luciferase reporter gene; M2H, mammalian 2‐hybrid; UAS, upstream‐activating sequence. Controls for screening assays described in panel (A), above. Error bars indicate mean ± SEM, n = 3. Results Strength of AF‐1 signaling does not determine cell‐specific signaling To compare ERα signaling induced by diverse ligand types, we synthesized and assayed a library of 241 ERα ligands containing 19 distinct molecular scaffolds. These include 15 indirect modulator series, which lack a SERM‐like side chain and modulate coactivator binding indirectly from the ligand‐binding pocket (Fig 2A–E; Dataset EV1) (Zheng et al, 2012) (Zhu et al, 2012) (Muthyala et al, 2003; Seo et al, 2006) (Srinivasan et al, 2013) (Wang et al, 2012) (Liao et al, 2014) (Min et al, 2013). We also generated four direct modulator series with side chains designed to directly dislocate h12 and thereby completely occlude the AF‐2 surface (Fig 2C and E; Dataset EV1) (Kieser et al, 2010). Ligand profiling using our quantitative bioassays revealed a wide range of ligand‐induced GREB1 expression, reporter gene activities, ERα‐coactivator interactions, and proliferative effects on MCF‐7 breast cancer cells (Figs EV1 and EV2A–J). This wide variance enabled us to probe specific features of ERα signaling using ligand class analyses, and identify signaling patterns shared by specific ligand series or scaffolds. Classes of compounds in the ERα ligand library Structure of the E2‐bound ERα LBD in complex with an NCOA2 peptide of (PDB 1GWR). Structural details of the ERα LBD bound to the indicated ligands. Unlike E2 (PDB 1GWR), TAM is a direct modulator with a BSC that dislocates h12 to block the NCOA2‐binding site (PDB 3ERT). OBHS is an indirect modulator that dislocates the h11 C‐terminus to destabilize the h11–h12 interface (PDB 4ZN9). The ERα ligand library contains 241 ligands representing 15 indirect modulator scaffolds, plus 4 direct modulator scaffolds. The number of compounds per scaffold is shown in parentheses (see Dataset EV1 for individual compound information and Appendix Supplementary Methods for synthetic protocols). ERα ligands induced a range of agonist activity profiles Screening data from individual ligands are shown, grouped by scaffold. Each data point represents the activity of a distinct compound. Error bars indicate the class average (mean) ± range. *Direct modulator. \nSource data are available online for this figure.\n We first asked whether direct modulation of the receptor with an extended side chain is required for cell‐specific signaling. To this end, we compared the average ligand‐induced GREB1 mRNA levels in MCF‐7 cells and 3×ERE‐Luc reporter gene activity in Ishikawa endometrial cancer cells (E‐Luc) or in HepG2 cells transfected with wild‐type ERα (L‐Luc ERα‐WT) (Figs 3A and EV2A–C). Direct modulators showed significant differences in average activity between cell types except OBHS‐ASC analogs, which had similar low agonist activities in the three cell types. The other direct modulators had low agonist activity in Ishikawa cells, no or inverse agonist activity in MCF‐7 cells, and more variable activity in HepG2 liver cells. While it was known that direct modulators such as tamoxifen drive cell‐specific signaling, these experiments reveal that indirect modulators also drive cell‐specific signaling, since eight of fourteen classes showed significant differences in average activity (Figs 3A and EV2A–C). Ligand‐specific signaling underlies ERα‐mediated cell proliferation (A) Ligand‐specific ERα activities in HepG2, Ishikawa and MCF‐7 cells. The ligand‐induced L‐Luc ERα‐WT and E‐Luc activities and GREB1 mRNA levels are shown by scaffold (mean + SD). (B) Ligand class analysis of the L‐Luc ERα‐WT and ERα‐ΔAB activities in HepG2 cells. Significant sensitivity to AB domain deletion was determined by Student's t‐test (n = number of ligands per scaffold in Fig 2). The average activities of ligands classes are shown (mean + SEM). Correlation and regression analyses in a large test set. The r\n2 values are plotted as a heat map. In cluster 1, the first three comparisons (rows) showed significant positive correlations (F‐test for nonzero slope, P ≤ 0.05). In cluster 2, only one of these comparisons revealed a significant positive correlation, while none was significant in cluster 3. +, statistically significant correlations gained by deletion of the AB or F domains. −, significant correlations lost upon deletion of AB or F domains. \nSource data are available online for this figure.\n Tamoxifen depends on AF‐1 for its cell‐specific activity (Sakamoto et al, 2002); therefore, we asked whether cell‐specific signaling observed here is due to a similar dependence on AF‐1 for activity (Fig EV1). To test this idea, we compared the average L‐Luc activities of each scaffold in HepG2 cells co‐transfected with wild‐type ERα or with ERα lacking the AB domain (Figs 1B and EV1). While E2 showed similar L‐Luc ERα‐WT and ERα‐ΔAB activities, tamoxifen showed complete loss of activity without the AB domain (Fig EV1B). Deletion of the AB domain significantly reduced the average L‐Luc activities of 14 scaffolds (Student's t‐test, P ≤ 0.05) (Fig 3B). These “AF‐1‐sensitive” activities were exhibited by both direct and indirect modulators, and were not limited to scaffolds that showed cell‐specific signaling (Fig 3A and B). Thus, the strength of AF‐1 signaling does not determine cell‐specific signaling. Identifying cell‐specific signaling clusters in ERα ligand classes As another approach to identifying cell‐specific signaling, we determined the degree of correlation between ligand‐induced activities in the different cell types. Here, we compared ligands within each class (Fig 3C), instead of comparing average activities (Fig 3A and B). For each ligand class or scaffold, we calculated the Pearson's correlation coefficient, r, for pairwise comparison of activity profiles in breast (GREB1), liver (L‐Luc), and endometrial cells (E‐Luc). The value of r ranges from −1 to 1, and it defines the extent to which the data fit a straight line when compounds show similar agonist/antagonist activity profiles between cell types (Fig EV3A). We also calculated the coefficient of determination, r 2, which describes the percentage of variance in a dependent variable such as proliferation that can be predicted by an independent variable such as GREB1 expression. We present both calculations as r 2 to readily compare signaling specificities using a heat map on which the red–yellow palette indicates significant positive correlations (P ≤ 0.05, F‐test for nonzero slope), while the blue palette denotes negative correlations (Fig 3C–F). The side chain of OBHS‐BSC analogs induces cell‐specific signaling Correlation analysis of OBHS versus OBHS‐BSC activity across cell types. Correlation analysis of L‐Luc ERα‐ΔAB activity versus endogenous ERα activity of OBHS analogs. In panel (D), L‐Luc ERα‐WT activity from panel (B) is shown for comparison. Correlation analysis of L‐Luc ERα‐ΔF activity versus endogenous ERα activities of OBHS analogs. Correlation analysis of MCF‐7 cell proliferation versus NCOA2/3 recruitment or GREB1 levels observed in response to (G) OBHS‐N and (H) OBHS‐BSC analogs. \nData information: In each panel, a data point indicates the activity of a distinct compound.Source data are available online for this figure.\n This analysis revealed diverse signaling specificities that we grouped into three clusters. Scaffolds in cluster 1 exhibited strongly correlated GREB1 levels, E‐Luc and L‐Luc activity profiles across the three cell types (Fig 3C lanes 1–4), suggesting these ligands use similar ERα signaling pathways in the breast, endometrial, and liver cell types. This cluster includes WAY‐C, OBHS, OBHS‐N, and triaryl‐ethylene analogs, all of which are indirect modulators. Cluster 2 contains scaffolds with activities that were positively correlated in only two of the three cell types, indicating cell‐specific signaling (Fig 3C lanes 5–12). This cluster includes two classes of direct modulators (cyclofenil‐ASC and WAY dimer), and six classes of indirect modulators (2,5‐DTP, 3,4‐DTP, S‐OBHS‐2 and S‐OBHS‐3, furan, and WAY‐D). In this cluster, the correlated activities varied by scaffold. For example, 3,4‐DTP, furan, and S‐OBHS‐2 drove positively correlated GREB1 levels and E‐Luc but not L‐Luc ERα‐WT activity (Fig 3C lanes 5–7). In contrast, WAY dimer and WAY‐D analogs drove positively correlated GREB1 levels and L‐Luc ERα‐WT but not E‐Luc activity (Fig 3C lanes 8 and 9). The last set of scaffolds, cluster 3, displayed cell‐specific activities that were not correlated in any of the three cell types (Fig 3C lanes 13–19). This cluster includes two direct modulator scaffolds (OBHS‐ASC and OBHS‐BSC), and five indirect modulator scaffolds (A‐CD, cyclofenil, 3,4‐DTPD, imine, and imidazopyridine). These results suggest that addition of an extended side chain to an ERα ligand scaffold is sufficient to induce cell‐specific signaling, where the relative activity profiles of the individual ligands change between cell types. This is demonstrated by directly comparing the signaling specificities of matched OBHS (indirect modulator, cluster 1) and OBHS‐BSC analogs (direct modulator, cluster 3), which differ only in the basic side chain (Fig 2E). The activities of OBHS analogs were positively correlated across the three cell types, but the side chain of OBHS‐BSC analogs was sufficient to abolish these correlations (Figs 3C lanes 1 and 19, and EV3A–C). The indirect modulator scaffolds in clusters 2 and 3 showed cell‐specific signaling patterns without the extended side chain typically viewed as the primary chemical and structural mechanism driving cell‐specific activity. Many of these scaffolds drove similar average activities of the ligand class in the different cell types (Fig 3A), but the individual ligands in each class had different cell‐specific activities (Fig EV2A–C). Thus, examining the correlated patterns of ERα activity within each scaffold demonstrates that an extended side chain is not required for cell‐specific signaling. Modulation of signaling specificity by AF‐1 To evaluate the role of AF‐1 and the F domain in ERα signaling specificity, we compared activity of truncated ERα constructs in HepG2 liver cells with endogenous ERα activity in the other cell types. The positive correlation between the L‐Luc and E‐Luc activities or GREB1 levels induced by scaffolds in cluster 1 was generally retained without the AB domain, or the F domain (Fig 3D lanes 1–4). This demonstrates that the signaling specificities underlying these positive correlations are not modified by AF‐1. OBHS analogs showed an average L‐Luc ERα‐ΔAB activity of 3.2% ± 3 (mean + SEM) relative to E2. Despite this nearly complete lack of activity, the pattern of L‐Luc ERα‐ΔAB activity was still highly correlated with the E‐Luc activity and GREB1 expression (Fig EV3D and E), demonstrating that very small AF‐2 activities can be amplified by AF‐1 to produce robust signals. Similarly, deletion of the F domain did not abolish correlations between the L‐Luc and E‐Luc or GREB1 levels induced by OBHS analogs (Fig EV3F). These similar patterns of ligand activity in the wild‐type and deletion mutants suggest that AF‐1 and the F domain purely amplify the AF‐2 activities of ligands in cluster 1. In contrast, AF‐1 was a determinant of signaling specificity for scaffolds in cluster 2. Deletion of the AB or F domain altered correlations for six of the eight scaffolds in this cluster (2,5‐DTP, 3,4‐DTP, S‐OBHS‐3, WAY‐D, WAY dimer, and cyclofenil‐ASC) (Fig 3D lanes 5–12). Comparing Fig 3C and D, the + and − signs indicate where the deletion mutant assays led to a gain or loss of statically significant correlation, respectively. Thus, in cluster 2, AF‐1 substantially modulated the specificity of ligands with cell‐specific activity (Fig 3D lanes 5–12). For ligands in cluster 3, we could not eliminate a role for AF‐1 in determining signaling specificity, since this cluster lacked positively correlated activity profiles (Fig 3C), and deletion of the AB or F domain rarely induced such correlations (Fig 3D), except for A‐CD and OBHS‐ASC analogs, where deletion of the AB domain or F domain led to positive correlations with E‐Luc activity and/or GREB1 levels (Fig 3D lanes 13 and 18). Thus, ligands in cluster 2 rely on AF‐1 for both activity (Fig 3B) and signaling specificity (Fig 3D). As discussed below, this cell specificity derives from alternate coactivator preferences. Ligand‐specific control of GREB1 expression To determine whether ligand classes control expression of native ERα target genes through the canonical linear signaling pathway, we performed pairwise linear regression analyses using ERα–NCOA1/2/3 interactions in M2H assay as independent predictors of GREB1 expression (the dependent variable) (Figs EV1 and EV2A, F–H). In cluster 1, the recruitment of NCOA1 and NCOA2 was highest for WAY‐C, followed by triaryl‐ethylene, OBHS‐N, and OBHS series, while for NCOA3, OBHS‐N compounds induced the most recruitment and OBHS ligands were inverse agonists (Fig EV2F–H). The average induction of GREB1 by cluster 1 ligands showed greater variance, with a range between ~25 and ~75% for OBHS and a range from full agonist to inverse agonist for the others in cluster 1 (Fig EV2A). GREB1 levels induced by OBHS analogs were determined by recruitment of NCOA1 but not NCOA2/3 (Fig 3E lane 1), suggesting that there may be alternate or preferential use of these coactivators by different classes. However, in cluster 1, NCOA1/2/3 recruitment generally predicted GREB1 levels (Fig 3E lanes 1–4), consistent with the canonical signaling model (Fig 1D). For clusters 2 and 3, GREB1 activity was generally not predicted by NCOA1/2/3 recruitment. Direct modulators showed low NCOA1/2/3 recruitment (Fig EV2F–H), but only OBHS‐ASC analogs had NCOA2 recruitment profiles that predicted a full range of effects on GREB1 levels (Figs 3E lanes 9, 11, 18–19, and EV2A). The indirect modulators in clusters 2 and 3 stimulated NCOA1/2/3 recruitment and GREB1 expression with substantial variance (Figs 3A and EV2F–H). However, ligand‐induced GREB1 levels were generally not determined by NCOA1/2/3 recruitment (Fig 3E lanes 5–19), consistent with an alternate causality model (Fig 1E). Out of 11 indirect modulator series in cluster 2 or 3, only the S‐OBHS‐3 class had NCOA1/2/3 recruitment profiles that predicted GREB1 levels (Fig 3E lane 12). These results suggest that compounds that show cell‐specific signaling do not activate GREB1, or use coactivators other than NCOA1/2/3 to control GREB1 expression (Fig 1E). Ligand‐specific control of cell proliferation To determine mechanisms for ligand‐dependent control of breast cancer cell proliferation, we performed linear regression analyses across the 19 scaffolds using MCF‐7 cell proliferation as the dependent variable, and the other activities as independent variables (Fig 3F). In cluster 1, E‐Luc and L‐Luc activities, NCOA1/2/3 recruitment, and GREB1 levels generally predicted the proliferative response (Fig 3F lanes 2–4). With the OBHS‐N compounds, NCOA3 and GREB1 showed near perfect prediction of proliferation (Fig EV3G), with unexplained variance similar to the noise in the assays. The lack of significant predictors for OBHS analogs (Fig 3F lane 1) reflects their small range of proliferative effects on MCF‐7 cells (Fig EV2I). The significant correlations with GREB1 expression and NCOA1/2/3 recruitment observed in this cluster are consistent with the canonical signaling model (Fig 1D), where NCOA1/2/3 recruitment determines GREB1 expression, which then drives proliferation. Ligands in cluster 2 and cluster 3 showed a wide range of proliferative effects on MCF‐7 cells (Fig EV2I). Despite this phenotypic variance, proliferation was not generally predicted by correlated NCOA1/2/3 recruitment and GREB1 induction (Figs 3F lanes 5–19, and EV3H). Out of 15 ligand series in these clusters, only 2,5‐DTP analogs induced a proliferative response that was predicted by GREB1 levels, which were not determined by NCOA1/2/3 recruitment (Fig 3E and F lane 10). 3,4‐DTP, cyclofenil, 3,4‐DTPD, and imidazopyridine analogs had NCOA1/3 recruitment profiles that predicted their proliferative effects, without determining GREB1 levels (Fig 3E and F, lanes 5 and 14–16). Similarly, S‐OBHS‐3, cyclofenil‐ASC, and OBHS‐ASC had positively correlated NCOA1/2/3 recruitment and GREB1 levels, but none of these activities determined their proliferative effects (Fig 3E and F lanes 11–12 and 18). For ligands that show cell‐specific signaling, ERα‐mediated recruitment of other coregulators and activation of other target genes likely determine their proliferative effects on MCF‐7 cells. NCOA3 occupancy at GREB1 did not predict the proliferative response We also questioned whether promoter occupancy by coactivators is statistically robust and reproducible for ligand class analysis using a chromatin immunoprecipitation (ChIP)‐based quantitative assay, and whether it has a better predictive power than the M2H assay. ERα and NCOA3 cycle on and off the GREB1 promoter (Nwachukwu et al, 2014). Therefore, we first performed a time‐course study, and found that E2 and the WAY‐C analog, AAPII‐151‐4, induced recruitment of NCOA3 to the GREB1 promoter in a temporal cycle that peaked after 45 min in MCF‐7 cells (Fig 4A). At this time point, other WAY‐C analogs also induced recruitment of NCOA3 at this site to varying degrees (Fig 4B). The Z’ for this assay was 0.6, showing statistical robustness (see Materials and Methods). We prepared biological replicates with different cell passage numbers and separately prepared samples, which showed r 2 of 0.81, demonstrating high reproducibility (Fig 4C). \nNCOA3 occupancy at GREB1 is statistically robust but does not predict transcriptional activity Kinetic ChIP assay examining recruitment of NCOA3 to the GREB1 gene in MCF‐7 cells stimulated with E2 or the indicated WAY‐C analog. The average of duplicate experiments (mean ± SEM) is shown. NCOA3 occupancy at GREB1 was compared by ChIP assay 45 min after stimulation with vehicle, E2, or the WAY‐C analogs. In panel (B), the average recruitment of two biological replicates are shown as mean + SEM, and the Z‐score is indicated. In panel (C), correlation analysis was performed for two biological replicates. Linear regression analyses comparing the ability of NCOA3 recruitment, measured by ChIP or M2H, to predict other agonist activities of WAY‐C analogs. *Significant positive correlation (F‐test for nonzero slope, P‐value). \nSource data are available online for this figure.\n The M2H assay for NCOA3 recruitment broadly correlated with the other assays, and was predictive for GREB1 expression and cell proliferation (Fig 3E). However, the ChIP assays for WAY‐C‐induced recruitment of NCOA3 to the GREB1 promoter did not correlate with any of the other WAY‐C activity profiles (Fig 4D), although the positive correlation between ChIP assays and NCOA3 recruitment via M2H assay showed a trend toward significance with r 2 = 0.36 and P = 0.09 (F‐test for nonzero slope). Thus, the simplified coactivator‐binding assay showed much greater predictive power than the ChIP assay for ligand‐specific effects on GREB1 expression and cell proliferation. ERβ activity is not an independent predictor of cell‐specific activity One difference between MCF‐7 breast cancer cells and Ishikawa endometrial cancer cells is the contribution of ERβ to estrogenic response, as Ishikawa cells may express ERβ (Bhat \u0026 Pezzuto, 2001). When overexpressed in MCF‐7 cells, ERβ alters E2‐induced expression of only a subset of ERα‐target genes (Wu et al, 2011), raising the possibility that ligand‐induced ERβ activity may contribute to E‐Luc activities, and thus underlie the lack of correlation between the E‐Luc and L‐Luc ERα‐WT activities or GREB1 levels induced by cell‐specific modulators in cluster 2 and cluster 3 (Fig 3C). To test this idea, we determined the L‐Luc ERβ activity profiles of the ligands (Fig EV1). All direct modulator and two indirect modulator scaffolds (OBHS and S‐OBHS‐3) lacked ERβ agonist activity. However, the other ligands showed a range of ERβ activities (Fig EV2J). For most scaffolds, L‐Luc ERβ and E‐Luc activities were not correlated, except for 2,5‐DTP and cyclofenil analogs, which showed moderate but significant correlations (Fig EV4A). Nevertheless, the E‐Luc activities of both 2,5‐DTP and cyclofenil analogs were better predicted by their L‐Luc ERα‐WT than L‐Luc ERβ activities (Fig EV4A and B). Thus, ERβ activity was not an independent determinant of the observed activity profiles. ERβ activity is not an independent predictor of E‐Luc activity ERβ activity in HepG2 cells rarely correlates with E‐Luc activity. ERα activity of 2,5‐DTP and cyclofenil analogs correlates with E‐Luc activity. \nData information: The r\n2 and P values for the indicated correlations are shown in both panels. *Significant positive correlation (F‐test for nonzero slope, P‐value) Structural features of consistent signaling across cell types To overcome barriers to crystallization of ERα LBD complexes, we developed a conformation‐trapping X‐ray crystallography approach using the ERα‐Y537S mutation (Nettles et al, 2008; Bruning et al, 2010; Srinivasan et al, 2013). To further validate this approach, we solved the structure of the ERα‐Y537S LBD in complex with diethylstilbestrol (DES), which bound identically in the wild‐type and ERα‐Y537S LBDs, demonstrating again that this surface mutation stabilizes h12 dynamics to facilitate crystallization without changing ligand binding (Appendix Fig S1A and B) (Nettles et al, 2008; Bruning et al, 2010; Delfosse et al, 2012). Using this approach, we solved 76 ERα LBD structures in the active conformation and bound to ligands studied here (Appendix Fig S1C). Eleven of these structures have been published, while 65 are new, including the DES‐bound ERα‐Y537S LBD. We present 57 of these new structures here (Dataset EV2), while the remaining eight new structures bound to OBHS‐N analogs will be published elsewhere (S. Srinivasan et al, in preparation). Examining many closely related structures allows us to visualize subtle structural differences, in effect using X‐ray crystallography as a systems biology tool. The indirect modulator scaffolds in cluster 1 did not show cell‐specific signaling (Fig 3C), but shared common structural perturbations that we designed to modulate h12 dynamics. Based on our original OBHS structure, the OBHS, OBHS‐N, and triaryl‐ethylene compounds were modified with h11‐directed pendant groups (Zheng et al, 2012; Zhu et al, 2012; Liao et al, 2014). Superposing the LBDs based on the class of bound ligands provides an ensemble view of the structural variance and clarifies what part of the ligand‐binding pocket is differentially perturbed or targeted. The 24 structures containing OBHS, OBHS‐N, or triaryl‐ethylene analogs showed structural diversity in the same part of the scaffolds (Figs 5A and EV5A), and the same region of the LBD—the C‐terminal end of h11 (Figs 5B and C, and EV5B), which in turn nudges h12 (Fig 5C and D). We observed that the OBHS‐N analogs displaced h11 along a vector away from Leu354 in a region of h3 that is unaffected by the ligands, and toward the dimer interface. For the triaryl‐ethylene analogs, the displacement of h11 was in a perpendicular direction, away from Ile424 in h8 and toward h12. Remarkably, these individual inter‐atomic distances showed a ligand class‐specific ability to significantly predict proliferative effects (Fig 5E and F), demonstrating the feasibility of developing a minimal set of activity predictors from crystal structures. Structural determinants of consistent signaling Structure‐class analysis of triaryl‐ethylene analogs. Triaryl‐ethylene analogs bound to the superposed crystal structures of the ERα LBD are shown. Arrows indicate chemical variance in the orientation of the different h11‐directed ligand side groups (PDB 5DK9, 5DKB, 5DKE, 5DKG, 5DKS, 5DL4, 5DLR, 5DMC, 5DMF and 5DP0). Triaryl‐ethylene analogs induce variance of ERα conformations at the C‐terminal region of h11. Panel (B) shows the crystal structure of a triaryl‐ethylene analog‐bound ERα LBD (PDB 5DLR). The h11–h12 interface (circled) includes the C‐terminal part of h11. This region was expanded in panel (C), where the 10 triaryl‐ethylene analog‐bound ERα LBD structures (see Datasets EV1 and EV2) were superposed to show variations in the h11 C‐terminus (PDB 5DK9, 5DKB, 5DKE, 5DKG, 5DKS, 5DL4, 5DLR, 5DMC, 5DMF, and 5DP0). ERα LBDs in complex with diethylstilbestrol (DES) or a triaryl‐ethylene analog were superposed to show that the ligand‐induced difference in h11 conformation is transmitted to the C‐terminus of h12 (PDB 4ZN7, 5DMC). Inter‐atomic distances predict the proliferative effects of specific ligand series. Ile424–His524 distance measured in the crystal structures correlates with the proliferative effect of triaryl‐ethylene analogs in MCF‐7 cells. In contrast, the Leu354–Leu525 distance correlates with the proliferative effects of OBHS‐N analogs in MCF‐7 cells. Structure‐class analysis of WAY‐C analogs. WAY‐C side groups subtly nudge h12 Leu540. ERα LBD structures bound to 4 distinct WAY‐C analogs were superposed (PDB 4 IU7, 4IV4, 4IVW, 4IW6) (see Datasets EV1 and EV2). \nSource data are available online for this figure.\n Structure‐class analysis of indirect modulators Structure‐class analysis of indirect modulators in cluster 1. Crystal structures of the ERα LBD bound to OBHS and OBHS‐N analogs were superposed. The bound ligands are shown in panel (A). Arrows indicate chemical variance in the orientation of the different h11‐directed ligand side groups. Panel (B) shows the ligand‐induced conformational variation at the C‐terminal region of h11 (OBHS: PDB 4ZN9, 4ZNH, 4ZNS, 4ZNT, 4ZNU, 4ZNV, and 4ZNW; OBHS‐N: PDB 4ZUB, 4ZUC, 4ZWH, 4ZWK, 5BNU, 5BP6, 5BPR, and 5BQ4). Structure‐class analysis of indirect modulators in clusters 2 and 3. Crystal structures of the ERα LBD bound to ligands with cell‐specific activities were superposed. The bound ligands are shown, and arrows indicate considerable variation in the orientation of the different h3‐, h8‐, h11‐, or h12‐directed ligand side groups. As visualized in four LBD structures (Srinivasan et al, 2013), WAY‐C analogs were designed with small substitutions that slightly nudge h12 Leu540, without exiting the ligand‐binding pocket (Fig 5G and H). Therefore, changing h12 dynamics maintains the canonical signaling pathway defined by E2 (Fig 1D) to support AF‐2‐driven signaling and recruit NCOA1/2/3 for GREB1‐stimulated proliferation. Ligands with cell‐specific activity alter the shape of the AF‐2 surface Direct modulators like tamoxifen drive AF‐1‐dependent cell‐specific activity by completely occluding AF‐2, but it is not known how indirect modulators produce cell‐specific ERα activity. Therefore, we examined another 50 LBD structures containing ligands in clusters 2 and 3. These structures demonstrated that cell‐specific activity derived from altering the shape of the AF‐2 surface without an extended side chain. Ligands in cluster 2 and cluster 3 showed conformational heterogeneity in parts of the scaffold that were directed toward multiple regions of the receptor including h3, h8, h11, h12, and/or the β‐sheets (Fig EV5C–G). For instance, S‐OBHS‐2 and S‐OBHS‐3 analogs (Fig 2) had similar ERα activity profiles in the different cell types (Fig EV2A–C), but the 2‐ versus 3‐methyl substituted phenol rings altered the correlated signaling patterns in different cell types (Fig 3B lanes 7 and 12). Structurally, the 2‐ versus 3‐methyl substitutions changed the binding position of the A‐ and E‐ring phenols by 1.0 Å and 2.2 Å, respectively (Fig EV5C). This difference in ligand positioning altered the AF‐2 surface via a shift in the N‐terminus of h12, which directly contacts the coactivator. This effect is evident in a single structure due to its 1 Å magnitude (Fig 6A and B). The shifts in h12 residues Asp538 and Leu539 led to rotation of the coactivator peptide (Fig 6C). Thus, cell‐specific activity can stem from perturbation of the AF‐2 surface without an extended side chain, which presumably alters the receptor–coregulator interaction profile. Structural correlates of cell‐specific signaling S‐OBHS‐2/3 analogs subtly distort the AF‐2 surface. Panel (A) shows the crystal structure of an S‐OBHS‐3‐bound ERα LBD (PDB 5DUH). The h3–h12 interface (circled) at AF‐2 (pink) was expanded in panels (B, C). The S‐OBHS‐2/3‐bound ERα LBDs were superposed to show shifts in h3 (panel B) and the NCOA2 peptide docked at the AF‐2 surface (panel C). Crystal structures show that 2,5‐DTP analogs shift h3 and h11 further apart compared to an A‐CD‐ring estrogen (PDB 4PPS, 5DRM, 5DRJ). The 2F\no‐F\nc electron density map and F\no‐F\nc difference map of a 2,5‐DTP‐bound structure (PDB 5DRJ) were contoured at 1.0 sigma and ± 3.0 sigma, respectively. Average (mean + SEM) α‐carbon distance measured from h3 Thr347 to h11 Leu525 of A‐CD‐, 2,5‐DTP‐, and 3,4‐DTPD‐bound ERα LBDs. *Two‐tailed Student's t‐test, P = 0.002 (PDB A‐CD: 5DI7, 5DID, 5DIE, 5DIG, and 4PPS; 2,5‐DTP: 4IWC, 5DRM, and 5DRJ; 3,4‐DTPD: 5DTV and 5DU5). Crystal structures show that a 3,4‐DTPD analog shifts h3 (F) and the NCOA2 (G) peptide compared to an A‐CD‐ring estrogen (PDB 4PPS, 5DTV). Hierarchical clustering of ligand‐specific binding of 154 interacting peptides to the ERα LBD was performed in triplicate by MARCoNI analysis. \nSource data are available online for this figure.\n The 2,5‐DTP analogs showed perturbation of h11, as well as h3, which forms part of the AF‐2 surface. These compounds bind the LBD in an unusual fashion because they have a phenol‐to‐phenol length of ~12 Å, which is longer than steroids and other prototypical ERα agonists that are ~10 Å in length. One phenol pushed further toward h3 (Fig 6D), while the other phenol pushed toward the C‐terminus of h11 to a greater extent than A‐CD‐ring estrogens (Nwachukwu et al, 2014), which are close structural analogs of E2 that lack a B‐ring (Fig 2). To quantify this difference, we compared the distance between α‐carbons at h3 Thr347 and h11 Leu525 in the set of structures containing 2,5‐DTP analogs (n = 3) or A‐CD‐ring analogs (n = 5) (Fig 6E). We observed a difference of 0.4 Å that was significant (two‐tailed Student's t‐test, P = 0.002) due to the very tight clustering of the 2,5‐DTP‐induced LBD conformation. The shifts in h3 suggest these compounds are positioned to alter coregulator preferences. The 2,5‐DTP and 3,4‐DTP scaffolds are isomeric, but with aryl groups at obtuse and acute angles, respectively (Fig 2). The crystal structure of ERα in complex with a 3,4‐DTP is unknown; however, we solved two crystal structures of ERα bound to 3,4‐DTPD analogs and one structure containing a furan ligand—all of which have a 3,4‐diaryl configuration (Fig 2; Datasets EV1 and EV2). In these structures, the A‐ring mimetic of the 3,4‐DTPD scaffold bound h3 Glu353 as expected, but the other phenol wrapped around h3 to form a hydrogen bond with Thr347, indicating a change in binding epitopes in the ERα ligand‐binding pocket (Fig 6F). The 3,4‐DTPD analogs also induced a shift in h3 positioning, which translated again into a shift in the bound coactivator peptide (Fig 6F). Therefore, these indirect modulators, including S‐OBHS‐2, S‐OBHS‐3, 2,5‐DTP, and 3,4‐DTPD analogs—all of which show cell‐specific activity profiles—induced shifts in h3 and h12 that were transmitted to the coactivator peptide via an altered AF‐2 surface. To test whether the AF‐2 surface shows changes in shape in solution, we used the microarray assay for real‐time coregulator–nuclear receptor interaction (MARCoNI) analysis (Aarts et al, 2013). Here, the ligand‐dependent interactions of the ERα LBD with over 150 distinct LxxLL motif peptides were assayed to define structural fingerprints for the AF‐2 surface, in a manner similar to the use of phage display peptides as structural probes (Connor et al, 2001). Despite the similar average activities of these ligand classes (Fig 3A and B), 2,5‐DTP and 3,4‐DTP analogs displayed remarkably different peptide recruitment patterns (Fig 6H), consistent with the structural analyses. Hierarchical clustering revealed that many of the 2,5‐DTP analogs recapitulated most of the peptide recruitment and dismissal patterns observed with E2 (Fig 6H). However, there was a unique cluster of peptides that were recruited by E2 but not the 2,5‐DTP analogs. In contrast, 3,4‐DTP analogs dismissed most of the peptides from the AF‐2 surface (Fig 6H). Thus, the isomeric attachment of diaryl groups to the thiophene core changed the AF‐2 surface from inside the ligand‐binding pocket, as predicted by the crystal structures. Together, these findings suggest that without an extended side chain, cell‐specific activity stems from different coregulator recruitment profiles, due to unique ligand‐induced conformations of the AF‐2 surface, in addition to differential usage of AF‐1. Indirect modulators in cluster 1 avoid this by perturbing the h11–h12 interface, and modulating the dynamics of h12 without changing the shape of AF‐2 when stabilized. Discussion Our goal was to identify a minimal set of predictors that would link specific structural perturbations to ERα signaling pathways that control cell‐specific signaling and proliferation. We found a very strong set of predictors, where ligands in cluster 1, defined by similar signaling across cell types, showed indirect modulation of h12 dynamics via the h11–12 interface or slight contact with h12. This perturbation determined proliferation that correlated strongly with AF‐2 activity, recruitment of NCOA1/2/3 family members, and induction of the GREB1 gene, consistent with the canonical ERα signaling pathway (Fig 1D). For ligands in cluster 1, deletion of AF‐1 reduced activity to varying degrees, but did not change the underlying signaling patterns established through AF‐2. In contrast, an extended side chain designed to directly reposition h12 and completely disrupt the AF‐2 surface results in cell‐specific signaling. This was demonstrated with direct modulators in clusters 2 and 3. Cluster 2 was defined by ligand classes that showed correlated activities in two of the three cell types tested, while ligand classes in cluster 3 did not show correlated activities among any of the three cell types. Compared to cluster 1, the structural rules are less clear in clusters 2 and 3, but a number of indirect modulator classes perturbed the LBD conformation at the intersection of h3, the h12 N‐terminus, and the AF‐2 surface. Ligands in these classes altered the shape of AF‐2 to affect coregulator preferences. For direct and indirect modulators in cluster 2 or 3, the canonical ERα signaling pathway involving recruitment of NCOA1/2/3 and induction of GREB1 did not generally predict their proliferative effects, indicating an alternate causal model (Fig 1E). These principles outlined above provide a structural basis for how the ligand–receptor interface leads to different signaling specificities through AF‐1 and AF‐2. It is noteworthy that regulation of h12 dynamics indirectly through h11 can virtually abolish AF‐2 activity, and yet still drive robust transcriptional activity through AF‐1, as demonstrated with the OBHS series. This finding can be explained by the fact that NCOA1/2/3 contain distinct binding sites for interaction with AF‐1 and AF‐2 (McInerney et al, 1996; Webb et al, 1998), which allows ligands to nucleate ERα–NCOA1/2/3 interaction through AF‐2, and reinforce this interaction with additional binding to AF‐1. Completely blocking AF‐2 with an extended side chain or altering the shape of AF‐2 changes the preference away from NCOA1/2/3 for determining GREB1 levels and proliferation of breast cancer cells. AF‐2 blockade also allows AF‐1 to function independently, which is important since AF‐1 drives tissue‐selective effects in vivo. This was demonstrated with AF‐1 knockout mice that show E2‐dependent vascular protection, but not uterine proliferation, thus highlighting the role of AF‐1 in tissue‐selective or cell‐specific signaling (Billon‐Gales et al, 2009; Abot et al, 2013). One current limitation to our approach is the identification of statistical variables that predict ligand‐specific activity. Here, we examined many LBD structures and tested several variables that were not predictive, including ERβ activity, the strength of AF‐1 signaling, and NCOA3 occupancy at the GREB1 gene. Similarly, we visualized structures to identify patterns. There are many systems biology approaches that could contribute to the unbiased identification of predictive variables for statistical modeling. For example, phage display was used to identify the androgen receptor interactome, which was cloned into an M2H library and used to identify clusters of ligand‐selective interactions (Norris et al, 2009). Also, we have used siRNA screening to identify a number of coregulators required for ERα‐mediated repression of the IL‐6 gene (Nwachukwu et al, 2014). However, the use of larger datasets to identify such predictor variables has its own limitations, one of the major ones being the probability of false positives from multiple hypothesis testing. If we calculated inter‐atomic distance matrices containing 4,000 atoms per structure × 76 ligand–receptor complexes, we would have 3 × 105 predictions. One way to address this issue is to use the cross‐validation concept, where hypotheses are generated on training sets of ligands and tested with another set of ligands. Based on this work, we propose several testable hypotheses for drug discovery. We have identified atomic vectors for the OBHS‐N and triaryl‐ethylene classes that predict ligand response (Fig 5E and F). These ligands in cluster 1 drive consistent, canonical signaling across cell types, which is desirable for generating full antagonists. Indeed, the most anti‐proliferative compound in the OBHS‐N series had a fulvestrant‐like profile across a battery of assays (S. Srinivasan et al, in preparation). Secondly, our finding that WAY‐C compounds do not rely of AF‐1 for signaling efficacy may derive from the slight contacts with h12 observed in crystal structures (Figs 3B and 5H), unlike other compounds in cluster 1 that dislocate h11 and rely on AF‐1 for signaling efficacy (Figs 3B and 5C, and EV5B). Thirdly, we found ligands that achieved cell‐specific activity without a prototypical extended side chain. Some of these ligands altered the shape of the AF‐2 surface by perturbing the h3–h12 interface, thus providing a route to new SERM‐like activity profiles by combining indirect and direct modulation of receptor structure. Incorporation of statistical approaches to understand relationships between structure and signaling variables moves us toward predictive models for complex ERα‐mediated responses such as in vivo uterine proliferation or tumor growth, and more generally toward structure‐based design for other allosteric drug targets including GPCRs and other nuclear receptors. Materials and Methods Statistical analysis Correlation and linear regression analyses were performed using GraphPad Prism software. For correlation analysis, the degree to which two datasets vary together was calculated with the Pearson correlation coefficient (r). However, we reported r 2 rather than r, to facilitate comparison with the linear regression results for which we calculated and reported r 2 (Fig 3C–F). Significance for r 2 was determined using the F‐test for nonzero slope. High‐throughput assays were considered statistically robust if they show Z’ \u003e 0.5, where Z’ = 1 − (3(σp+σn)/|μp−μn|), for the mean (σ) and standard deviations (μ) of the positive and negative controls (Fig EV1A and B). ERα ligand library The library of compounds examined includes both previously reported (Srinivasan et al, 2013) and newly synthesized compound series (see Dataset EV1 for individual compound information, and Appendix Supplementary Methods for synthetic protocols). Luciferase reporter assays Cells were transfected with FugeneHD reagent (Roche Applied Sciences, Indianapolis, IN) in 384‐well plates. After 24 h, cells were stimulated with 10 μM compounds dispensed using a 100‐nl pintool Biomeck NXP workstation (Beckman Coulter Inc.). Luciferase activity was measured 24 h later (see Appendix Supplementary Methods for more details). Mammalian 2‐hybrid (M2H) assays HEK293T cells were transfected with 5× UAS‐luciferase reporter, and wild‐type ERα‐VP16 activation domain plus full‐length NCOA1/2/3‐GAL4 DBD fusion protein expression plasmids, using the TransIT‐LT1 transfection reagent (Mirus Bio LLC, Madison, WI). The next day, cells were stimulated with 10 μM compounds using a 100‐nl pintool Biomeck NXP workstation (Beckman Coulter Inc.). Luciferase activity was measured after 24 h (see Appendix Supplementary Methods for more details). Cell proliferation assay MCF‐7 cells were plated on 384‐well plates in phenol red‐free media plus 10% FBS and stimulated with 10 μM compounds using 100‐nl pintool Biomeck NXP workstation (Beckman Coulter Inc.). Cell numbers determined 1 week later (see Appendix Supplementary Methods for more details). Quantitative RT–PCR MCF‐7 cells were steroid‐deprived and stimulated with compounds for 24 h. Total RNA was extracted and reverse‐transcribed. The cDNA was analyzed using TaqMan Gene Expression Master Mix (Life Technologies, Grand Island, NY), GREB1 and GAPDH (control) primers, and hybridization probes (see Appendix Supplementary Methods for more details). MARCoNI coregulator‐interaction profiling This assay was performed as previously described with the ERα LBD, 10 μM compounds, and a PamChiP peptide microarray (PamGene International) containing 154 unique coregulator peptides (Aarts et al, 2013) (see Appendix Supplementary Methods for more details). Protein production and X‐ray crystallography ERα protein was produced as previously described (Bruning et al, 2010). New ERα LBD structures (see Dataset EV2 for data collection and refinement statistics) were solved by molecular replacement using PHENIX (Adams et al, 2010), refined using ExCoR as previously described (Nwachukwu et al, 2013), and COOT (Emsley \u0026 Cowtan, 2004) for ligand‐docking and rebuilding. Data availability Crystal structures analyzed in this study include the following: 1GWR (Warnmark et al, 2002), 3ERD and 3ERT (Shiau et al, 1998), 4ZN9 (Zheng et al, 2012), 4IWC, 4 IU7, 4IV4, 4IVW, 4IW6, 4IUI, 4IV2, 4IVY and 4IW8 (Srinivasan et al, 2013), and 4PPS (Nwachukwu et al, 2014). New crystal structures analyzed in this study were deposited in the RCSB protein data bank (http://www.pdb.org): 4ZN7, 4ZNH, 4ZNS, 4ZNT, 4ZNU, 4ZNV, 4ZNW, 5DI7, 5DID, 5DIE, 5DIG, 5DK9, 5DKB, 5DKE, 5DKG, 5DKS, 5DL4, 5DLR, 5DMC, 5DMF, 5DP0, 5DRM, 5DRJ, 5DTV, 5DU5, 5DUE, 5DUG, 5DUH, 5DXK, 5DXM, 5DXP, 5DXQ, 5DXR, 5EHJ, 5DY8, 5DYB, 5DYD, 5DZ0, 5DZ1, 5DZ3, 5DZH, 5DZI, 5E0W, 5E0X, 5E14, 5E15, 5E19, 5E1C, 5DVS, 5DVV, 5DWE, 5DWG, 5DWI, 5DWJ, 5EGV, 5EI1, 5EIT. Author contributions JCN and SS contributed equally to this work. JCN and SS designed and performed experiments and wrote the manuscript; YZ, KEC, SW, JM, CD, ZL, VC, JN, NJW, JSJ, and RH performed experiments; HBZ designed experiments; and JAK and KWN designed experiments and wrote the manuscript. Conflict of Interest The authors declare that they have no conflict of interest. Supporting information References Robust array‐based coregulator binding assay predicting ERalpha‐agonist potency and generating binding profiles reflecting ligand structure The AF‐1 activation function of estrogen receptor alpha is necessary and sufficient for uterine epithelial cell proliferation in vivo PHENIX: a comprehensive Python‐based system for macromolecular structure solution Role of the two activating domains of the oestrogen receptor in the cell‐type and promoter‐context dependent agonistic activity of the anti‐oestrogen 4‐hydroxytamoxifen Resveratrol exhibits cytostatic and antiestrogenic properties with human endometrial adenocarcinoma (Ishikawa) cells The transactivating function 1 of estrogen receptor alpha is dispensable for the vasculoprotective actions of 17beta‐estradiol Coupling of receptor conformation and ligand orientation determine graded activity Circumventing tamoxifen resistance in breast cancers using antiestrogens that induce unique conformational changes in the estrogen receptor Analysis of estrogen receptor interaction with a repressor of estrogen receptor activity (REA) and the regulation of estrogen receptor transcriptional activity by REA Structural and mechanistic insights into bisphenols action provide guidelines for risk assessment and discovery of bisphenol A substitutes Regulation of GREB1 transcription by estrogen receptor alpha through a multipartite enhancer spread over 20 kb of upstream flanking sequences Coot: model‐building tools for molecular graphics Purification and identification of p68 RNA helicase acting as a transcriptional coactivator specific for the activation function 1 of human estrogen receptor alpha Tamoxifen for prevention of breast cancer: report of the National Surgical Adjuvant Breast and Bowel Project P‐1 Study Time‐dependent changes in biochemical bone markers and serum cholesterol in ovariectomized rats: effects of raloxifene HCl, tamoxifen, estrogen, and alendronate PDZK1 and GREB1 are estrogen‐regulated genes expressed in hormone‐responsive breast cancer Steroid receptor coactivators 1, 2, and 3: critical regulators of nuclear receptor activity and steroid receptor modulator (SRM)‐based cancer therapy Antiestrogens and selective estrogen receptor modulators as multifunctional medicines. 1. Receptor interactions Diversity and modularity of G protein‐coupled receptor structures Characterization of the pharmacophore properties of novel selective estrogen receptor downregulators (SERDs) Triaryl‐substituted Schiff bases are high‐affinity subtype‐selective ligands for the estrogen receptor Genome‐wide protein‐DNA binding dynamics suggest a molecular clutch for transcription factor function GREB1 functions as a growth promoter and is modulated by IL6/STAT3 in breast cancer Definition of the molecular and cellular mechanisms underlying the tissue‐selective agonist/antagonist activities of selective estrogen receptor modulators Analysis of estrogen receptor transcriptional enhancement by a nuclear hormone receptor coactivator Estrogen receptor‐alpha directs ordered, cyclical, and combinatorial recruitment of cofactors on a natural target promoter Thiophene‐core estrogen receptor ligands having superagonist activity Bridged bicyclic cores containing a 1,1‐diarylethylene motif are high‐affinity subtype‐selective ligands for the estrogen receptor Ligand control of coregulator recruitment to nuclear receptors NFκB selectivity of estrogen receptor ligands revealed by comparative crystallographic analyses Differential presentation of protein interaction surfaces on the androgen receptor defines the pharmacological actions of bound ligands Improved crystallographic structures using extensive combinatorial refinement Resveratrol modulates the inflammatory response via an estrogen receptor‐signal integration network How many drug targets are there? GREB 1 is a critical regulator of hormone dependent breast cancer growth Estrogen receptor‐mediated effects of tamoxifen on human endometrial cancer cells Fluorine‐substituted cyclofenil derivatives as estrogen receptor ligands: synthesis and structure‐affinity relationship study of potential positron emission tomography agents for imaging estrogen receptors in breast cancer Molecular determinants for the tissue specificity of SERMs The structural basis of estrogen receptor/coactivator recognition and the antagonism of this interaction by tamoxifen Ligand‐binding dynamics rewire cellular signaling via estrogen receptor‐alpha Identification and structure‐activity relationships of a novel series of estrogen receptor ligands based on 7‐thiabicyclo[2.2.1]hept‐2‐ene‐7‐oxide Interaction of transcriptional intermediary factor 2 nuclear receptor box peptides with the coactivator binding site of estrogen receptor alpha Estrogen receptor activation function 1 works by binding p160 coactivator proteins Recent developments in biased agonism Gene‐specific patterns of coregulator requirements by estrogen receptor‐alpha in breast cancer cells Estrogen receptor‐beta sensitizes breast cancer cells to the anti‐estrogenic actions of endoxifen Structure of a biologically active estrogen receptor‐coactivator complex on DNA Development of selective estrogen receptor modulator (SERM)‐like activity through an indirect mechanism of estrogen receptor antagonism: defining the binding mode of 7‐oxabicyclo[2.2.1]hept‐5‐ene scaffold core ligands Bicyclic core estrogens as full antagonists: synthesis, biological evaluation and structure‐activity relationships of estrogen receptor ligands based on bridged oxabicyclic core 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+[{"sourceid":"4850273","sourcedb":"","project":"","target":"","text":"Molecular Dissection of Xyloglucan Recognition in a Prominent Human Gut Symbiont ABSTRACT Polysaccharide utilization loci (PUL) within the genomes of resident human gut Bacteroidetes are central to the metabolism of the otherwise indigestible complex carbohydrates known as “dietary fiber.” However, functional characterization of PUL lags significantly behind sequencing efforts, which limits physiological understanding of the human-bacterial symbiosis. In particular, the molecular basis of complex polysaccharide recognition, an essential prerequisite to hydrolysis by cell surface glycosidases and subsequent metabolism, is generally poorly understood. Here, we present the biochemical, structural, and reverse genetic characterization of two unique cell surface glycan-binding proteins (SGBPs) encoded by a xyloglucan utilization locus (XyGUL) from Bacteroides ovatus, which are integral to growth on this key dietary vegetable polysaccharide. Biochemical analysis reveals that these outer membrane-anchored proteins are in fact exquisitely specific for the highly branched xyloglucan (XyG) polysaccharide. The crystal structure of SGBP-A, a SusD homolog, with a bound XyG tetradecasaccharide reveals an extended carbohydrate-binding platform that primarily relies on recognition of the β-glucan backbone. The unique, tetra-modular structure of SGBP-B is comprised of tandem Ig-like folds, with XyG binding mediated at the distal C-terminal domain. Despite displaying similar affinities for XyG, reverse-genetic analysis reveals that SGBP-B is only required for the efficient capture of smaller oligosaccharides, whereas the presence of SGBP-A is more critical than its carbohydrate-binding ability for growth on XyG. Together, these data demonstrate that SGBP-A and SGBP-B play complementary, specialized roles in carbohydrate capture by B. ovatus and elaborate a model of how vegetable xyloglucans are accessed by the Bacteroidetes. IMPORTANCE The Bacteroidetes are dominant bacteria in the human gut that are responsible for the digestion of the complex polysaccharides that constitute “dietary fiber.” Although this symbiotic relationship has been appreciated for decades, little is currently known about how Bacteroidetes seek out and bind plant cell wall polysaccharides as a necessary first step in their metabolism. Here, we provide the first biochemical, crystallographic, and genetic insight into how two surface glycan-binding proteins from the complex Bacteroides ovatus xyloglucan utilization locus (XyGUL) enable recognition and uptake of this ubiquitous vegetable polysaccharide. Our combined analysis illuminates new fundamental aspects of complex polysaccharide recognition, cleavage, and import at the Bacteroidetes cell surface that may facilitate the development of prebiotics to target this phylum of gut bacteria. INTRODUCTION The human gut microbiota influences the course of human development and health, playing key roles in immune stimulation, intestinal cell proliferation, and metabolic balance. This microbial community is largely bacterial, with the Bacteroidetes, Firmicutes, and Actinobacteria comprising the dominant phyla. The ability to acquire energy from carbohydrates of dietary or host origin is central to the adaptation of human gut bacterial species to their niche. More importantly, this makes diet a tractable way to manipulate the abundance and metabolic output of the microbiota toward improved human health. However, there is a paucity of data regarding how the vast array of complex carbohydrate structures are selectively recognized and imported by members of the microbiota, a critical process that enables these organisms to thrive in the competitive gut environment. The human gut bacteria Bacteroidetes share a profound capacity for dietary glycan degradation, with many species containing \u003e250 predicted carbohydrate-active enzymes (CAZymes), compared to 50 to 100 within many Firmicutes and only 17 in the human genome devoted toward carbohydrate utilization. A remarkable feature of the Bacteroidetes is the packaging of genes for carbohydrate catabolism into discrete polysaccharide utilization loci (PUL), which are transcriptionally regulated by specific substrate signatures. The archetypal PUL-encoded system is the starch utilization system (Sus) (Fig. 1B) of Bacteroides thetaiotaomicron. The Sus includes a lipid-anchored, outer membrane endo-amylase, SusG; a TonB-dependent transporter (TBDT), SusC, which imports oligosaccharides with the help of an associated starch-binding protein, SusD; two additional carbohydrate-binding lipoproteins, SusE and SusF; and two periplasmic exo-glucosidases, SusA and SusB, which generate glucose for transport into the cytoplasm. The importance of PUL as a successful evolutionary strategy is underscored by the observation that Bacteroidetes such as B. thetaiotaomicron and Bacteroides ovatus devote ~18% of their genomes to these systems. Moving beyond seminal genomic and transcriptomic analyses, the current state-of-the-art PUL characterization involves combined reverse-genetic, biochemical, and structural studies to illuminate the molecular details of PUL function. Xyloglucan and the Bacteroides ovatus xyloglucan utilization locus (XyGUL). (A) Representative structures of common xyloglucans using the Consortium for Functional Glycomics Symbol Nomenclature (http://www.functionalglycomics.org/static/consortium/Nomenclature.shtml). Cleavage sites for BoXyGUL glycosidases (GHs) are indicated for solanaceous xyloglucan. (B) BtSus and BoXyGUL. (C) Localization of BoXyGUL-encoded proteins in cellular membranes and concerted modes of action in the degradation of xyloglucans to monosaccharides. The location of SGBP-A/B is presented in this work; the location of GH5 has been empirically determined, and the enzymes have been placed based upon their predicted cellular location. We recently reported the detailed molecular characterization of a PUL that confers the ability of the human gut commensal B. ovatus ATCC 8483 to grow on a prominent family of plant cell wall glycans, the xyloglucans (XyG). XyG variants (Fig. 1A) constitute up to 25% of the dry weight of common vegetables. Analogous to the Sus locus, the xyloglucan utilization locus (XyGUL) encodes a cohort of carbohydrate-binding, -hydrolyzing, and -importing proteins (Fig. 1B and C). The number of glycoside hydrolases (GHs) encoded by the XyGUL is, however, more expansive than that by the Sus locus (Fig. 1B), which reflects the greater complexity of glycosidic linkages found in XyG vis-à-vis starch. Whereas our previous study focused on the characterization of the linkage specificity of these GHs, a key outstanding question regarding this locus is how XyG recognition is mediated at the cell surface. In the archetypal starch utilization system of B. thetaiotaomicron, starch binding to the cell surface is mediated at eight distinct starch-binding sites distributed among four surface glycan-binding proteins (SGBPs): two within the amylase SusG, one within SusD, two within SusE, and three within SusF. The functional redundancy of many of these sites is high: whereas SusD is essential for growth on starch, combined mutations of the SusE, SusF, and SusG binding sites are required to impair growth on the polysaccharide. Bacteroidetes PUL ubiquitously encode homologs of SusC and SusD, as well as proteins whose genes are immediately downstream of susD, akin to susE/F, and these are typically annotated as “putative lipoproteins”. The genes coding for these proteins, sometimes referred to as “susE/F positioned,” display products with a wide variation in amino acid sequence and which have little or no homology to other PUL-encoded proteins or known carbohydrate-binding proteins. As the Sus SGBPs remain the only structurally characterized cohort to date, we therefore wondered whether such glycan binding and function are extended to other PUL that target more complex and heterogeneous polysaccharides, such as XyG. We describe here the detailed functional and structural characterization of the noncatalytic SGBPs encoded by Bacova_02651 and Bacova_02650 of the XyGUL, here referred to as SGBP-A and SGBP-B, to elucidate their molecular roles in carbohydrate acquisition in vivo. Combined biochemical, structural, and reverse-genetic approaches clearly illuminate the distinct, yet complementary, functions that these two proteins play in XyG recognition as it impacts the physiology of B. ovatus. These data extend our current understanding of the Sus-like glycan uptake paradigm within the Bacteroidetes and reveals how the complex dietary polysaccharide xyloglucan is recognized at the cell surface. RESULTS AND DISCUSSION SGBP-A and SGBP-B are cell-surface-localized, xyloglucan-specific binding proteins. SGBP-A, encoded by the XyGUL locus tag Bacova_02651 (Fig. 1B), shares 26% amino acid sequence identity (40% similarity) with its homolog, B. thetaiotaomicron SusD, and similar homology with the SusD-like proteins encoded within syntenic XyGUL identified in our earlier work. In contrast, SGBP-B, encoded by locus tag Bacova_02650, displays little sequence similarity to the products of similarly positioned genes in syntenic XyGUL nor to any other gene product among the diversity of Bacteroidetes PUL. Whereas sequence similarity among SusC/SusD homolog pairs often serves as a hallmark for PUL identification, the sequence similarities of downstream genes encoding SGBPs are generally too low to allow reliable bioinformatic classification of their products into protein families, let alone prediction of function. Hence, there is a critical need for the elucidation of detailed structure-function relationships among PUL SGBPs, in light of the manifold glycan structures in nature. Immunofluorescence of formaldehyde-fixed, nonpermeabilized cells grown in minimal medium with XyG as the sole carbon source to induce XyGUL expression, reveals that both SGBP-A and SGBP-B are presented on the cell surface by N-terminal lipidation, as predicted by signal peptide analysis with SignalP (Fig. 2). Here, the SGBPs very likely work in concert with the cell-surface-localized endo-xyloglucanase B. ovatus GH5 (BoGH5) to recruit and cleave XyG for subsequent periplasmic import via the SusC-like TBDT of the XyGUL (Fig. 1B and C). SGBP-A and SGBP-B visualized by immunofluorescence. Formalin-fixed, nonpermeabilized B. ovatus cells were grown in minimal medium plus XyG, probed with custom rabbit antibodies to SGBP-A or SGBP-B, and then stained with Alexa Fluor 488 goat anti-rabbit IgG. (A) Overlay of bright-field and FITC images of B. ovatus cells labeled with anti-SGBP-A. (B) Overlay of bright-field and FITC images of B. ovatus cells labeled with anti-SGBP-B. (C) Bright-field image of ΔSGBP-B cells labeled with anti-SGBP-B antibodies. (D) FITC images of ΔSGBP-B cells labeled with anti-SGBP-B antibodies. Cells lacking SGBP-A (ΔSGBP-A) do not grow on XyG and therefore could not be tested in parallel. In our initial study focused on the functional characterization of the glycoside hydrolases of the XyGUL, we reported preliminary affinity PAGE and isothermal titration calorimetry (ITC) data indicating that both SGBP-A and SGBP-B are competent xyloglucan-binding proteins (affinity constant [Ka] values of 3.74 × 105 M−1 and 4.98 × 104 M−1, respectively [23]). Additional affinity PAGE analysis (Fig. 3) demonstrates that SGBP-A also has moderate affinity for the artificial soluble cellulose derivative hydroxyethyl cellulose [HEC; a β(1 → 4)-glucan] and limited affinity for mixed-linkage β(1→3)/β(1→4)-glucan (MLG) and glucomannan (GM; mixed glucosyl and mannosyl backbone), which together indicate general binding to polysaccharide backbone residues and major contributions from side-chain recognition. In contrast, SGBP-B bound to HEC more weakly than SGBP-A and did not bind to MLG or GM. Neither SGBP recognized galactomannan (GGM), starch, carboxymethylcellulose, or mucin (see Fig. S1 in the supplemental material). Together, these results highlight the high specificities of SGBP-A and SGBP-B for XyG, which is concordant with their association with XyG-specific GHs in the XyGUL, as well as transcriptomic analysis indicating that B. ovatus has discrete PUL for MLG, GM, and GGM (11). Notably, the absence of carbohydrate-binding modules in the GHs encoded by the XyGUL implies that noncatalytic recognition of xyloglucan is mediated entirely by SGBP-A and -B. SGBP-A and SGBP-B preferentially bind xyloglucan. Affinity electrophoresis (10% acrylamide) of SGBP-A and SGBP-B with BSA as a control protein. All samples were loaded on the same gel next to the BSA controls; thin black lines indicate where intervening lanes were removed from the final image for both space and clarity. The percentage of polysaccharide incorporated into each native gel is displayed. The vanguard endo-xyloglucanase of the XyGUL, BoGH5, preferentially cleaves the polysaccharide at unbranched glucosyl residues to generate xylogluco-oligosaccharides (XyGOs) comprising a Glc4 backbone with variable side-chain galactosylation (XyGO1) (Fig. 1A; n = 1) as the limit of digestion products in vitro; controlled digestion and fractionation by size exclusion chromatography allow the production of higher-order oligosaccharides (e.g., XyGO2) (Fig. 1A; n = 2). ITC demonstrates that SGBP-A binds to XyG polysaccharide and XyGO2 (based on a Glc8 backbone) with essentially equal affinities, while no binding of XyGO1 (Glc4 backbone) was detectable (Table 1; see Fig. S2 and S3 in the supplemental material). Similarly, SGBP-B also bound to XyG and XyGO2 with approximately equal affinities, although in both cases, Ka values were nearly 10-fold lower than those for SGBP-A. Also in contrast to SGBP-A, SGBP-B also bound to XyGO1, yet the affinity for this minimal repeating unit was poor, with a Ka value of ca. 1 order of magnitude lower than for XyG and XyGO2. Together, these data clearly suggest that polysaccharide binding of both SGBPs is fulfilled by a dimer of the minimal repeat, corresponding to XyGO2 (cf. Fig. 1A). The observation by affinity PAGE that these proteins specifically recognize XyG is further substantiated by their lack of binding for the undecorated oligosaccharide cellotetraose (Table 1; see Fig. S3). Furthermore, SGBP-A binds cellohexaose with ~770-fold weaker affinity than XyG, while SGBP-B displays no detectable binding to this linear hexasaccharide. To provide molecular-level insight into how the XyGUL SGBPs equip B. ovatus to specifically harvest XyG from the gut environment, we performed X-ray crystallography analysis of both SGBP-A and SGPB-B in oligosaccharide-complex forms. Summary of thermodynamic parameters for wild-type SGBP-A and SGBP-B obtained by isothermal titration calorimetry at 25°Ca Carbohydrate\tKa (M−1)\tΔG (kcal ⋅ mol−1)\tΔH (kcal ⋅ mol−1)\tTΔS (kcal ⋅ mol−1)\t \tSGBP-A\tSGBP-B\tSGBP-A\tSGBP-B\tSGBP-A\tSGBP-B\tSGBP-A\tSGBP-B\t \tXyGb\t(4.4 ± 0.1) × 105\t(5.7 ± 0.2) × 104\t−7.7\t−6.5\t−14 ± 3\t−14 ± 2\t−6.5\t−7.6\t \tXyGO2c\t3.0 × 105\t2.0 × 104\t−7.5\t−5.9\t−17.2\t−17.6\t−9.7\t−11.7\t \tXyGO1\tNBd\t(2.4 ± 0.1) × 103\tNB\t−4.6\tNB\t−4.4 ± 0.2\tNB\t0.2\t \tCellohexaose\t568.0 ± 291.0\tNB\t−3.8\tNB\t−16 ± 8\tNB\t−12.7\tNB\t \tCellotetraose\tNB\tNB\tNB\tNB\tNB\tNB\tNB\tNB\t \t Shown are average values ± standard errors from two independent titrations, unless otherwise indicated. Binding thermodynamics for XyG based on the concentration of the binding unit, XyGO2. Values from a single titration. NB, no binding observed. SGBP-A is a SusD homolog with an extensive glycan-binding platform. As anticipated by sequence similarity, the high-resolution tertiary structure of apo-SGBP-A (1.36 Å, Rwork = 14.7%, Rfree = 17.4%, residues 28 to 546) (Table 2) displays the canonical “SusD-like” protein fold dominated by four tetratrico-peptide repeat (TPR) motifs that cradle the rest of the structure (Fig. 4A). Specifically, SGBP-A overlays B. thetaiotaomicron SusD (BtSusD) with a root mean square deviation (RMSD) value of 2.2 Å for 363 Cα pairs, which is notable given the 26% amino acid identity (40% similarity) between these homologs (Fig. 4C). Cocrystallization of SGBP-A with XyGO2 generated a substrate complex structure (2.3 Å, Rwork = 21.8%, Rfree = 24.8%, residues 36 to 546) (Fig. 4A and B; Table 2) that revealed the distinct binding-site architecture of the XyG binding protein. The SGBP-A:XyGO2 complex superimposes closely with the apo structure (RMSD of 0.6 Å) and demonstrates that no major conformational change occurs upon substrate binding; small deviations in the orientation of several surface loops are likely the result of differential crystal packing. It is particularly notable that although the location of the ligand-binding site is conserved between SGBP-A and SusD, that of SGBP-A displays an ~29-Å-long aromatic platform to accommodate the extended, linear XyG chain (see reference for a review of XyG secondary structure), versus the shorter, ~18-Å-long, site within SusD that complements the helical conformation of amylose (Fig. 4C and D). Molecular structure of SGBP-A (Bacova_02651). (A) Overlay of SGBP-A from the apo (rainbow) and XyGO2 (gray) structures. The apo structure is color ramped from blue to red. An omit map (2σ) for XyGO2 (orange and red sticks) is displayed. (B) Close-up view of the omit map as in panel A, rotated 90° clockwise. (C) Overlay of the Cα backbones of SGBP-A (black) with XyGO2 (orange and red spheres) and BtSusD (blue) with maltoheptaose (pink and red spheres), highlighting the conservation of the glycan-binding site location. (D) Close-up of the SGBP-A (black and orange) and SusD (blue and pink) glycan-binding sites. The approximate length of each glycan-binding site is displayed, colored to match the protein structures. (E) Stereo view of the xyloglucan-binding site of SGBP-A, displaying all residues within 4 Å of the ligand. The backbone glucose residues are numbered from the nonreducing end; xylose residues are labeled X1 and X2. Potential hydrogen-bonding interactions are shown as dashed lines, and the distance is shown in angstroms. X-ray data collection and refinement statistics Parameter\tValue(s) fora:\t \tSGBP-A apo\tSGBP-A/XyGO2\tSGBP-B/XyGO2\tSGBP-B (CD)/XyGO2\t \tPDB ID no.\t5E75\t5E76\t5E7G\t5E7H\t \tResolution (Å)\t21.48–1.36 (1.409–1.36)\t56.13–2.3 (2.382–2.3)\t39.19–2.37 (2.455–2.37)\t30.69–1.57 (1.626–1.570)\t \tSpace group\tP21\tI422\tR32\tP6122\t \tUnit cell dimensions, a, b, c (Å)\t52.8, 81.4, 57.7; β = 107.85°\t131.5, 131.5, 188\t207.4, 207.4, 117.9\t87.1, 87.1, 201.6\t \tNo. of reflections\t\t\t\t\t \t    Total\t355,272 (26,772)\t1,068,014 (102,923)\t324,544 (32,355)\t1,366,812 (129,645)\t \t    Unique\t99,136 (9,762)\t36,775 (3,625)\t39,362 (3,898)\t62,808 (6,068)\t \tMultiplicity\t3.6 (2.7)\t29.0 (28.4)\t8.2 (8.3)\t21.8 (21.4)\t \tCompleteness (%)\t99.71 (98.82)\t99.63 (99.42)\t99.96 (100.00)\t98.4 (96.98)\t \tMean I/σ〈I〉\t15.57 (2.29)\t24.93 (6.71)\t20.98 (2.36)\t38.52 (5.03)\t \tWilson B-factor\t11.91\t31.14\t43.91\t17.86\t \tRmerge\t0.04759 (0.4513)\t0.1428 (0.7178)\t0.09159 (1.197)\t0.05559 (0.7748)\t \tCC1/2b\t0.999 (0.759)\t0.999 (0.982)\t0.999 (0.794)\t1.000 (0.933)\t \tCC*c\t1.000 (0.929)\t1.000 (0.995)\t1.000 (0.941)\t1.000 (0.982)\t \tRwork\t0.1468 (0.2597)\t0.2178 (0.2788)\t0.1975 (0.3018)\t0.1560 (0.2008)\t \tRfree\t0.1738 (0.2632)\t0.2482 (0.2978)\t0.2260 (0.3219)\t0.1712 (0.2019)\t \tNo. of non-hydrogen atoms\t\t\t\t\t \t    All\t4,562\t4,319\t3,678\t2,328\t \t    Macromolecules\t4,079\t3,974\t3,425\t1,985\t \t    Ligands\t39\t116\t127\t25\t \t    Water\t444\t229\t126\t318\t \tNo. of protein residues\t506\t492\t446\t260\t \tRMSD\t\t\t\t\t \t    Bond length (Å)\t0.008\t0.007\t0.005\t0.009\t \t    Bond angle (°)\t1.15\t0.96\t0.87\t1.18\t \tRamachandran statistics\t\t\t\t\t \t    Favored (%)\t98\t95\t97\t98\t \t    Outliers (%)\t0\t0.41\t0.23\t0\t \t    Clash score\t0.5\t2.13\t0.86\t1.27\t \tAvg B-factors\t\t\t\t\t \t    All\t16.1\t53.2\t53\t25.4\t \t    Macromolecules\t15.2\t53.5\t52.5\t22.9\t \t    Ligands\t24.7\t61\t71.1\t47\t \t    Solvent\t24.4\t42.9\t47.6\t39\t \t Numbers in parentheses are for the highest-resolution shell. CC1/2, Pearson correlation coefficient between the average intensities of each subset. CC*, Pearson correlation coefficient for correlation between the observed data set and true signal. Seven of the eight backbone glucosyl residues of XyGO2 could be convincingly modeled in the ligand electron density, and only two α(1→6)-linked xylosyl residues were observed (Fig. 4B; cf. Fig. 1). Indeed, the electron density for the ligand suggests some disorder, which may arise from multiple oligosaccharide orientations along the binding site. Three aromatic residues—W82, W283, W306—comprise the flat platform that stacks along the naturally twisted β-glucan backbone (Fig. 4E). The functional importance of this platform is underscored by the observation that the W82A W283A W306A mutant of SGBP-A, designated SGBP-A*, is completely devoid of XyG affinity (Table 3; see Fig. S4 in the supplemental material). Dissection of the individual contribution of these residues reveals that the W82A mutant displays a significant 4.9-fold decrease in the Ka value for XyG, while the W306A substitution completely abolishes XyG binding. Contrasting with the clear importance of these hydrophobic interactions, there are remarkably few hydrogen-bonding interactions with the ligand, which are provided by R65, N83, and S308, which are proximal to Glc5 and Glc3. Most surprising in light of the saccharide-binding data, however, was a lack of extensive recognition of the XyG side chains; only Y84 appeared to provide a hydrophobic interface for a xylosyl residue (Xyl1). Summary of thermodynamic parameters for site-directed mutants of SGBP-A and SGBP-B obtained by ITC with XyG at 25°Ca Protein name\tKa\tΔG (kcal ⋅ mol−1)\tΔH (kcal ⋅ mol−1)\tTΔS (kcal ⋅ mol−1)\t \tFold changeb\tM−1\t \tSGBP-A(W82A W283A W306A)\tND\tNB\tNB\tNB\tNB\t \tSGBP-A(W82A)c\t4.9\t9.1 × 104\t−6.8\t−6.3\t0.5\t \tSGBP-A(W306)\tND\tNB\tNB\tNB\tNB\t \tSGBP-B(230–489)\t0.7\t(8.6 ± 0.20) × 104\t−6.7\t−14.9 ± 0.1\t−8.2\t \tSGBP-B(Y363A)\t19.7\t(2.9 ± 0.10) × 103\t−4.7\t−18.1 ± 0.1\t−13.3\t \tSGBP-B(W364A)\tND\tWeak\tWeak\tWeak\tWeak\t \tSGBP-B(F414A)\t3.2\t(1.80 ± 0.03) × 104\t−5.8\t−11.4 ± 0.1\t−5.6\t \t Shown are average values ± standard deviations from two independent titrations, unless otherwise indicated. Binding thermodynamics are based on the concentration of the binding unit, XyGO2. Weak binding represents a Ka of \u003c500 M−1. ND, not determined; NB, no binding. Ka fold change = Ka of wild-type protein/Ka of mutant protein for xyloglucan binding. Values from a single titration. SGBP-B has a multimodular structure with a single, C-terminal glycan-binding domain. The crystal structure of full-length SGBP-B in complex with XyGO2 (2.37 Å, Rwork = 19.9%, Rfree = 23.9%, residues 34 to 489) (Table 2) revealed an extended structure composed of three tandem immunoglobulin (Ig)-like domains (domains A, B, and C) followed at the C terminus by a novel xyloglucan-binding domain (domain D) (Fig. 5A). Domains A, B, and C display similar β-sandwich folds; domains B (residues 134 to 230) and C (residues 231 to 313) can be superimposed onto domain A (residues 34 to 133) with RMSDs of 1.1 and 1.2 Å, respectively, for 47 atom pairs (23% and 16% sequence identity, respectively). These domains also display similarity to the C-terminal β-sandwich domains of many GH13 enzymes, including the cyclodextrin glucanotransferase of Geobacillus stearothermophilus (Fig. 5B). Such domains are not typically involved in carbohydrate binding. Indeed, visual inspection of the SGBP-B structure, as well as individual production of the A and B domains and affinity PAGE analysis (see Fig. S5 in the supplemental material), indicates that these domains do not contribute to XyG capture. On the other hand, production of the fused domains C and D in tandem (SGBP-B residues 230 to 489) retains complete binding of xyloglucan in vitro, with the observed slight increase in affinity likely arising from a reduced potential for steric hindrance of the smaller protein construct during polysaccharide interactions (Table 3). While neither the full-length protein nor domain D displays structural homology to known XyG-binding proteins, the topology of SGBP-B resembles the xylan-binding protein Bacova_04391 (PDB 3ORJ) encoded within a xylan-targeting PUL of B. ovatus (Fig. 5C). The structure-based alignment of these proteins reveals 17% sequence identity, with a core RMSD of 3.6 Å for 253 aligned residues. While there is no substrate-complexed structure of Bacova_04391 available, the binding site is predicted to include W241 and Y404, which are proximal to the XyGO binding site in SGBP-B. However, the opposing, clamp-like arrangement of these residues in Bacova_04391 is clearly distinct from the planar surface arrangement of the residues that interact with XyG in SGBP-B (described below). Multimodular structure of SGBP-B (Bacova_02650). (A) Full-length structure of SGBP-B, color coded by domain as indicated. Prolines between domains are indicated as spheres. An omit map (2σ) for XyGO2 is displayed to highlight the location of the glycan-binding site. (B) Overlay of SGBP-B domains A, B, and C (colored as in panel A), with a C-terminal Ig-like domain of the G. stearothermophilus cyclodextrin glucanotransferase (PDB 1CYG [residues 375 to 493]) in green. (C) Cα overlay of SGBP-B (gray) and Bacova_04391 (PDB 3ORJ) (pink). (D) Close-up omit map for the XyGO2 ligand, contoured at 2σ. (E) Stereo view of the xyloglucan-binding site of SGBP-B, displaying all residues within 4 Å of the ligand. The backbone glucose residues are numbered from the nonreducing end, xylose residues are shown as X1, X2, and X3, potential hydrogen-bonding interactions are shown as dashed lines, and the distance is shown in angstroms. Inspection of the tertiary structure indicates that domains C and D are effectively inseparable, with a contact interface of 396 Å2. Domains A, B, and C do not pack against each other. Moreover, the five-residue linkers between these first three domains all feature a proline as the middle residue, suggesting significant conformational rigidity (Fig. 5A). Despite the lack of sequence and structural conservation, a similarly positioned proline joins the Ig-like domains of the xylan-binding Bacova_04391 and the starch-binding proteins SusE and SusF. We speculate that this is a biologically important adaptation that serves to project the glycan binding site of these proteins far from the membrane surface. Any mobility of SGBP-B on the surface of the cell (beyond lateral diffusion within the membrane) is likely imparted by the eight-residue linker that spans the predicted lipidated Cys (C28) and the first β-strand of domain A. Other outer membrane proteins from various Sus-like systems possess a similar 10- to 20-amino-acid flexible linker between the lipidated Cys that tethers the protein to the outside the cell and the first secondary structure element. Analogously, the outer membrane-anchored endo-xyloglucanase BoGH5 of the XyGUL contains a 100-amino-acid, all-β-strand, N-terminal module and flexible linker that imparts conformational flexibility and distances the catalytic module from the cell surface. XyG binds to domain D of SGBP-B at the concave interface of the top β-sheet, with binding mediated by loops connecting the β-strands. Six glucosyl residues, comprising the main chain, and three branching xylosyl residues of XyGO2 can be modeled in the density (Fig. 5D; cf. Fig. 1A). The backbone is flat, with less of the “twisted-ribbon” geometry observed in some cello- and xylogluco-oligosaccharides. The aromatic platform created by W330, W364, and Y363 spans four glucosyl residues, compared to the longer platform of SGBP-A, which supports six glucosyl residues (Fig. 5E). The Y363A site-directed mutant of SGBP-B displays a 20-fold decrease in the Ka for XyG, while the W364A mutant lacks XyG binding (Table 3; see Fig. S6 in the supplemental material). There are no additional contacts between the protein and the β-glucan backbone and surprisingly few interactions with the side-chain xylosyl residues, despite that fact that ITC data demonstrate that SGBP-B does not measurably bind the cellohexaose (Table 1). F414 stacks with the xylosyl residue of Glc3, while Q407 is positioned for hydrogen bonding with the O4 of xylosyl residue Xyl1. Surprisingly, an F414A mutant of SGBP-B displays only a mild 3-fold decrease in the Ka value for XyG, again suggesting that glycan recognition is primarily mediated via contact with the β-glucan backbone (Table 3; see Fig. S6). Additional residues surrounding the binding site, including Y369 and E412, may contribute to the recognition of more highly decorated XyG, but precisely how this is mediated is presently unclear. Hoping to achieve a higher-resolution view of the SGBP-B–xyloglucan interaction, we solved the crystal structure of the fused CD domains in complex with XyGO2 (1.57 Å, Rwork = 15.6%, Rfree = 17.1%, residues 230 to 489) (Table 2). The CD domains of the truncated and full-length proteins superimpose with a 0.4-Å RMSD of the Cα backbone, with no differences in the position of any of the glycan-binding residues (see Fig. S7A in the supplemental material). While density is observed for XyGO2, the ligand could not be unambiguously modeled into this density to achieve a reasonable fit between the X-ray data and the known stereochemistry of the sugar (see Fig. S7B and C). While this may occur for a number of reasons in crystal structures, it is likely that the poor ligand density even at higher resolution is due to movement or multiple orientations of the sugar averaged throughout the lattice. SGBP-A and SGBP-B have distinct, coordinated functions in vivo. The similarity of the glycan specificity of SGBP-A and SGBP-B presents an intriguing conundrum regarding their individual roles in XyG utilization by B. ovatus. To disentangle the functions of SGBP-A and SGBP-B in XyG recognition and uptake, we created individual in-frame deletion and complementation mutant strains of B. ovatus. In these growth experiments, overnight cultures of strains grown on minimal medium plus glucose were back-diluted 1:100-fold into minimal medium containing 5 mg/ml of the reported carbohydrate. Growth on glucose displayed the shortest lag time for each strain, and so lag times were normalized for each carbohydrate by subtracting the lag time of that strain in glucose (Fig. 6; see Fig. S8 in the supplemental material). A strain in which the entire XyGUL is deleted displays a lag of 24.5 h during growth on glucose compared to the isogenic parental wild-type (WT) Δtdk strain, for which exponential growth lags for 19.8 h (see Fig. S8D). It is unknown whether this is because cultures were not normalized by the starting optical density (OD) or viable cells or reflects a minor defect for glucose utilization. The former seems more likely as the growth rates are nearly identical for these strains on glucose and xylose. The ΔXyGUL and WT Δtdk strains display normalized lag times on xylose within experimental error, and curiously some of the mutant and complemented strains display a nominally shorter lag time on xylose than the WT Δtdk strain. Complementation of the ΔSGBP-A strain (ΔSGBP-A::SGBP-A) restores growth to wild-type rates on xyloglucan and XyGO1, yet the calculated rate of the complemented strain is ~72% that of the WT Δtdk strain on XyGO2; similar results were obtained for the SGBP-B complemented strain despite the fact that the growth curves do not appear much different (see Fig. S8C and F). The reason for this observation on XyGO2 is unclear, as the ΔSGBP-B mutant does not have a significantly different growth rate from the WT on XyGO2. Therefore, we limit our discussion to those mutants that displayed the most obvious defects in growth on particular substrates. Growth of select XyGUL mutants on xyloglucan and oligosaccharides. B. ovatus mutants were created in a thymidine kinase deletion (Δtdk) mutant as described previously. SGBP-A* denotes the Bacova_02651 (W82A W283A W306A) allele, and the GH9 gene is Bacova_02649. Growth was measured over time in minimal medium containing (A) XyG, (B) XyGO2, (C) XyGO1, (D) glucose, and (E) xylose. In panel F, the growth rate of each strain on the five carbon sources is displayed, and in panel G, the normalized lag time of each culture, relative to its growth on glucose, is displayed. Solid bars indicate conditions that are not statistically significant from the WT Δtdk cultures grown on the indicated carbohydrate, while open bars indicate a P value of \u003c0.005 compared to the WT Δtdk strain. Conditions denoted by the same letter (b, c, or d) are not statistically significant from each other but are significantly different from the condition labeled “a.” Complementation of ΔSGBP-A and ΔSBGP-B was performed by allelic exchange of the wild-type genes back into the genome for expression via the native promoter: these growth curves, quantified rates and lag times are displayed in Fig. S8 in the supplemental material. The ΔSGBP-A (ΔBacova_02651) strain (cf. Fig. 1B) was completely incapable of growth on XyG, XyGO1, and XyGO2, indicating that SGBP-A is essential for XyG utilization (Fig. 6). This result mirrors our previous data for the canonical Sus of B. thetaiotaomicron, which revealed that a homologous ΔsusD mutant is unable to grow on starch or malto-oligosaccharides, despite normal cell surface expression of all other PUL-encoded proteins. More recently, we demonstrated that this phenotype is due to the loss of the physical presence of SusD; complementation of ΔsusD with SusD*, a triple site-directed mutant (W96A W320A Y296A) that ablates glycan binding, restores B. thetaiotaomicron growth on malto-oligosaccharides and starch when sus transcription is induced by maltose addition. Similarly, the function of SGBP-A extends beyond glycan binding. Complementation of ΔSGBP-A with the SGBP-A* (W82A W283A W306A) variant, which does not bind XyG, supports growth on XyG and XyGOs (Fig. 6; ΔSGBP-A::SGBP-A*), with growth rates that are ~70% that of the WT. In previous studies, we observed that carbohydrate binding by SusD enhanced the sensitivity of the cells to limiting concentrations of malto-oligosaccharides by several orders of magnitude, such that the addition of 0.5 g/liter maltose was required to restore growth of the ΔsusD::SusD* strain on starch, which nonetheless occurred following an extended lag phase. In contrast, the ΔSGBP-A::SGBP-A* strain does not display an extended lag time on any of the xyloglucan substrates compared to the WT (Fig. 6). The specific glycan signal that upregulates BoXyGUL is currently unknown. From our present data, we cannot eliminate the possibility that the glycan binding by SGBP-A enhances transcriptional activation of the XyGUL. However, the modest rate defect displayed by the SGBP-A::SGBP-A* strain suggests that recognition of XyG and product import is somewhat less efficient in these cells. Intriguingly, the ΔSGBP-B strain (ΔBacova_02650) (cf. Fig. 1B) exhibited a minor growth defect on both XyG and XyGO2, with rates 84.6% and 93.9% that of the WT Δtdk strain. However, growth of the ΔSGBP-B strain on XyGO1 was 54.2% the rate of the parental strain, despite the fact that SGBP-B binds this substrate ca. 10-fold more weakly than XyGO2 and XyG (Fig. 6; Table 1). As such, the data suggest that SGBP-A can compensate for the loss of function of SGBP-B on longer oligo- and polysaccharides, while SGBP-B may adapt the cell to recognize smaller oligosaccharides efficiently. Indeed, a double mutant, consisting of a crippled SGBP-A and a deletion of SGBP-B (ΔSGBP-A::SGBP-A*/ΔSGBP-B), exhibits an extended lag time on both XyG and XyGO2, as well as XyGO1. Taken together, the data indicate that SGBP-A and SGBP-B functionally complement each other in the capture of XyG polysaccharide, while SGBP-B may allow B. ovatus to scavenge smaller XyGOs liberated by other gut commensals. This additional role of SGBP-B is especially notable in the context of studies on BtSusE and BtSusF (positioned similarly in the archetypal Sus locus) (Fig. 1B), for which growth defects on starch or malto-oligosaccharides have never been observed. Beyond SGBP-A and SGBP-B, we speculated that the catalytically feeble endo-xyloglucanase GH9, which is expendable for growth in the presence of GH5, might also play a role in glycan binding to the cell surface. However, combined deletion of the genes encoding GH9 (encoded by Bacova_02649) and SGBP-B does not exacerbate the growth defect on XyGO1 (Fig. 6; ΔSGBP-B/ΔGH9). The necessity of SGBP-B is elevated in the SGBP-A* strain, as the ΔSGBP-A::SGBP-A*/ ΔSGBP-B mutant displays an extended lag during growth on XyG and xylogluco-oligosaccharides, while growth rate differences are more subtle. The precise reason for this lag is unclear, but recapitulating our findings on the role of SusD in malto-oligosaccharide sensing in B. thetaiotaomicron, this extended lag may be due to inefficient import and thus sensing of xyloglucan in the environment in the absence of glycan binding by essential SGBPs. Our previous work demonstrates that B. ovatus cells grown in minimal medium plus glucose express low levels of the XyGUL transcript. Thus, in our experiments, we presume that each strain, initially grown in glucose, expresses low levels of the XyGUL transcript and thus low levels of the XyGUL-encoded surface proteins, including the vanguard GH5. Presumably without glycan binding by the SGBPs, the GH5 protein cannot efficiently process xyloglucan, and/or the lack of SGBP function prevents efficient capture and import of the processed oligosaccharides. It may then be that only after a sufficient amount of glycan is processed and imported by the cell is XyGUL upregulated and exponential growth on the glycan can begin. We hypothesize that during exponential growth the essential role of SGBP-A extends beyond glycan recognition, perhaps due to a critical interaction with the TBDT. In the BtSus, SusD and the TBDT SusC interact, and we speculate that this interaction is necessary for glycan uptake, as suggested by the fact that a ΔsusD mutant cannot grow on starch, but a ΔsusD::SusD* strain regains this ability if a transcriptional activator of the sus operon is supplied. Likewise, such cognate interactions between homologous protein pairs such as SGBP-A and its TBDT may underlie our observation that a ΔSGBP-A mutant cannot grow on xyloglucan. However, unlike the Sus, in which elimination of SusE and SusF does not affect growth on starch, SGBP-B appears to have a dedicated role in growth on small xylogluco-oligosaccharides. Conclusions. The ability of gut-adapted microorganisms to thrive in the gastrointestinal tract is critically dependent upon their ability to efficiently recognize, cleave, and import glycans. The human gut, in particular, is a densely packed ecosystem with hundreds of species, in which there is potential for both competition and synergy in the utilization of different substrates. Recent work has elucidated that Bacteroidetes cross-feed during growth on many glycans; the glycoside hydrolases expressed by one species liberate oligosaccharides for consumption by other members of the community. Thus, understanding glycan capture at the cell surface is fundamental to explaining, and eventually predicting, how the carbohydrate content of the diet shapes the gut community structure as well as its causative health effects. Here, we demonstrate that the surface glycan binding proteins encoded within the BoXyGUL play unique and essential roles in the acquisition of the ubiquitous and abundant vegetable polysaccharide xyloglucan. Yet, a number of questions remain regarding the molecular interplay of SGBPs with their cotranscribed cohort of glycoside hydrolases and TonB-dependent transporters. A particularly understudied aspect of glycan utilization is the mechanism of import via TBDTs (SusC homologs) (Fig. 1), which are ubiquitous and defining components of all PUL. PUL-encoded TBDTs in Bacteroidetes are larger than the well-characterized iron-targeting TBDTs from many Proteobacteria and are further distinguished as the only known glycan-importing TBDTs coexpressed with an SGBP. A direct interaction between the BtSusC TBDT and the SusD SGBP has been previously demonstrated, as has an interaction between the homologous components encoded by an N-glycan-scavenging PUL of Capnocytophaga canimorsus. Our observation here that the physical presence of the SusD homolog SGBP-A, independent of XyG-binding ability, is both necessary and sufficient for XyG utilization further supports a model of glycan import whereby the SusC-like TBDTs and the SusD-like SGBPs must be intimately associated to support glycan uptake (Fig. 1C). It is yet presently unclear whether this interaction is static or dynamic and to what extent the association of cognate TBDT/SGBPs is dependent upon the structure of the carbohydrate to be imported. On the other hand, there is clear evidence for independent TBDTs in Bacteroidetes that do not require SGBP association for activity. For example, it was recently demonstrated that expression of nanO, which encodes a SusC-like TBDT as part of a sialic-acid-targeting PUL from B. fragilis, restored growth on this monosaccharide in a mutant strain of E. coli. In this instance, coexpression of the susD-like gene nanU was not required, nor did the expression of the nanU gene enhance growth kinetics. Similarly, the deletion of BT1762 encoding a fructan-targeting SusD-like protein in B. thetaiotaomicron did not result in a dramatic loss of growth on fructans. Thus, the strict dependence on a SusD-like SGBP for glycan uptake in the Bacteroidetes may be variable and substrate dependent. Furthermore, considering the broader distribution of TBDTs in PUL lacking SGBPs (sometimes known as carbohydrate utilization containing TBDT [CUT] loci; see reference and reviewed in reference) across bacterial phyla, it appears that the intimate biophysical association of these substrate-transport and -binding proteins is the result of specific evolution within the Bacteroidetes. Equally intriguing is the observation that while SusD-like proteins such as SGBP-A share moderate primary and high tertiary structural conservation, the genes for the SGBPs encoded immediately downstream (Fig. 1B [sometimes referred to as “susE positioned”]) encode glycan-binding lipoproteins with little or no sequence or structural conservation, even among syntenic PUL that target the same polysaccharide. Such is the case for XyGUL from related Bacteroides species, which may encode either one or two of these predicted SGBPs, and these proteins vary considerably in length. The extremely low similarity of these SGBPs is striking in light of the moderate sequence conservation observed among homologous GHs in syntenic PUL. This, together with the observation that these SGBPs, as exemplified by BtSusE and BtSusF and the XyGUL SGBP-B of the present study, are expendable for polysaccharide growth, implies a high degree of evolutionary flexibility to enhance glycan capture at the cell surface. Because the intestinal ecosystem is a dense consortium of bacteria that must compete for their nutrients, these multimodular SGBPs may reflect ongoing evolutionary experiments to enhance glycan uptake efficiency. Whether organisms that express longer SGBPs, extending further above the cell surface toward the extracellular environment, are better equipped to compete for available carbohydrates is presently unknown. However, the natural diversity of these proteins represents a rich source for the discovery of unique carbohydrate-binding motifs to both inform gut microbiology and generate new, specific carbohydrate analytical reagents. In conclusion, the present study further illuminates the essential role that surface-glycan binding proteins play in facilitating the catabolism of complex dietary carbohydrates by Bacteroidetes. The ability of our resident gut bacteria to recognize polysaccharides is the first committed step of glycan consumption by these organisms, a critical process that influences the community structure and thus the metabolic output (i.e., short-chain fatty acid and metabolite profile) of these organisms. A molecular understanding of glycan uptake by human gut bacteria is therefore central to the development of strategies to improve human health through manipulation of the microbiota. MATERIALS AND METHODS Protein production and purification. The gene fragments corresponding to Bacova_02650 (encoding SGBP-B residues 34 to 489) and Bacova_02651 (encoding SGBP-A residues 28 to 546) were amplified from Bacteroidetes ovatus ATCC 8483 genomic DNA by PCR using forward primers, including NdeI restriction sites, and reverse primers, including XhoI. The gene products were ligated into a modified version of pET-28a (EMD Biosciences) containing a recombinant tobacco etch virus (rTEV) protease recognition site (pET-28aTEV) preceding an N-terminal 6-His tag for affinity purification. The expression vector (pET-28TEV) containing SGBP-B was used for subsequent cloning of the domains A (residues 34 to 133), B (residues 134 to 229), and CD (residues 230 to 489). The pET-28TEV vector expressing residues 28 to 546 of SGBP-A was utilized for carbohydrate-binding experiments and crystallization of the apo structure. To obtain crystals of SGBP-A with XyGO2, the DNA sequence coding for residues 36 to 546 was PCR amplified from genomic DNA for ligation-independent cloning into the pETite N-His vector (Lucigen, Madison, WI) according to the manufacturer’s instructions. The N-terminal primer for pETite N-His insertion contained a TEV cleavage site immediately downstream of the complementary 18-bp overlap (encoding the His tag) to create a TEV-cleavable His-tagged protein. The site-directed mutants of SGBP-A and SGBP-B in pET-28TEV were created using the QuikChange II site-directed mutagenesis kit (Stratagene) according to the manufacturer’s instructions. The sequences of all primers to generate these constructs are displayed in Table S1 in the supplemental material. The plasmids containing the SGBP-A and SGBP-B genes were transformed into Escherichia coli BL21(DE3) or Rosetta(DE3) cells. For native protein expression, cells were cultured in Terrific Broth containing kanamycin (50 µg/ml) and chloramphenicol (20 µg/ml) at 37°C to the mid-exponential phase (A600 of ≈0.6), induced by the addition of 0.5 mM isopropyl β-d-1-thiogalactopyranoside (IPTG), and then incubated for 2 days at 16°C or 1 day at 20°C. Cells were harvested by centrifugation and frozen at −80°C prior to protein purification. For selenomethionine-substituted SGBP-B, the pET-28TEV-SGBP-B plasmid was transformed into E. coli Rosetta(DE3)/pLysS and plated onto LB supplemented with kanamycin (50 µg/ml) and chloramphenicol (20 µg/ml). After 16 h of growth at 37°C, colonies were harvested from the plates, used to inoculate 100 ml of M9 minimal medium supplemented with kanamycin (30 µg/ml) and chloramphenicol (20 µg/ml), and then grown at 37°C for 16 h. This overnight culture was used to inoculate a 2-liter baffled flask containing 1 liter of Molecular Dimensions SelenoMet premade medium supplemented with 50 ml of the recommended sterile nutrient mix, chloramphenicol, and kanamycin. Cultures were grown at 37°C to an A600 of ≈0.45 before adjusting the temperature to 20°C and supplementing each flask with 100 mg each of l-lysine, l-threonine, and l-phenylalanine and 50 mg each of l-leucine, l-isoleucine, l-valine, and l-selenomethionine. After 20 additional minutes of growth, the cells were induced with 0.5 mM IPTG, and cultures were grown for an additional 48 h. For the purification of native and selenomethionine-substituted protein, cells were thawed and lysed via sonication in His buffer (25 mM NaH2PO4, 500 mM NaCl, 20 mM imidazole, pH 7.5) and purified via immobilized nickel affinity chromatography (His-Trap; GE Healthcare) using a gradient of 20 to 300 mM imidazole, according to the manufacturer’s instructions. The His tag was removed by incubation with TEV protease (1:100 molar ratio relative to protein) at room temperature for 2 h and then overnight at 4°C while being dialyzed against His buffer. The cleaved protein was then repurified via nickel affinity chromatography to remove undigested target protein, the cleaved His tag, and His-tagged TEV protease. Purified proteins were dialyzed against 20 mM HEPES–100 mM NaCl (pH 7.0) prior to crystallization and concentrated using Vivaspin 15 (10,000-molecular-weight-cutoff [MWCO]) centrifugal concentrators (Vivaproducts, Inc.). Glycans. Xyloglucan from tamarind seed, β-glucan from barley, and konjac glucomannan were purchased from Megazyme. Starch, guar, and mucin were purchased from Sigma. Hydroxyethyl cellulose was purchased from AMRESCO. Carboxymethyl cellulose was purchased from Acros Organics. Xylogluco-oligosaccharides XyGO1 and XyGO2 for biophysical analyses were prepared from tamarind seed XyG according to the method of Martinez-Fleites et al. with minor modifications. XyGO2 for cocrystallization was purchased from Megazyme (O-XGHDP). Affinity gel electrophoresis. Affinity PAGE was performed as described previously, with minor modification. Various polysaccharides were used at a concentration of 0.05 to 0.1% (wt/vol), and electrophoresis was carried out for 90 min at room temperature in native 10% (wt/vol) polyacrylamide gels. BSA was used as noninteracting negative-control protein. ITC. Isothermal titration calorimetry (ITC) of glycan binding by the SGPB-A mutants was performed using the TA Nano isothermal titration calorimeter calibrated to 25°C. Proteins were dialyzed against 20 mM HEPES–100 mM NaCl (pH 7.0), and sugars were prepared using the dialysis buffer. The protein (45 to 50 µM) was placed in the sample cell, and the syringe was loaded with 2.5 to 4 mg/ml XyG polysaccharide. Following an initial injection of 0.5 µl, 26 subsequent injections of 2 µl were performed with stirring at 350 rpm, and the resulting heat of reaction was recorded. Data were analyzed using the Nano Analyze software. All other ITC experiments were performed using a MicroCal VP-ITC titration calorimeter calibrated to 25°C. Proteins were dialyzed into 20 mM HEPES–100 mM NaCl (pH 7.0), and polysaccharides were prepared using the dialysis buffer. Proteins (micromolar concentrations) were placed in the sample cell, and a first injection of 2 µl was performed followed by 24 subsequent injections of 10 µl of 2 to 20 mM oligosaccharide (cellotetraose, cellohexaose, XyGO1, or XyGO2) or 1 to 2.5 mg/ml XyG polysaccharide. The solution was stirred at 280 rpm, and the resulting heat of reaction was recorded. Data were analyzed using the Origin software program. DSC. Structural integrity of the SGBP-B mutants was verified by differential scanning calorimetry (DSC). DSC studies were performed on a MicroCal VP-DSC (Malvern Instruments). Experiments were carried out in 50 mM HEPES (pH 7.0) at a scan rate of 60°C/h. All samples (40 µM protein) were degassed for 7 min with gentle stirring under vacuum prior to being loaded into the calorimeter. Background excess thermal power scans were obtained with buffer in both the sample and reference cells and subtracted from the scans for each sample solution to generate excess heat capacity versus temperature thermograms. The melting temperature decreased from 57.8 ± 0.9°C for the wild-type SGBP-B protein to 54.6 ± 0.1°C for the Y363A mutant, 54.2 ± 0.1°C for the W364A mutant, and 52 ± 1°C for the F414A mutant. All proteins were therefore in their stable folded state for the ITC measurements (see Fig. S9 in the supplemental material). Bacteroides ovatus mutagenesis. Gene deletions and complementations were performed via allelic exchange in a Bacteroides ovatus thymidine kinase gene (Bacova_03071) deletion (Δtdk) derivative strain of ATCC 8483 using the vector pExchange-tdk, as previously described. Primers for the construction of B. ovatus mutants are listed in Table S1. The B. ovatus Δtdk strain and the B. ovatus ΔXyGUL mutant were a generous gift from Eric Martens, University of Michigan Medical School. Bacteroides growth experiments. All Bacteroides ovatus culturing was performed in a Coy anaerobic chamber (85% N2, 10% H2, 5% CO2) at 37°C. Prior to growth on minimal medium plus the carbohydrates indicated (Fig. 6; see Fig. S8 in the supplemental material), each strain was grown for 16 h from a freezer stock in tryptone-yeast extract-glucose (TYG) medium and then back diluted 1:100 into Bacteroides minimal medium supplemented with 5 mg/ml glucose, as previously described. After growth for 24 h, cultures were back-diluted 1:100 into Bacteroides minimal medium supplemented with 5 mg/ml of glucose, xylose, XyG, XyGO1, or XyGO2. Growth experiments were performed in replicates of 12 (glucose, xylose, and xyloglucan) or 5 (XyGO1 and XyGO2) as 200-µl cultures in 96-well plates. Plates were loaded into a Biostack automated plate handling device coupled to a Powerwave HT absorbance reader (both devices from Biotek Instruments, Winooski, VT). Absorbance at 600 nm (A600; i.e., optical density at 600 nm [OD600]) was measured for each well at 20-min intervals. Data were processed using Gen5 software (BioTek) and Microsoft Excel. Growth was quantified in each assay by first identifying a minimum time point (Amin) at which A600 had increased by 15% over a baseline reading taken during the first 500 min of incubation. Next, we identified the time point at which A600 reached its maximum (Amax) immediately after exponential growth. The growth rate for each well was defined by (Amax − Amin)/(Tmax − Tmin), where Tmax and Tmin are the corresponding time values for each absorbance. To account for variations in inoculum density, for each strain, the lag time (Tmin) on glucose was subtracted from the lag time for the substrate of interest; in all cases, cultures had shorter lag times on glucose than other glycans. Immunofluorescence. Custom rabbit antibodies to recombinant SGBP-A and SGBP-B were generated by Cocalico Biologicals, Inc. (Reamstown, PA). The B. ovatus ATCC 8483 Δtdk and ΔSGBP-B strains were grown in 1 ml minimal Bacteroides medium supplemented with 5 mg/ml tamarind xyloglucan to an A600 of ≈0.6 and then harvested via centrifugation (7,000 × g for 3 min) and washed twice with phosphate-buffered saline (PBS). Cells were resuspended in 0.25 ml PBS, and 0.75 ml of 6% formalin in PBS was added. Cells were incubated with rocking at 20°C for 1.5 h and then washed twice with PBS. Cells were resuspended in 0.5 to 1 ml blocking solution (2% goat serum, 0.02% NaN3 in PBS) and incubated for 16 h at 4°C. Cells were centrifuged and resuspended in 0.5 ml of a 1/100 dilution of custom rabbit antibody sera in blocking solution and incubated by rocking for 2 h at 20°C. Cells were washed with PBS and then resuspended in 0.4 ml of a 1/500 dilution of Alexa Fluor 488 goat anti-rabbit IgG (Life Technologies) in blocking solution and incubated with rocking for 1 h at 20°C. Cells were washed three times with an excess of PBS and then resuspended in 20 µl of PBS plus 1 µl of ProLong Gold antifade (Life Technologies). Cells were spotted on 0.8% agarose pads and imaged at the Center for Live Cell Imaging at the University of Michigan Medical School, using an Olympus IX70 inverted confocal microscope. Images were processed with Metamorph Software. Crystallization and data collection. All X-ray diffraction data for both native and selenomethionine-substituted protein crystals were collected at the Life Science Consortium (LS-CAT) at the Advance Photon Source at Argonne National Laboratory in Argonne, IL. The native protein SGBP-B (residues 34 to 489) was concentrated to an A280 of 12.25 prior to crystallization and mixed with 10 mM XyGO2 (Megazyme, O-XGHDP). Hanging drop vapor diffusion was performed against mother liquor consisting of 1.1 to 1.5 M ammonium sulfate and 30 to 70 mM sodium cacodylate (pH 6.5). To decrease crystal nucleation, 0.3 ml of paraffin oil was overlaid on top of 0.5 ml of mother liquor yielding diffraction-quality crystals within 2 weeks. Selenomethionine-substituted crystals of SGBP-B were generated using the same conditions as the native crystals. Crystals of the truncated SGBP-B (domains CD, residues 230 to 489) were obtained by mixing concentrated protein (A600 of 20.6) with 10 mM XyGO2 for hanging drop vapor diffusion against a solution containing 2 M sodium formate and 0.1 M sodium acetate (pH 4.6). All SGBP-B crystals were flash-frozen prior to data collection by briefly soaking in a solution of 80% mother liquor–20% glycerol plus 10 mM xylogluco-oligosaccharide. Data were processed and scaled using HKL2000 and Scalepack. SAD phasing from a selenomethionine-substituted protein crystals was used to determine the structure of SGBP-B. The AutoSol and Autobuild algorithms within the Phenix suite of programs were used to locate and refine the selenium positions and automatically build an initial model of the protein structure, respectively. Successive rounds of manual model building and refinement in Coot and Phenix, respectively, were utilized to build a 2.7-Å model of the selenomethionine-substituted protein, which then was placed in the unit cell of the native data set. Additional rounds of manual model building and refinement were performed to complete the 2.37-Å structure of SGBP-B with XyGO2. The structure of the truncated protein (CD domains, residues 230 to 489) was solved via molecular replacement with Phaser using the CD domains of the full-length protein as a model. The native protein SGBP-A (residues 28 to 546) was concentrated to an A280 of 28.6 and crystallized via hanging drop vapor diffusion from the Morpheus crystal screen (Molecular Dimensions). Crystals formed in well A1 (30 mM MgCl2, 30 mM CaCl2, 20% polyethylene glycol [PEG 500], 10% PEG 20K, 0.1 M imidazole-MES [morpholinethanesulfonic acid], pH 6.5), and were flash-frozen in liquid nitrogen without additional cryoprotectant. The truncated SGBP-A (residues 36 to 546) concentrated to an A280 of 38.2 yielded crystals with 10 mM XyGO2 via hanging drop vapor diffusion against 1.2 to 1.8 M sodium citrate (pH 6.15 to 6.25), and were flash-frozen in a cryoprotectant solution of 80% mother liquor–20% ethylene glycol with the glycan. Data were processed and scaled using HKL2000 and Scalepack. The structure of the apo protein was solved via molecular replacement with BALBES using the homologous structure PDB 3JYS, followed by successive rounds of automatic and manual model building with Autobuild and Coot. The structure of SGBP-A with XyGO2 was solved via molecular replacement with Phaser and refined with Phenix. X-data collection and refinement statistics are presented in Table 2. SUPPLEMENTAL MATERIAL Citation Tauzin AS, Kwiatkowski KJ, Orlovsky NI, Smith CJ, Creagh AL, Haynes CA, Wawrzak Z, Brumer H, Koropatkin NM. 2016. Molecular dissection of xyloglucan recognition in a prominent human gut symbiont. mBio 7(2):e02134-15. doi:10.1128/mBio.02134-15. 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+[{"sourceid":"4850288","sourcedb":"","project":"","target":"","text":"Crystal Structure and Activity Studies of the C11 Cysteine Peptidase from Parabacteroides merdae in the Human Gut Microbiome* Clan CD cysteine peptidases, a structurally related group of peptidases that include mammalian caspases, exhibit a wide range of important functions, along with a variety of specificities and activation mechanisms. However, for the clostripain family (denoted C11), little is currently known. Here, we describe the first crystal structure of a C11 protein from the human gut bacterium, Parabacteroides merdae (PmC11), determined to 1.7-Å resolution. PmC11 is a monomeric cysteine peptidase that comprises an extended caspase-like α/β/α sandwich and an unusual C-terminal domain. It shares core structural elements with clan CD cysteine peptidases but otherwise structurally differs from the other families in the clan. These studies also revealed a well ordered break in the polypeptide chain at Lys147, resulting in a large conformational rearrangement close to the active site. Biochemical and kinetic analysis revealed Lys147 to be an intramolecular processing site at which cleavage is required for full activation of the enzyme, suggesting an autoinhibitory mechanism for self-preservation. PmC11 has an acidic binding pocket and a preference for basic substrates, and accepts substrates with Arg and Lys in P1 and does not require Ca2+ for activity. Collectively, these data provide insights into the mechanism and activity of PmC11 and a detailed framework for studies on C11 peptidases from other phylogenetic kingdoms. Introduction Cysteine peptidases play crucial roles in the virulence of bacterial and other eukaryotic pathogens. In the MEROPS peptidase database, clan CD contains groups (or families) of cysteine peptidases that share some highly conserved structural elements. Clan CD families are typically described using the name of their archetypal, or founding, member and also given an identification number preceded by a “C,” to denote cysteine peptidase. Although seven families (C14 is additionally split into three subfamilies) have been described for this clan, crystal structures have only been determined from four: legumain (C13), caspase (C14a), paracaspase (C14b(P), metacaspase (C14b(M), gingipain (C25), and the cysteine peptidase domain (CPD) of various toxins (C80). No structural information is available for clostripain (C11), separase (C50), or PrtH-peptidase (C85). Clan CD enzymes have a highly conserved His/Cys catalytic dyad and exhibit strict specificity for the P1 residue of their substrates. However, despite these similarities, clan CD forms a functionally diverse group of enzymes: the overall structural diversity between (and at times within) the various families provides these peptidases with a wide variety of substrate specificities and activation mechanisms. Several members are initially expressed as proenzymes, demonstrating self-inhibition prior to full activation. The archetypal and arguably most notable family in the clan is that of the mammalian caspases (C14a), although clan CD members are distributed throughout the entire phylogenetic kingdom and are often required in fundamental biological processes. Interestingly, little is known about the structure or function of the C11 proteins, despite their widespread distribution and its archetypal member, clostripain from Clostridium histolyticum, first reported in the literature in 1938. Clostripain has been described as an arginine-specific peptidase with a requirement for Ca2+ and loss of an internal nonapeptide for full activation; lack of structural information on the family appears to have prohibited further investigation. As part of an ongoing project to characterize commensal bacteria in the microbiome that inhabit the human gut, the structure of C11 peptidase, PmC11, from Parabacteroides merdae was determined using the Joint Center for Structural Genomics (JCSG)4 HTP structural biology pipeline. The structure was analyzed, and the enzyme was biochemically characterized to provide the first structure/function correlation for a C11 peptidase. Experimental Procedures Cloning, expression, purification, crystallization, and structure determination of PmC11 were carried out using standard JCSG protocols as follows. Cloning Clones were generated using the polymerase incomplete primer extension (PIPE) cloning method. The gene encoding PmC11 (SP5111E) was amplified by polymerase chain reaction (PCR) from P. merdae genomic DNA using PfuTurbo DNA polymerase (Stratagene), using I-PIPE primers that included sequences for the predicted 5′ and 3′ ends (shown below). The expression vector, pSpeedET, which encodes an amino-terminal tobacco etch virus protease-cleavable expression and purification tag (MGSDKIHHHHHHENLYFQ/G), was PCR amplified with V-PIPE (Vector) primers. V-PIPE and I-PIPE PCR products were mixed to anneal the amplified DNA fragments together. Escherichia coli GeneHogs (Invitrogen) competent cells were transformed with the I-PIPE/V-PIPE mixture and dispensed on selective LB-agar plates. The cloning junctions were confirmed by DNA sequencing. The plasmid encoding the full-length protein was deposited in the PSI:Biology Materials Repository at the DNASU plasmid repository (PmCD00547516). For structure determination, to obtain soluble protein using the PIPE, method the gene segment encoding residues Met1-Asn22 was deleted because these residues were predicted to correspond to a signal peptide using SignalP. Protein Expression and Selenomethionine Incorporation The expression plasmid for the truncated PmC11 construct was transformed into E. coli GeneHogs competent cells and grown in minimal media supplemented with selenomethionine and 30 μg ml−1 of kanamycin at 37 °C using a GNF fermentor. A methionine auxotrophic strain was not required as selenomethionine is incorporated via the inhibition of methionine biosynthesis. Protein expression was induced using 0.1% (w/v) l-arabinose and the cells were left to grow for a further 3 h at 37 °C. At the end of the cell culture, lysozyme was added to all samples to a final concentration of 250 μg ml−1 and the cells were harvested and stored at −20 °C, until required. Primers used in this section are as follows: I-PIPE (forward): CTGTACTTCCAGGGCGAGACTCCGGAACCCCGGACAACCCGC; I-PIPE (reverse): AATTAAGTCGCGTTATTCATAAACTGCCTTATACCAGCCGAC; V-PIPE (forward): TAACGCGACTTAATTAACTCGTTTAAACGGTCTCCAGC; and V-PIPE (reverse): GCCCTGGAAGTACAGGTTTTCGTGATGATGATGATGAT. Protein Purification for Crystallization Cells were resuspended, homogenized, and lysed by sonication in 40 mm Tris (pH 8.0), 300 mm NaCl, 10 mm imidazole, and 1 mm Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (Lysis Buffer 1) containing 0.4 mm MgSO4 and 1 μl of 250 unit/μl−1 of benzonase (Sigma). The cell lysate was then clarified by centrifugation (32,500 × g for 25 min at 4 °C) before being passed over Ni2+-chelating resin equilibrated in Lysis Buffer 1 and washed in the same buffer supplemented with 40 mm imidazole and 10% (v/v) glycerol. The protein was subsequently eluted in 20 mm Tris (pH 8.0), 150 mm NaCl, 10% (v/v) glycerol, 1 mm TCEP, and 300 mm imidazole, and the fractions containing the protein were pooled. To remove the His tag, PmC11 was exchanged into 20 mm Tris (pH 8.0), 150 mm NaCl, 30 mm imidazole, and 1 mm TCEP using a PD-10 column (GE Healthcare), followed by incubation with 1 mg of His-tagged tobacco etch virus protease per 15 mg of protein for 2 h at room temperature and subsequent overnight incubation at 4 °C. The sample was centrifuged to remove any precipitated material (13,000 × g for 10 min at 4 °C) and the supernatant loaded onto Ni2+-chelating resin equilibrated with 20 mm Tris (pH 8.0), 150 mm NaCl, 30 mm imidazole, and 1 mm TCEP and washed with the same buffer. The flow-through and wash fractions were collected and concentrated to 13.3 mg ml−1 using Amicon Ultra-15 5K centrifugal concentrators (Millipore). Crystallization and Data Collection PmC11 was crystallized using the nanodroplet vapor diffusion method using standard JCSG crystallization protocols. Drops were comprised of 200 nl of protein solution mixed with 200 nl of crystallization solution in 96-well sitting-drop plates, equilibrated against a 50-μl reservoir. Crystals of PmC11 were grown at 4 °C in mother liquor consisting of 0.2 m NH4H2PO4, 20% PEG-3350 (JCSG Core Suite I). Crystals were flash cooled in liquid nitrogen using 10% ethylene glycol as a cryoprotectant prior to data collection and initial screening for diffraction was carried out using the Stanford Automated Mounting system at the Stanford Synchrotron Radiation Lightsource (SSRL, Menlo Park, CA). Single wavelength anomalous dispersion data were collected using a wavelength of 0.9793 Å, at the Advanced Light Source (ALS, beamline 8.2.2, Berkeley, CA) on an ADSC Quantum 315 CCD detector. The data were indexed and integrated with XDS and scaled using XSCALE. The diffraction data were indexed in space group P21 with a = 39.11, b = 108.68, c = 77.97 Å, and β = 94.32°. The unit cell contained two molecules in the asymmetric unit resulting in a solvent content of 39% (Matthews' coefficient (Vm) of 2.4 Å3 Da−1). Structure Determination The PmC11 structure was determined by the single wavelength anomalous dispersion method using an x-ray wavelength corresponding to the peak of the selenium K edge. Initial phases were derived using the autoSHARP interface, which included density modification with SOLOMON. Good quality electron density was obtained at 1.7-Å resolution, allowing an initial model to be obtained by automated model building with ARP/wARP. Model completion and refinement were iteratively performed with COOT and REFMAC to produce a final model with an Rcryst and Rfree of 14.3 and 17.5%, respectively. The refinement included experimental phase restraints in the form of Hendrickson-Lattman coefficients, TLS refinement with one TLS group per molecule in the asymmetric unit, and NCS restraints. The refined structure contains residues 24–375 and 28–375 for the two molecules in the crystallographic asymmetric unit. Structural validation was carried using the JCSG Quality Control Server that analyzes both the coordinates and data using a variety of structural validation tools to confirm the stereochemical quality of the model (ADIT, MOLPROBITY, and WHATIF 5.0) and agreement between model and data (SGCHECK and RESOLVE). All of the main-chain torsion angles were in the allowed regions of the Ramachandran plot and the MolProbity overall clash score for the structure was 2.09 (within the 99th percentile for its resolution). The atomic coordinates and structure factors for PmC11 have been deposited in the Protein Data Bank (PDB) with the accession code 3UWS. Data collection, model, and refinement statistics are reported in Table 1. Crystallographic statistics for PDB code 3UWS Values in parentheses are for the highest resolution shell. Data collection\t\t \t    Wavelength (Å)\t0.9793\t \t    Space group\tP21\t \t    Unit cell dimensions a, b, c (Å); β°\t39.11, 108.68, 77.97; β = 94.32°\t \t    Resolution range (Å)\t28.73–1.70 (1.79–1.70)\t \t    Unique reflections\t70,913\t \t    Rmergea on I (%)\t10.2 (49.0)\t \t    Rmeasb on I (%)\t11.0 (52.7)\t \t    Rpimc on I (%)\t4.1 (19.2)\t \t    I/σI\t15.6 (4.6)\t \t    Wilson B (Å2)\t15.9\t \t    Completeness (%)\t99.6 (99.8)\t \t    Multiplicity\t7.3 (7.5)\t \t\t \tModel and refinement\t\t \t    Reflections (total/test)\t70,883/3,577\t \t    Rcryst/Rfreed (%)\t14.3/17.5\t \t    No. protein residues/atoms\t700/5612\t \t    No. of water/EDO molecules\t690/7\t \t    ESUe based on Rfree (Å)\t0.095\t \t    B-values (Å2)\t\t \t        Average isotropic B (overall)\t20.0\t \t        Protein overall\t18.8\t \t        All main/side chains\t16.7/20.8\t \t        Solvent/EDO\t29.4/35.6\t \t    RMSDg\t\t \t        Bond lengths (Å)\t0.01\t \t        Bond angles (°)\t1.6\t \t    Ramachandran analysis (%)\t\t \t        Favored regions\t97.0\t \t        Allowed regions\t3.0\t \t        Outliers\t0.0\t \t a Rmerge = ΣhklΣi|Ii(hkl) − 〈I(hkl)〉|/Σhkl Σi(hkl). b Rmeas = Σhkl[N/(N-1)]1/2Σi|Ii(hkl) − 〈I(hkl)〉|/ΣhklΣiIi(hkl). c Rpim (precision-indicating Rmerge) = Σhkl[(1/(N-1)]1/2 Σi|Ii (hkl) − 〈I(hkl)〉|/ΣhklΣi Ii(hkl), where n is the multiplicity of reflection hkl, and Ii(hkl) and 〈I(hkl)〉 are the intensity of the ith measurement and the average intensity of reflection hkl, respectively. d Rcryst and Rfree = Σ‖Fobs| − |Fcalc‖/Σ|Fobs| for reflections in the working and test sets, respectively, where Fobs and Fcalc are the observed and calculated structure-factor amplitudes, respectively. Rfree is the same as Rcryst but for 5% of the total reflections chosen at random and omitted from structural refinement. e ESU is the estimated standard uncertainties of atoms. f The average isotropic B includes TLS and residual B components. g RMSD, root-mean-square deviation. Structural Analysis The primary sequence alignment with assigned secondary structure was prepared using CLUSTAL OMEGA and ALINE. The topology diagram was produced with TOPDRAW and all three-dimensional structural figures were prepared with PyMol with the electrostatic surface potential calculated with APBS and contoured at ±5 kT/e. Architectural comparisons with known structures revealed that PmC11 was most structurally similar to caspase-7, gingipain-K, and legumain (PBD codes 4hq0, 4tkx, and 4aw9, respectively). The statistical significance of the structural alignment between PmC11 and both caspase-7 and gingipain-K is equivalent (Z-score of 9.2) with legumain giving a very similar result (Z-score of 9.1). Of note, the β-strand topology of the CDP domains of Clostridium difficile toxin B (family C80; TcdB; PDB code 3pee) is identical to that observed in the PmC11 β-sheet, but the Z-score from DaliLite was notably less at 7.6. It is possible that the PmC11 structure is more closely related to the C80 family than other families in clan CD, and appear to reside on the same branch of the phylogenetic tree based on structure. Protein Production for Biochemical Assays The PmCD00547516 plasmid described above was obtained from the PSI:Biology Materials Repository and used to generate a cleavage site mutant PmC11K147A and an active-site mutant PmC11C179A using the QuikChange Site-directed Mutagenesis kit (Stratagene) as per the manufacturer's instructions using the following primers: K147A mutant (forward): CAGAATAAGCTGGCAGCGTTCGGACAGGACG, and K147A mutant (reverse): CGTCCTGTCCGAACGCTGCCAGCTTATTCTG; C179A mutant (forward): CCTGTTCGATGCCGCCTACATGGCAAGC, and C179A mutant (reverse): GCTTGCCATGTAGGCGGCATCGAACAGG. The expression plasmids containing PmC11 were transformed into E. coli BL21 Star (DE3) and grown in Luria-Bertani media containing 30 μg ml−1 of kanamycin at 37 °C until an optical density (600 nm) of ∼0.6 was reached. l-Arabinose was added to a final concentration of 0.2% (w/v) and the cells incubated overnight at 25 °C. Compared with the protein production for crystallography, a slightly modified purification protocol was employed for biochemical assays. Initially, the cells were resuspended in 20 mm sodium phosphate (pH 7.5), 150 mm NaCl (Lysis Buffer 2) containing an EDTA-free protease inhibitor mixture (cOmplete, Roche Applied Science). Cells were disrupted by three passages (15 KPSI) through a One-Shot cell disruptor (Constant Systems) followed by centrifugation at 20,000 × g for 20 min at 4 °C. The supernatant was collected and sterile-filtered (0.2 μm) before being applied to a 5-ml HisTrap HP column (GE Healthcare) equilibrated in Lysis Buffer 2 containing 25 mm imidazole, and the protein was eluted in the same buffer containing 250 mm imidazole. The peak fractions were pooled and buffer exchanged into the assay buffer (20 mm Tris, 150 mm NaCl, pH 8.0) using a PD-10 column. When required, purified PmC11 was concentrated using Vivaspin 2 30-K centrifugal concentrators (Sartorius). Protein concentration was routinely measured using Bradford's reagent (Bio-Rad) with a BSA standard. Fluorogenic Substrate Activity Assays The release of the fluorescent group AMC (7-amino-4-methylcoumarin) from potential peptide substrates was used to assess the activity of PmC11. Peptidase activity was tested using 20 μg of PmC11 and 100 μm substrate (unless otherwise stated) in assay buffer to a final reaction volume of 200 μl and all samples were incubated (without substrate) at 37 °C for 16 h prior to carrying out the assay. The substrate and plate reader were brought to 37 °C for 20 min prior to the addition of the PmC11 and samples prepared without PmC11 were used as blanks (negative controls). The curves were plotted using the blank-corrected fluorescence units against the time of acquisition (in min). The assays were carried out in black 96-well flat-bottomed plates (Greiner). AMC fluorescence was measured using a PHERAstar FS plate reader (BMG Labtech) with excitation and emission wavelengths of 355 and 460 nm, respectively. To investigate the substrate specificity of PmC11, substrates Z-GGR-AMC, Bz-R-AMC, Z-GP-AMC, Z-HGP-AMC, Ac-DEVD-AMC (all Bachem), BOC-VLK-AMC, and BOC-K-AMC (both PeptaNova) were prepared at 100 mm in 100% dimethyl sulfoxide. The amount of AMC (micromoles) released was calculated by generating an AMC standard curve (as described in Ref.) and the specific activity of PmC11 was calculated as picomoles of AMC released per min per mg of the protein preparation. The reaction rates (Vmax) and Km values were determined for mutants PmC11K147A and PmC11C179A by carrying out the activity assay at varying concentrations of Bz-R-AMC between 0 and 600 μm. The blank-corrected relative fluorescence units were plotted against time (min) with ΔFU/T giving the reaction rate. The Km and Vmax of PmC11 and PmC11K147A against an R-AMC substrate were determined from the Lineweaver-Burk plot as described, calculated using GraphPad Prism6 software. All experiments were carried out in triplicate. Effect of VRPR-FMK on PmC11 To test the effect of the inhibitor on the activity of PmC11, 25 μm Z-VRPR-FMK (100 mm stock in 100% dimethyl sulfoxide, Enzo Life Sciences), 20 μg of PmC11, 100 μm R-AMC substrate, 1 mm EGTA were prepared in the assay buffer and the activity assay carried out as described above. A gel-shift assay, to observe Z-VRPR-FMK binding to PmC11, was also set up using 20 μg of PmC11, 25 μm inhibitor, 1 mm EGTA in assay buffer. The reactions were incubated at 37 °C for 20 min before being stopped by the addition SDS-PAGE sample buffer. Samples were analyzed by loading the reaction mixture on a 10% NuPAGE BisTris gel using MES buffer. Effect of Cations on PmC11 The enzyme activity of PmC11 was tested in the presence of various divalent cations: Mg2+, Ca2+, Mn2+, Co2+, Fe2+, Zn2+, and Cu2+. The final concentration of the salts (MgSO4, CaCl2, MnCl2, CoCl2, FeSO4, ZnCl2, and CuSO4) was 1 mm and the control was set up without divalent ions but with addition of 1 mm EGTA. The assay was set up using 20 mg of PmC11, 1 mm salts, 100 μm R-AMC substrate, and the assay buffer, and incubated at 37 °C for 16 h. The activity assay was carried out as described above. Size Exclusion Chromatography Affinity-purified PmC11 was loaded onto a HiLoad 16/60 Superdex 200 gel filtration column (GE Healthcare) equilibrated in the assay buffer. The apparent molecular weight of PmC11 was determined from calibration curves based on protein standards of known molecular weights. Autoprocessing Profile of PmC11 Autoprocessing of PmC11 was evaluated by incubating the enzyme at 37 °C and removing samples at 1-h intervals from 0 to 16 h and placing into SDS-PAGE loading buffer to stop the processing. Samples were then analyzed on a 4–12% NuPAGE (Thermo Fisher) Novex BisTris gel run in MES buffer. Autoprocessing Cleavage Site Analysis To investigate whether processing is a result of intra- or inter-molecular cleavage, the PmC11C179A mutant was incubated with increasing concentrations of activated PmC11 (0, 0.1, 0.2, 0.5, 1, 2, and 5 μg). The final assay volume was 40 μl and the proteins were incubated at 37 °C for 16 h in the PmC11 assay buffer. To stop the reaction, NuPAGE sample buffer was added to the protein samples and 20 μl was analyzed on 10% NuPAGE Novex BisTris gel using MES buffer. These studies revealed no apparent cleavage of PmC11C179A by the active enzyme at low concentrations of PmC11 and that only limited cleavage was observed when the ratio of active enzyme (PmC11: PmC11C179A) was increased to ∼1:10 and 1:4. Results Structure of PmC11 The crystal structure of the catalytically active form of PmC11 revealed an extended caspase-like α/β/α sandwich architecture comprised of a central nine-stranded β-sheet, with an unusual C-terminal domain (CTD), starting at Lys250. A single cleavage was observed in the polypeptide chain at Lys147 (Fig. 1, A and B), where both ends of the cleavage site are fully visible and well ordered in the electron density. The central nine-stranded β-sheet (β1–β9) of PmC11 consists of six parallel and three anti-parallel β-strands with 4↑3↓2↑1↑5↑6↑7↓8↓9↑ topology (Fig. 1A) and the overall structure includes 14 α-helices with six (α1–α2 and α4–α7) closely surrounding the β-sheet in an approximately parallel orientation. Helices α1, α7, and α6 are located on one side of the β-sheet with α2, α4, and α5 on the opposite side (Fig. 1A). Helix α3 sits at the end of the loop following β5 (L5), just preceding the Lys147 cleavage site, with both L5 and α3 pointing away from the central β-sheet and toward the CTD, which starts with α8. The structure also includes two short β-hairpins (βA–βB and βD–βE) and a small β-sheet (βC–βF), which is formed from two distinct regions of the sequence (βC precedes α11, α12 and β9, whereas βF follows the βD-βE hairpin) in the middle of the CTD (Fig. 1B). Crystal structure of a C11 peptidase from P. merdae.\nA, primary sequence alignment of PmC11 (Uniprot ID A7A9N3) and clostripain (Uniprot ID P09870) from C. histolyticum with identical residues highlighted in gray shading. The secondary structure of PmC11 from the crystal structure is mapped onto its sequence with the position of the PmC11 catalytic dyad, autocatalytic cleavage site (Lys147), and S1 binding pocket Asp (Asp177) highlighted by a red star, a red downturned triangle, and a red upturned triangle, respectively. Connecting loops are colored gray, the main β-sheet is in orange, with other strands in olive, α-helices are in blue, and the nonapeptide linker of clostripain that is excised upon autocleavage is underlined in red. Sequences around the catalytic site of clostripain and PmC11 align well. B, topology diagram of PmC11 colored as in A except that additional (non-core) β-strands are in yellow. Helices found on either side of the central β-sheet are shown above and below the sheet, respectively. The position of the catalytic dyad (H, C) and the processing site (Lys147) are highlighted. Helices (1–14) and β-strands (1–9 and A-F) are numbered from the N terminus. The core caspase-fold is highlighted in a box. C, tertiary structure of PmC11. The N and C termini (N and C) of PmC11 along with the central β-sheet (1–9), helix α5, and helices α8, α11, and α13 from the C-terminal domain, are all labeled. Loops are colored gray, the main β-sheet is in orange, with other β-strands in yellow, and α-helices are in blue. The CTD of PmC11 is composed of a tight helical bundle formed from helices α8–α14 and includes strands βC and βF, and β-hairpin βD–βE. The CTD sits entirely on one side of the enzyme interacting only with α3, α5, β9, and the loops surrounding β8. Of the interacting secondary structure elements, α5 is perhaps the most interesting. This helix makes a total of eight hydrogen bonds with the CTD, including one salt bridge (Arg191-Asp255) and is surrounded by the CTD on one side and the main core of the enzyme on the other, acting like a linchpin holding both components together (Fig. 1C). Structural Comparisons PmC11 is, as expected, most structurally similar to other members of clan CD with the top hits in a search of known structures being caspase-7, gingipain-K, and legumain (PBD codes 4hq0, 4tkx, and 4aw9, respectively) (Table 2). The C-terminal domain is unique to PmC11 within clan CD and structure comparisons for this domain alone does not produce any hits in the PDB (DaliLite, PDBeFold), suggesting a completely novel fold. As the archetypal and arguably most well studied member of clan CD, the caspases were used as the basis to investigate the structure/function relationships in PmC11, with caspase-7 as the representative member. Six of the central β-strands in PmC11 (β1–β2 and β5–β8) share the same topology as the six-stranded β-sheet found in caspases, with strands β3, β4, and β9 located on the outside of this core structure (Fig. 1B, box). His133 and Cys179 were found at locations structurally homologous to the caspase catalytic dyad, and other clan CD structures, at the C termini of strands β5 and β6, respectively (Figs. 1, A and B, and 2A). A multiple sequence alignment of C11 proteins revealed that these residues are highly conserved (data not shown). Summary of PDBeFOLD superposition of structures found to be most similar to PmC11 in the PBD based on DaliLite  The results are ordered in terms of structural homology (QH), where %SSEPC-X is the percentage of the SSEs in the PmC11 that can be identified in the target X (where X = caspase-7, legumain, gingipain, and TcdB-CPD; % SSEX-PC is the percentage of SSEs in X (as above) that can be identified in PmC11 (as above); % sequence ID is the percentage sequence identity after structural alignment; Nalign is the number of matched residues; and r.m.s. deviation the root mean squared deviation on the Cα positions of the matched residues. Enzyme\tFamily\tPDB code\tQH\tZ-score\t%SSEPC-X\t%SSEX-PC\t% Seq. ID\tNalign\tRMSD (Å)\tNStrands\t \tPmC11\tC11\t3UWS\t1.00\t33.4\t100\t100\t100\t352\t0.00\t9\t \tCaspase-7\tC14A\t4HQ0\t0.16\t4.3\t38\t79\t14\t162\t3.27\t6\t \tLegumain\tC13\t4AW9\t0.13\t5.5\t31\t53\t13\t161\t2.05\t6\t \tTcdB-CPD\tC80\t3PEE\t0.10\t4.9\t28\t50\t12\t138\t3.18\t9\t \tGingipain\tC25\t4TKX\t0.07\t5.4\t28\t27\t12\t153\t2.97\t10\t \t Biochemical and structural characterization of PmC11.\nA, ribbon representation of the overall structure of PmC11 illustrating the catalytic site, cleavage site displacement, and potential S1 binding site. The overall structure of PmC11 is shown in gray, looking down into the catalytic site with the catalytic dyad in red. The two ends of the autolytic cleavage site (Lys147 and Ala148, green) are displaced by 19.5 Å (thin black line) from one another and residues in the potential substrate binding pocket are highlighted in blue. B, size exclusion chromatography of PmC11. PmC11 migrates as a monomer with a molecular mass around 41 kDa calculated from protein standards of known molecular weights. Elution fractions across the major peak (1–6) were analyzed by SDS-PAGE on a 4–12% gel in MES buffer. C, the active form of PmC11 and two mutants, PmC11C179A (C) and PmC11K147A (K), were examined by SDS-PAGE (lane 1) and Western blot analysis using an anti-His antibody (lane 2) show that PmC11 autoprocesses, whereas mutants, PmC11C179A and PmC11K147A, do not show autoprocessing in vitro. D, cysteine peptidase activity of PmC11. Km and Vmax of PmC11 and K147A mutant were determined by monitoring change in the fluorescence corresponding to AMC release from Bz-R-AMC. Reactions were performed in triplicate and representative values ± S.D. are shown. E, intermolecular processing of PmC11C179A by PmC11. PmC11C179A (20 μg) was incubated overnight at 37 °C with increasing amounts of processed PmC11 and analyzed on a 10% SDS-PAGE gel. Inactive PmC11C179A was not processed to a major extent by active PmC11 until around a ratio of 1:4 (5 μg of active PmC11). A single lane of 20 μg of active PmC11 (labeled 20) is shown for comparison. F, activity of PmC11 against basic substrates. Specific activity is shown ± S.D. from three independent experiments. G, electrostatic surface potential of PmC11 shown in a similar orientation, where blue and red denote positively and negatively charged surface potential, respectively, contoured at ±5 kT/e. The position of the catalytic dyad, one potential key substrate binding residue Asp177, and the ends of the cleavage site Lys147 and Ala148 are indicated. Five of the α-helices surrounding the β-sheet of PmC11 (α1, α2, α4, α6, and α7) are found in similar positions to the five structurally conserved helices in caspases and other members of clan CD, apart from family C80. Other than its more extended β-sheet, PmC11 differs most significantly from other clan CD members at its C terminus, where the CTD contains a further seven α-helices and four β-strands after β8. Autoprocessing of PmC11 Purification of recombinant PmC11 (molecular mass = 42.6 kDa) revealed partial processing into two cleavage products of 26.4 and 16.2 kDa, related to the observed cleavage at Lys147 in the crystal structure (Fig. 2A). Incubation of PmC11 at 37 °C for 16 h, resulted in a fully processed enzyme that remained as an intact monomer when applied to a size-exclusion column (Fig. 2B). The single cleavage site of PmC11 at Lys147 is found immediately after α3, in loop L5 within the central β-sheet (Figs. 1, A and B, and 2A). The two ends of the cleavage site are remarkably well ordered in the crystal structure and displaced from one another by 19.5 Å (Fig. 2A). Moreover, the C-terminal side of the cleavage site resides near the catalytic dyad with Ala148 being 4.5 and 5.7 Å from His133 and Cys179, respectively. Consequently, it appears feasible that the helix attached to Lys147 (α3) could be responsible for steric autoinhibition of PmC11 when Lys147 is covalently bonded to Ala148. Thus, the cleavage would be required for full activation of PmC11. To investigate this possibility, two mutant forms of the enzyme were created: PmC11C179A (a catalytically inactive mutant) and PmC11K147A (a cleavage-site mutant). Initial SDS-PAGE and Western blot analysis of both mutants revealed no discernible processing occurred as compared with active PmC11 (Fig. 2C). The PmC11K147A mutant enzyme had a markedly different reaction rate (Vmax) compared with WT, where the reaction velocity of PmC11 was 10 times greater than that of PmC11K147A (Fig. 2D). Taken together, these data reveal that PmC11 requires processing at Lys147 for optimum activity. To investigate whether processing is a result of intra- or intermolecular cleavage, the PmC11C179A mutant was incubated with increasing concentrations of processed and activated PmC11. These studies revealed that there was no apparent cleavage of PmC11C179A by the active enzyme at low concentrations of PmC11 and that only limited cleavage was observed when the ratio of active enzyme (PmC11:PmC11C179A) was increased to ∼1:10 and 1:4, with complete cleavage observed at a ratio of 1:1 (Fig. 2E). This suggests that cleavage of PmC11C179A was most likely an effect of the increasing concentration of PmC11 and intermolecular cleavage. Collectively, these data suggest that the pro-form of PmC11 is autoinhibited by a section of L5 blocking access to the active site, prior to intramolecular cleavage at Lys147. This cleavage subsequently allows movement of the region containing Lys147 and the active site to open up for substrate access. Substrate Specificity of PmC11 The autocatalytic cleavage of PmC11 at Lys147 (sequence KLK∧A) demonstrates that the enzyme accepts substrates with Lys in the P1 position. The substrate specificity of the enzyme was further tested using a variety of fluorogenic substrates. As expected, PmC11 showed no activity against substrates with Pro or Asp in P1 but was active toward substrates with a basic residue in P1 such as Bz-R-AMC, Z-GGR-AMC, and BOC-VLK-AMC. The rate of cleavage was ∼3-fold greater toward the single Arg substrate Bz-R-AMC than for the other two (Fig. 2F) and, unexpectedly, PmC11 showed no activity toward BOC-K-AMC. These results confirm that PmC11 accepts substrates containing Arg or Lys in P1 with a possible preference for Arg. The catalytic dyad of PmC11 sits near the bottom of an open pocket on the surface of the enzyme at a conserved location in the clan CD family. The PmC11 structure reveals that the catalytic dyad forms part of a large acidic pocket (Fig. 2G), consistent with a binding site for a basic substrate. This pocket is lined with the potential functional side chains of Asn50, Asp177, and Thr204 with Gly134, Asp207, and Met205 also contributing to the pocket (Fig. 2A). Interestingly, these residues are in regions that are structurally similar to those involved in the S1 binding pockets of other clan CD members (shown in Ref.). Because PmC11 recognizes basic substrates, the tetrapeptide inhibitor Z-VRPR-FMK was tested as an enzyme inhibitor and was found to inhibit both the autoprocessing and activity of PmC11 (Fig. 3A). Z-VRPR-FMK was also shown to bind to the enzyme: a size-shift was observed, by SDS-PAGE analysis, in the larger processed product of PmC11 suggesting that the inhibitor bound to the active site (Fig. 3B). A structure overlay of PmC11 with the MALT1-paracacaspase (MALT1-P), in complex with Z-VRPR-FMK, revealed that the PmC11 dyad sits in a very similar position to that of active MALT1-P and that Asn50, Asp177, and Asp207 superimpose well with the principal MALT1-P inhibitor binding residues (Asp365, Asp462, and Glu500, respectively (VRPR-FMK from MALT1-P with the corresponding PmC11 residues from the structural overlay is shown in Fig. 1D), as described in Ref.). Asp177 is located near the catalytic cysteine and is conserved throughout the C11 family, suggesting it is the primary S1 binding site residue. In the structure of PmC11, Asp207 resides on a flexible loop pointing away from the S1 binding pocket (Fig. 3C). However, this loop has been shown to be important for substrate binding in clan CD and this residue could easily rotate and be involved in substrate binding in PmC11. Thus, Asn50, Asp177, and Asp207 are most likely responsible for the substrate specificity of PmC11. Asp177 is highly conserved throughout the clan CD C11 peptidases and is thought to be primarily responsible for substrate specificity of the clan CD enzymes, as also illustrated from the proximity of these residues relative to the inhibitor Z-VRPR-FMK when PmC11 is overlaid on the MALT1-P structure (Fig. 3C). PmC11 binds and is inhibited by Z-VRPR-FMK and does not require Ca2+ for activity.\nA, PmC11 activity is inhibited by Z-VRPR-FMK. Cleavage of Bz-R-AMC by PmC11 was measured in a fluorometric activity assay with (+, purple) and without (−, red) Z-VRPR-FMK. The relative fluorescence units of AMC released are plotted against time (min) (n = 3; ±S.D.). B, gel-shift assay reveals that Z-VRPR-FMK binds to PmC11. PmC11 was incubated with (+) or without (−) Z-VRPR-FMK and the samples analyzed on a 10% SDS-PAGE gel. A size shift can be observed in the larger processed product of PmC11 (26.1 kDa). C, PmC11 with the Z-VRPR-FMK from the MALT1-paracacaspase (MALT1-P) superimposed. A three-dimensional structural overlay of Z-VRPR-FMK from the MALT1-P complex onto PmC11. The position and orientation of Z-VRPR-FMK was taken from superposition of the PmC11 and MALTI_P structures and indicates the presumed active site of PmC11. Residues surrounding the inhibitor are labeled and represent potentially important binding site residues, labeled in black and shown in an atomic representation. Carbon atoms are shown in gray, nitrogen in blue, and oxygen in red. C, divalent cations do not increase the activity of PmC11. The cleavage of Bz-R-AMC by PmC11 was measured in the presence of the cations Ca2+, Mn2+, Zn2+, Co2+, Cu2+, Mg2+, and Fe3+ with EGTA as a negative control, and relative fluorescence measured against time (min). The addition of cations produced no improvement in activity of PmC11 when compared in the presence of EGTA, suggesting that PmC11 does not require metal ions for proteolytic activity. Furthermore, Cu2+, Fe2+, and Zn2+ appear to inhibit PmC11. Comparison with Clostripain Clostripain from C. histolyticum is the founding member of the C11 family of peptidases and contains an additional 149 residues compared with PmC11. A multiple sequence alignment revealed that most of the secondary structural elements are conserved between the two enzymes, although they are only ∼23% identical (Fig. 1A). Nevertheless, PmC11 may be a good model for the core structure of clostripain. The primary structural alignment also shows that the catalytic dyad in PmC11 is structurally conserved in clostripain (Fig. 1A). Unlike PmC11, clostripain has two cleavage sites (Arg181 and Arg190), which results in the removal of a nonapeptide, and is required for full activation of the enzyme (highlighted in Fig. 1A). Interestingly, Arg190 was found to align with Lys147 in PmC11. In addition, the predicted primary S1-binding residue in PmC11 Asp177 also overlays with the residue predicted to be the P1 specificity determining residue in clostripain (Asp229, Fig. 1A). As studies on clostripain revealed addition of Ca2+ ions are required for full activation, the Ca2+ dependence of PmC11 was examined. Surprisingly, Ca2+ did not enhance PmC11 activity and, furthermore, other divalent cations, Mg2+, Mn2+, Co2+, Fe2+, Zn2+, and Cu2+, were not necessary for PmC11 activity (Fig. 3D). In support of these findings, EGTA did not inhibit PmC11 suggesting that, unlike clostripain, PmC11 does not require Ca2+ or other divalent cations, for activity. Discussion The crystal structure of PmC11 now provides three-dimensional information for a member of the clostripain C11 family of cysteine peptidases. The enzyme exhibits all of the key structural elements of clan CD members, but is unusual in that it has a nine-stranded central β-sheet with a novel C-terminal domain. The structural similarity of PmC11 with its nearest structural neighbors in the PDB is decidedly low, overlaying better with six-stranded caspase-7 than any of the other larger members of the clan (Table 2). The substrate specificity of PmC11 is Arg/Lys and the crystal structure revealed an acidic pocket for specific binding of such basic substrates. In addition, the structure suggested a mechanism of self-inhibition in both PmC11 and clostripain and an activation mechanism that requires autoprocessing. PmC11 differs from clostripain in that is does not appear to require divalent cations for activation. Several other members of clan CD require processing for full activation including legumain, gingipain-R, MARTX-CPD, and the effector caspases, e.g. caspase-7. To date, the effector caspases are the only group of enzymes that require cleavage of a loop within the central β-sheet. This is also the case in PmC11, although the cleavage loop is structurally different to that found in the caspases and follows the catalytic His (Fig. 1A), as opposed to the Cys in the caspases. All other clan CD members requiring cleavage for full activation do so at sites external to their central sheets. The caspases and gingipain-R both undergo intermolecular (trans) cleavage and legumain and MARTX-CPD are reported to perform intramolecular (cis) cleavage. In addition, several members of clan CD exhibit self-inhibition, whereby regions of the enzyme block access to the active site. Like PmC11, these structures show preformed catalytic machinery and, for a substrate to gain access, movement and/or cleavage of the blocking region is required. The structure of PmC11 gives the first insight into this class of relatively unexplored family of proteins and should allow important catalytic and substrate binding residues to be identified in a variety of orthologues. Indeed, insights gained from an analysis of the PmC11 structure revealed the identity of the Trypanosoma brucei PNT1 protein as a C11 cysteine peptidase with an essential role in organelle replication. The PmC11 structure should provide a good basis for structural modeling and, given the importance of other clan CD enzymes, this work should also advance the exploration of these peptidases and potentially identify new biologically important substrates. Author Contributions K. M., J. S. G., D. D., I. A. W., and J. C. M. designed the research; K. M., J. S. G., and D. D. performed the research; K. M., J. S. G., D. D., G. H. C., A. S., M. A. E., and J. C. M. analyzed the data; A. G., S. A. L., A. M. D., M. A. E., and I. A. W. supervised various components of the JCSG structural genomics pipeline; M. K. G., A. G., S. A. L., A. M. D., and M. A. E. contributed reagents, materials, and analysis tools; and K. M., J. S. G., G. H. C., M. A. E., I. A. W., and J. C. M. wrote the paper. This work was supported by the Medical Research Council Grant MR/K019384, Wellcome Trust Grants 091790 and 104111, and National Institutes of Health Grant U54 GM094586 (JCSG). The SSRL Structural Molecular Biology Program is supported by the DOE Office of Biological and Environmental Research and National Institutes of Health (NIH), National Center for Research Resources, Biomedical Technology Program Grant P41RR001209, and the NIMGS. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIGMS or NIH. The authors declare that they have no conflicts of interest with the contents of this article. The atomic coordinates and structure factors (code 3UWS) have been deposited in the Protein Data Bank (http://wwpdb.org/). JCSG Joint Center for Structural Genomics PIPE polymerase incomplete primer extension TCEP Tris(2-carboxyethyl)phosphine AMC 7-amino-4-methylcoumarin PDB Protein Data Bank BisTris 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol Z benzyloxycarbonyl FMK fluoromethyl ketone CTD C-terminal domain Bz-R-AMC benzoyl-l-Arg-4-methylcoumaryl-7-amide Z-GGR-AMC benzyloxycarbonyl-Gly-Gly-Arg-AMC BOC-VLK-AMC t-butyloxycarbonyl-Val-Leu-Lys. 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diff --git a/annotated_BioC_JSON/PMC4852598_ann.json b/annotated_BioC_JSON/PMC4852598_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..84df4100577592013822190fafd45613eeb49333
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4852598_ann.json
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+[{"sourceid":"4852598","sourcedb":"","project":"","target":"","text":"Structural basis for Mep2 ammonium transceptor activation by phosphorylation Mep2 proteins are fungal transceptors that play an important role as ammonium sensors in fungal development. Mep2 activity is tightly regulated by phosphorylation, but how this is achieved at the molecular level is not clear. Here we report X-ray crystal structures of the Mep2 orthologues from Saccharomyces cerevisiae and Candida albicans and show that under nitrogen-sufficient conditions the transporters are not phosphorylated and present in closed, inactive conformations. Relative to the open bacterial ammonium transporters, non-phosphorylated Mep2 exhibits shifts in cytoplasmic loops and the C-terminal region (CTR) to occlude the cytoplasmic exit of the channel and to interact with His2 of the twin-His motif. The phosphorylation site in the CTR is solvent accessible and located in a negatively charged pocket ∼30 Å away from the channel exit. The crystal structure of phosphorylation-mimicking Mep2 variants from C. albicans show large conformational changes in a conserved and functionally important region of the CTR. The results allow us to propose a model for regulation of eukaryotic ammonium transport by phosphorylation.  Mep2 proteins are tightly regulated fungal ammonium transporters. Here, the authors report the crystal structures of closed states of Mep2 proteins and propose a model for their regulation by comparing them with the open ammonium transporters of bacteria. Transceptors are membrane proteins that function not only as transporters but also as receptors/sensors during nutrient sensing to activate downstream signalling pathways. A common feature of transceptors is that they are induced when cells are starved for their substrate. While most studies have focused on the Saccharomyces cerevisiae transceptors for phosphate (Pho84), amino acids (Gap1) and ammonium (Mep2), transceptors are found in higher eukaryotes as well (for example, the mammalian SNAT2 amino-acid transporter and the GLUT2 glucose transporter). One of the most important unresolved questions in the field is how the transceptors couple to downstream signalling pathways. One hypothesis is that downstream signalling is dependent on a specific conformation of the transporter. Mep2 (methylammonium (MA) permease) proteins are ammonium transceptors that are ubiquitous in fungi. They belong to the Amt/Mep/Rh family of transporters that are present in all kingdoms of life and they take up ammonium from the extracellular environment. Fungi typically have more than one Mep paralogue, for example, Mep1-3 in S. cerevisiae. Of these, only Mep2 proteins function as ammonium receptors/sensors in fungal development. Under conditions of nitrogen limitation, Mep2 initiates a signalling cascade that results in a switch from the yeast form to filamentous (pseudohyphal) growth that may be required for fungal pathogenicity. As is the case for other transceptors, it is not clear how Mep2 interacts with downstream signalling partners, but the protein kinase A and mitogen-activated protein kinase pathways have been proposed as downstream effectors of Mep2 (refs). Compared with Mep1 and Mep3, Mep2 is highly expressed and functions as a low-capacity, high-affinity transporter in the uptake of MA. In addition, Mep2 is also important for uptake of ammonium produced by growth on other nitrogen sources. With the exception of the human RhCG structure, no structural information is available for eukaryotic ammonium transporters. By contrast, several bacterial Amt orthologues have been characterized in detail via high-resolution crystal structures and a number of molecular dynamics (MD) studies. All the solved structures including that of RhCG are very similar, establishing the basic architecture of ammonium transporters. The proteins form stable trimers, with each monomer having 11 transmembrane (TM) helices and a central channel for the transport of ammonium. All structures show the transporters in open conformations. Intriguingly, fundamental questions such as the nature of the transported substrate and the transport mechanism are still controversial. Where earlier studies favoured the transport of ammonia gas, recent data and theoretical considerations suggest that Amt/Mep proteins are instead active, electrogenic transporters of either NH4+ (uniport) or NH3/H+ (symport). A highly conserved pair of channel-lining histidine residues dubbed the twin-His motif may serve as a proton relay system while NH3 moves through the channel during NH3/H+ symport. Ammonium transport is tightly regulated. In animals, this is due to toxicity of elevated intracellular ammonium levels, whereas for microorganisms ammonium is a preferred nitrogen source. In bacteria, amt genes are present in an operon with glnK, encoding a PII-like signal transduction class protein. By binding tightly to Amt proteins without inducing a conformational change in the transporter, GlnK sterically blocks ammonium conductance when nitrogen levels are sufficient. Under conditions of nitrogen limitation, GlnK becomes uridylated, blocking its ability to bind and inhibit Amt proteins. Importantly, eukaryotes do not have GlnK orthologues and have a different mechanism for regulation of ammonium transport activity. In plants, transporter phosphorylation and dephosphorylation are known to regulate activity. In S. cerevisiae, phosphorylation of Ser457 within the C-terminal region (CTR) in the cytoplasm was recently proposed to cause Mep2 opening, possibly via inducing a conformational change. To elucidate the mechanism of Mep2 transport regulation, we present here X-ray crystal structures of the Mep2 transceptors from S. cerevisiae and C. albicans. The structures are similar to each other but show considerable differences to all other ammonium transporter structures. The most striking difference is the fact that the Mep2 proteins have closed conformations. The putative phosphorylation site is solvent accessible and located in a negatively charged pocket ∼30 Å away from the channel exit. The channels of phosphorylation-mimicking mutants of C. albicans Mep2 are still closed but show large conformational changes within a conserved part of the CTR. Together with a structure of a C-terminal Mep2 variant lacking the segment containing the phosphorylation site, the results allow us to propose a structural model for phosphorylation-based regulation of eukaryotic ammonium transport. Results General architecture of Mep2 ammonium transceptors The Mep2 protein of S. cerevisiae (ScMep2) was overexpressed in S. cerevisiae in high yields, enabling structure determination by X-ray crystallography using data to 3.2 Å resolution by molecular replacement (MR) with the archaebacterial Amt-1 structure (see Methods section). Given that the modest resolution of the structure and the limited detergent stability of ScMep2 would likely complicate structure–function studies, several other fungal Mep2 orthologues were subsequently overexpressed and screened for diffraction-quality crystals. Of these, Mep2 from C. albicans (CaMep2) showed superior stability in relatively harsh detergents such as nonyl-glucoside, allowing structure determination in two different crystal forms to high resolution (up to 1.5 Å). Despite different crystal packing (Supplementary Table 1), the two CaMep2 structures are identical to each other and very similar to ScMep2 (Cα r.m.s.d. (root mean square deviation)=0.7 Å for 434 residues), with the main differences confined to the N terminus and the CTR (Fig. 1). Electron density is visible for the entire polypeptide chains, with the exception of the C-terminal 43 (ScMep2) and 25 residues (CaMep2), which are poorly conserved and presumably disordered. Both Mep2 proteins show the archetypal trimeric assemblies in which each monomer consists of 11 TM helices surrounding a central pore. Important functional features such as the extracellular ammonium binding site, the Phe gate and the twin-His motif within the hydrophobic channel are all very similar to those present in the bacterial transporters and RhCG. In the remainder of the manuscript, we will specifically discuss CaMep2 due to the superior resolution of the structure. Unless specifically stated, the drawn conclusions also apply to ScMep2. While the overall architecture of Mep2 is similar to that of the prokaryotic transporters (Cα r.m.s.d. with Amt-1=1.4 Å for 361 residues), there are large differences within the N terminus, intracellular loops (ICLs) ICL1 and ICL3, and the CTR. The N termini of the Mep2 proteins are ∼20–25 residues longer compared with their bacterial counterparts (Figs 1 and 2), substantially increasing the size of the extracellular domain. Moreover, the N terminus of one monomer interacts with the extended extracellular loop ECL5 of a neighbouring monomer. Together with additional, smaller differences in other extracellular loops, these changes generate a distinct vestibule leading to the ammonium binding site that is much more pronounced than in the bacterial proteins. The N-terminal vestibule and the resulting inter-monomer interactions likely increase the stability of the Mep2 trimer, in support of data for plant AMT proteins. However, given that an N-terminal deletion mutant (2-27Δ) grows as well as wild-type (WT) Mep2 on minimal ammonium medium (Fig. 3 and Supplementary Fig. 1), the importance of the N terminus for Mep2 activity is not clear. Mep2 channels are closed by a two-tier channel block The largest differences between the Mep2 structures and the other known ammonium transporter structures are located on the intracellular side of the membrane. In the vicinity of the Mep2 channel exit, the cytoplasmic end of TM2 has unwound, generating a longer ICL1 even though there are no insertions in this region compared to the bacterial proteins (Figs 2 and 4). ICL1 has also moved inwards relative to its position in the bacterial Amts. The largest backbone movements of equivalent residues within ICL1 are ∼10 Å, markedly affecting the conserved basic RxK motif (Fig. 4). The head group of Arg54 has moved ∼11 Å relative to that in Amt-1, whereas the shift of the head group of the variable Lys55 residue is almost 20 Å. The side chain of Lys56 in the basic motif points in an opposite direction in the Mep2 structures compared with that of, for example, Amt-1 (Fig. 4). In addition to changing the RxK motif, the movement of ICL1 has another, crucial functional consequence. At the C-terminal end of TM1, the side-chain hydroxyl group of the relatively conserved Tyr49 (Tyr53 in ScMep2) makes a strong hydrogen bond with the ɛ2 nitrogen atom of the absolutely conserved His342 of the twin-His motif (His348 in ScMep2), closing the channel (Figs 4 and 5). In bacterial Amt proteins, this Tyr side chain is rotated ∼4 Å away as a result of the different conformation of TM1, leaving the channel open and the histidine available for its putative role in substrate transport (Supplementary Fig. 2). Compared with ICL1, the backbone conformational changes observed for the neighbouring ICL2 are smaller, but large shifts are nevertheless observed for the conserved residues Glu140 and Arg141 (Fig. 4). Finally, the important ICL3 linking the pseudo-symmetrical halves (TM1-5 and TM6-10) of the transporter is also shifted up to ∼10 Å and forms an additional barrier that closes the channel on the cytoplasmic side (Fig. 5). This two-tier channel block likely ensures that very little ammonium transport will take place under nitrogen-sufficient conditions. The closed state of the channel might also explain why no density, which could correspond to ammonium (or water), is observed in the hydrophobic part of the Mep2 channel close to the twin-His motif. Significantly, this is also true for ScMep2, which was crystallized in the presence of 0.2 M ammonium ions (see Methods section). The final region in Mep2 that shows large differences compared with the bacterial transporters is the CTR. In Mep2, the CTR has moved away and makes relatively few contacts with the main body of the transporter, generating a more elongated protein (Figs 1 and 4). By contrast, in the structures of bacterial proteins, the CTR is docked tightly onto the N-terminal half of the transporters (corresponding to TM1-5), resulting in a more compact structure. This is illustrated by the positions of the five universally conserved residues within the CTR, that is, Arg415(370), Glu421(376), Gly424(379), Asp426(381) and Tyr 435(390) in CaMep2(Amt-1) (Fig. 2). These residues include those of the ‘ExxGxD' motif, which when mutated generate inactive transporters. In Amt-1 and other bacterial ammonium transporters, these CTR residues interact with residues within the N-terminal half of the protein. On one side, the Tyr390 hydroxyl in Amt-1 is hydrogen bonded with the side chain of the conserved His185 at the C-terminal end of loop ICL3. At the other end of ICL3, the backbone carbonyl groups of Gly172 and Lys173 are hydrogen bonded to the side chain of Arg370. Similar interactions were also modelled in the active, non-phosphorylated plant AtAmt-1;1 structure (for example, Y467-H239 and D458-K71). The result of these interactions is that the CTR ‘hugs' the N-terminal half of the transporters (Fig. 4). Also noteworthy is Asp381, the side chain of which interacts strongly with the positive dipole on the N-terminal end of TM2. This interaction in the centre of the protein may be particularly important to stabilize the open conformations of ammonium transporters. In the Mep2 structures, none of the interactions mentioned above are present. Phosphorylation target site is at the periphery of Mep2 Recently Boeckstaens et al. provided evidence that Ser457 in ScMep2 (corresponding to Ser453 in CaMep2) is phosphorylated by the TORC1 effector kinase Npr1 under nitrogen-limiting conditions. In the absence of Npr1, plasmid-encoded WT Mep2 in a S. cerevisiae mep1-3Δ strain (triple mepΔ) does not allow growth on low concentrations of ammonium, suggesting that the transporter is inactive (Fig. 3 and Supplementary Fig. 1). Conversely, the phosphorylation-mimicking S457D variant is active both in the triple mepΔ background and in a triple mepΔ npr1Δ strain (Fig. 3). Mutation of other potential phosphorylation sites in the CTR did not support growth in the npr1Δ background. Collectively, these data suggest that phosphorylation of Ser457 opens the Mep2 channel to allow ammonium uptake. Ser457 is located in a part of the CTR that is conserved in a subgroup of Mep2 proteins, but which is not present in bacterial proteins (Fig. 2). This segment (residues 450–457 in ScMep2 and 446–453 in CaMep2) was dubbed an autoinhibitory (AI) region based on the fact that its removal generates an active transporter in the absence of Npr1 (Fig. 3). Where is the AI region and the Npr1 phosphorylation site located? Our structures reveal that surprisingly, the AI region is folded back onto the CTR and is not located near the centre of the trimer as expected from the bacterial structures (Fig. 4). The AI region packs against the cytoplasmic ends of TM2 and TM4, physically linking the main body of the transporter with the CTR via main chain interactions and side-chain interactions of Val447, Asp449, Pro450 and Arg452 (Fig. 6). The AI regions have very similar conformations in CaMep2 and ScMep2, despite considerable differences in the rest of the CTR (Fig. 6). Strikingly, the Npr1 target serine residue is located at the periphery of the trimer, far away (∼30 Å) from any channel exit (Fig. 6). Despite its location at the periphery of the trimer, the electron density for the serine is well defined in both Mep2 structures and corresponds to the non-phosphorylated state (Fig. 6). This makes sense since the proteins were expressed in rich medium and confirms the recent suggestion by Boeckstaens et al. that the non-phosphorylated form of Mep2 corresponds to the inactive state. For ScMep2, Ser457 is the most C-terminal residue for which electron density is visible, indicating that the region beyond Ser457 is disordered. In CaMep2, the visible part of the sequence extends for two residues beyond Ser453 (Fig. 6). The peripheral location and disorder of the CTR beyond the kinase target site should facilitate the phosphorylation by Npr1. The disordered part of the CTR is not conserved in ammonium transporters (Fig. 2), suggesting that it is not important for transport. Interestingly, a ScMep2 457Δ truncation mutant in which a His-tag directly follows Ser457 is highly expressed but has low activity (Fig. 3 and Supplementary Fig. 1b), suggesting that the His-tag interferes with phosphorylation by Npr1. The same mutant lacking the His-tag has WT properties (Supplementary Fig. 1b), confirming that the region following the phosphorylation site is dispensable for function. Mep2 lacking the AI region is conformationally heterogeneous Given that Ser457/453 is far from any channel exit (Fig. 6), the crucial question is how phosphorylation opens the Mep2 channel to generate an active transporter. Boeckstaens et al. proposed that phosphorylation does not affect channel activity directly, but instead relieves inhibition by the AI region. The data behind this hypothesis is the observation that a ScMep2 449-485Δ deletion mutant lacking the AI region is highly active in MA uptake both in the triple mepΔ and triple mepΔ npr1Δ backgrounds, implying that this Mep2 variant has a constitutively open channel. We obtained a similar result for ammonium uptake by the 446Δ mutant (Fig. 3), supporting the data from Marini et al. We then constructed and purified the analogous CaMep2 442Δ truncation mutant and determined the crystal structure using data to 3.4 Å resolution. The structure shows that removal of the AI region markedly increases the dynamics of the cytoplasmic parts of the transporter. This is not unexpected given the fact that the AI region bridges the CTR and the main body of Mep2 (Fig. 6). Density for ICL3 and the CTR beyond residue Arg415 is missing in the 442Δ mutant, and the density for the other ICLs including ICL1 is generally poor with visible parts of the structure having high B-factors (Fig. 7). Interestingly, however, the Tyr49-His342 hydrogen bond that closes the channel in the WT protein is still present (Fig. 7 and Supplementary Fig. 2). Why then does this mutant appear to be constitutively active? We propose two possibilities. The first one is that the open state is disfavoured by crystallization because of lower stability or due to crystal packing constraints. The second possibility is that the Tyr–His hydrogen bond has to be disrupted by the incoming substrate to open the channel. The latter model would fit well with the NH3/H+ symport model in which the proton is relayed by the twin-His motif. The importance of the Tyr–His hydrogen bond is underscored by the fact that its removal in the ScMep2 Y53A mutant results in a constitutively active transporter (Fig. 3). Phosphorylation causes a conformational change in the CTR Do the Mep2 structures provide any clues regarding the potential effect of phosphorylation? The side-chain hydroxyl of Ser457/453 is located in a well-defined electronegative pocket that is solvent accessible (Fig. 6). The closest atoms to the serine hydroxyl group are the backbone carbonyl atoms of Asp419, Glu420 and Glu421, which are 3–4 Å away. We therefore predict that phosphorylation of Ser453 will result in steric clashes as well as electrostatic repulsion, which in turn might cause substantial conformational changes within the CTR. To test this hypothesis, we determined the structure of the phosphorylation-mimicking R452D/S453D protein (hereafter termed ‘DD mutant'), using data to a resolution of 2.4 Å. The additional mutation of the arginine preceding the phosphorylation site was introduced (i) to increase the negative charge density and make it more comparable to a phosphate at neutral pH, and (ii) to further destabilize the interactions of the AI region with the main body of the transporter (Fig. 6). The ammonium uptake activity of the S. cerevisiae version of the DD mutant is the same as that of WT Mep2 and the S453D mutant, indicating that the mutations do not affect transporter functionality in the triple mepΔ background (Fig. 3). Unexpectedly, the AI segment containing the mutated residues has only undergone a slight shift compared with the WT protein (Fig. 8 and Supplementary Fig. 3). By contrast, the conserved part of the CTR has undergone a large conformational change involving formation of a 12-residue-long α-helix from Leu427 to Asp438. In addition, residues Glu420-Leu423 including Glu421 of the ExxGxD motif are now disordered (Fig. 8 and Supplementary Fig. 3). Overall, ∼20 residues are affected by the introduced mutations. This is the first time a large conformational change has been observed in an ammonium transporter as a result of a mutation, and confirms previous hypotheses that phosphorylation causes structural changes in the CTR. To exclude the possibility that the additional R452D mutation is responsible for the observed changes, we also determined the structure of the ‘single D' S453D mutant. As shown in Supplementary Fig. 4, the consequence of the single D mutation is very similar to that of the DD substitution, with conformational changes and increased dynamics confined to the conserved part of the CTR (Supplementary Fig. 4). To supplement the crystal structures, we also performed modelling and MD studies of WT CaMep2, the DD mutant and phosphorylated protein (S453J). In the WT structure, the acidic residues Asp419, Glu420 and Glu421 are within hydrogen bonding distance of Ser453. After 200 ns of MD simulation, the interactions between these residues and Ser453 remain intact. The protein backbone has an average r.m.s.d. of only ∼3 Å during the 200-ns simulation, indicating that the protein is stable. There is flexibility in the side chains of the acidic residues so that they are able to form stable hydrogen bonds with Ser453. In particular, persistent hydrogen bonds are observed between the Ser453 hydroxyl group and the acidic group of Glu420, and also between the amine group of Ser453 and the backbone carbonyl of Glu420 (Supplementary Fig. 5). The DD mutant is also stable during the simulations, but the average backbone r.m.s.d of ∼3.6 Å suggests slightly more conformational flexibility than WT. As the simulation proceeds, the side chains of the acidic residues move away from Asp452 and Asp453, presumably to avoid electrostatic repulsion. For example, the distance between the Asp453 acidic oxygens and the Glu420 acidic oxygens increases from ∼7 to \u003e22 Å after 200 ns simulations, and thus these residues are not interacting. The protein is structurally stable throughout the simulation with little deviation in the other parts of the protein. Finally, the S453J mutant is also stable throughout the 200-ns simulation and has an average backbone deviation of ∼3.8 Å, which is similar to the DD mutant. The movement of the acidic residues away from Arg452 and Sep453 is more pronounced in this simulation in comparison with the movement away from Asp452 and Asp453 in the DD mutant. The distance between the phosphate of Sep453 and the acidic oxygen atoms of Glu420 is initially ∼11 Å, but increases to \u003e30 Å after 200 ns. The short helix formed by residues Leu427 to Asp438 unravels during the simulations to a disordered state. The remainder of the protein is not affected (Supplementary Fig. 5). Thus, the MD simulations support the notion from the crystal structures that phosphorylation generates conformational changes in the conserved part of the CTR. However, the conformational changes for the phosphomimetic mutants in the crystals are confined to the CTR (Fig. 8), and the channels are still closed (Supplementary Fig. 2). One possible explanation is that the mutants do not accurately mimic a phosphoserine, but the observation that the S453D and DD mutants are fully active in the absence of Npr1 suggests that the mutations do mimic the effect of phosphorylation (Fig. 3). The fact that the S453D structure was obtained in the presence of 10 mM ammonium ions suggests that the crystallization process favours closed states of the Mep2 channels. Discussion Knowledge about ammonium transporter structure has been obtained from experimental and theoretical studies on bacterial family members. In addition, a number of biochemical and genetic studies are available for bacterial, fungal and plant proteins. These efforts have advanced our knowledge considerably but have not yet yielded atomic-level answers to several important mechanistic questions, including how ammonium transport is regulated in eukaryotes and the mechanism of ammonium signalling. In Arabidopsis thaliana Amt-1;1, phosphorylation of the CTR residue T460 under conditions of high ammonium inhibits transport activity, that is, the default (non-phosphorylated) state of the plant transporter is open. Interestingly, phosphomimetic mutations introduced into one monomer inactivate the entire trimer, indicating that (i) heterotrimerization occurs and (ii) the CTR mediates allosteric regulation of ammonium transport activity via phosphorylation. Owing to the lack of structural information for plant AMTs, the details of channel closure and inter-monomer crosstalk are not yet clear. Contrasting with the plant transporters, the inactive states of Mep2 proteins under conditions of high ammonium are non-phosphorylated, with channels that are closed on the cytoplasmic side. The reason why similar transporters such as A. thaliana Amt-1;1 and Mep2 are regulated in opposite ways by phosphorylation (inactivation in plants and activation in fungi) is not known. In fungi, preventing ammonium entry via channel closure in ammonium transporters would be one way to alleviate ammonium toxicity, in addition to ammonium excretion via Ato transporters and amino-acid secretion. By determining the first structures of closed ammonium transporters and comparing those structures with the permanently open bacterial proteins, we demonstrate that Mep2 channel closure is likely due to movements of the CTR and ICL1 and ICL3. More specifically, the close interactions between the CTR and ICL1/ICL3 present in open transporters are disrupted, causing ICL3 to move outwards and block the channel (Figs 4 and 9a). In addition, ICL1 has shifted inwards to contribute to the channel closure by engaging His2 from the twin-His motif via hydrogen bonding with a highly conserved tyrosine hydroxyl group. Upon phosphorylation by the Npr1 kinase in response to nitrogen limitation, the region around the conserved ExxGxD motif undergoes a conformational change that opens the channel (Fig. 9). Importantly, the structural similarities in the TM parts of Mep2 and AfAmt-1 (Fig. 5a) suggest that channel opening/closure does not require substantial changes in the residues lining the channel. How exactly the channel opens and whether opening is intra-monomeric are still open questions; it is possible that the change in the CTR may disrupt its interactions with ICL3 of the neighbouring monomer (Fig. 9b), which could result in opening of the neighbouring channel via inward movement of its ICL3. Owing to the crosstalk between monomers, a single phosphorylation event might lead to opening of the entire trimer, although this has not yet been tested (Fig. 9b). Whether or not Mep2 channel opening requires, in addition to phosphorylation, disruption of the Tyr–His2 interaction by the ammonium substrate is not yet clear. Is our model for opening and closing of Mep2 channels valid for other eukaryotic ammonium transporters? Our structural data support previous studies and clarify the central role of the CTR and cytoplasmic loops in the transition between closed and open states. However, even the otherwise highly similar Mep2 proteins of S. cerevisiae and C. albicans have different structures for their CTRs (Fig. 1 and Supplementary Fig. 6). In addition, the AI region of the CTR containing the Npr1 kinase site is conserved in only a subset of fungal transporters, suggesting that the details of the structural changes underpinning regulation vary. Nevertheless, given the central role of absolutely conserved residues within the ICL1-ICL3-CTR interaction network (Fig. 4), we propose that the structural basics of fungal ammonium transporter activation are conserved. The fact that Mep2 orthologues of distantly related fungi are fully functional in ammonium transport and signalling in S. cerevisiae supports this notion. It should also be noted that the tyrosine residue interacting with His2 is highly conserved in fungal Mep2 orthologues, suggesting that the Tyr–His2 hydrogen bond might be a general way to close Mep2 proteins. With regards to plant AMTs, it has been proposed that phosphorylation at T460 generates conformational changes that would close the neighbouring pore via the C terminus. This assumption was based partly on a homology model for Amt-1;1 based on the (open) archaebacterial AfAmt-1 structure, which suggested that the C terminus of Amt-1;1 would extend further to the neighbouring monomer. Our Mep2 structures show that this assumption may not be correct (Fig. 4 and Supplementary Fig. 6). In addition, the considerable differences between structurally resolved CTR domains means that the exact environment of T460 in Amt-1;1 is also not known (Supplementary Fig. 6). Based on the available structural information, we consider it more likely that phosphorylation-mediated pore closure in Amt-1;1 is intra-monomeric, via disruption of the interactions between the CTR and ICL1/ICL3 (for example, Y467-H239 and D458-K71). There is generally no equivalent for CaMep2 Tyr49 in plant AMTs, indicating that a Tyr–His2 hydrogen bond as observed in Mep2 may not contribute to the closed state in plant transporters. We propose that intra-monomeric CTR-ICL1/ICL3 interactions lie at the basis of regulation of both fungal and plant ammonium transporters; close interactions generate open channels, whereas the lack of ‘intra-' interactions leads to inactive states. The need to regulate in opposite ways may be the reason why the phosphorylation sites are in different parts of the CTR, that is, centrally located close to the ExxGxD motif in AMTs and peripherally in Mep2. In this way, phosphorylation can either lead to channel closing (in the case of AMTs) or channel opening in the case of Mep2. Our model also provides an explanation for the observation that certain mutations within the CTR completely abolish transport activity. An example of an inactivating residue is the glycine of the ExxGxD motif of the CTR. Mutation of this residue (G393 in EcAmtB; G456 in AtAmt-1;1) inactivates transporters as diverse as Escherichia coli AmtB and A. thaliana Amt-1;1 (refs). Such mutations likely cause structural changes in the CTR that prevent close contacts between the CTR and ICL1/ICL3, thereby stabilizing a closed state that may be similar to that observed in Mep2. Regulation and modulation of membrane transport by phosphorylation is known to occur in, for example, aquaporins and urea transporters, and is likely to be a common theme for eukaryotic channels and transporters. Recently, phosphorylation was also shown to modulate substrate affinity in nitrate transporters. With respect to ammonium transport, phosphorylation has thus far only been shown for A. thaliana AMTs and for S. cerevisiae Mep2 (refs). However, the absence of GlnK proteins in eukaryotes suggests that phosphorylation-based regulation of ammonium transport may be widespread. Nevertheless, as discussed above, considerable differences may exist between different species. With respect to Mep2-mediated signalling to induce pseudohyphal growth, two models have been put forward as to how this occurs and why it is specific to Mep2 proteins. In one model, signalling is proposed to depend on the nature of the transported substrate, which might be different in certain subfamilies of ammonium transporters (for example, Mep1/Mep3 versus Mep2). For example, NH3 uniport or symport of NH3/H+ might result in changes in local pH, but NH4+ uniport might not, and this difference might determine signalling. In the other model, signalling is thought to require a distinct conformation of the Mep2 transporter occurring during the transport cycle. While the current study does not specifically address the mechanism of signalling underlying pseudohyphal growth, our structures do show that Mep2 proteins can assume different conformations. It is clear that ammonium transport across biomembranes remains a fascinating and challenging field in large part due to the unique properties of the substrate. Our Mep2 structural work now provides a foundation for future studies to uncover the details of the structural changes that occur during eukaryotic ammonium transport and signaling, and to assess the possibility to utilize small molecules to shut down ammonium sensing and downstream signalling pathways in pathogenic fungi. Methods Mep2 overexpression and purification Ammonium transporter genes were amplified from genomic DNA or cDNA by PCR (Phusion, New England Biolabs). In both ScMEP2 and CaMEP2, Asn4 was replaced by a glutamine to prevent glycosylation. In order to allow transformation of yeast by recombination, the following primer extensions were used: forward 5′-GAAAAAACCCCGGATTCTAGAACTAGTGGATCCTCC-3′ and reverse 5′-TGACTCGAGTTATGCACCGTGGTGGTGATGGTGATG-3′. These primers result in a construct that lacks the cleavable N- and C-terminal tags present in the original vector, and replaces these with a C-terminal hexa-histidine tag. Recombination in yeast strain W303 pep4Δ was carried out using ∼50–100 ng of SmaI-digested vector 83νΔ (ref.) and at least a fourfold molar excess of PCR product via the lithium acetate method. Transformants were selected on SCD -His plates incubated at 30 °C. Construction of mutant CaMEP2 genes was done using the Q5 site-directed mutagenesis kit (NEB) per manufacturer's instructions. Three CaMep2 mutants were made for crystallization: the first mutant is a C-terminal truncation mutant 442Δ, lacking residues 443–480 including the AI domain. The second mutant, R452D/S453D, mimics the protein phosphorylated at Ser453. Given that phosphate is predominantly charged −2 at physiological pH, we introduced the second aspartate residue for Arg452. However, we also constructed the ‘single D', S453D CaMep2 variant. For expression, cells were grown in shaker flasks at 30 °C for ∼24 h in synthetic minimal medium lacking histidine and with 1% (w/v) glucose to a typical OD600 of 6–8. Cells were subsequently spun down for 15 min at 4,000g and resuspended in YP medium containing 1.5% (w/v) galactose, followed by another 16–20 h growth at 30 °C/160 r.p.m. and harvesting by centrifugation. Final OD600 values typically reached 18–20. Cells were lysed by bead beating (Biospec) for 5 × 1 min with 1 min intervals on ice, or by 1–2 passes through a cell disrupter operated at 35,000 p.s.i. (TS-Series 0.75 kW; Constant Systems). Membranes were collected from the suspension by centrifugation at 200,000g for 90 min (45Ti rotor; Beckmann Coulter). Membrane protein extraction was performed by homogenization in a 1:1 (w/w) mixture of dodecyl-β-D-maltoside and decyl-β-D-maltoside (DDM/DM) followed by stirring at 4 °C overnight. Typically, 1 g (1% w/v) of total detergent was used for membranes from 2 l of cells. The membrane extract was centrifuged for 35 min at 200,000g and the supernatant was loaded onto a 10-ml Nickel column (Chelating Sepharose; GE Healthcare) equilibrated in 20 mM Tris/300 mM NaCl/0.2% DDM, pH 8. The column was washed with 15 column volumes buffer containing 30 mM imidazole and eluted in 3 column volumes with 250 mM imidazole. Proteins were purified to homogeneity by gel filtration chromatography in 10 mM HEPES/100 mM NaCl/0.05% DDM, pH 7–7.5. For polishing and detergent exchange, a second gel filtration column was performed using various detergents. In the case of ScMep2, diffracting crystals were obtained only with 0.05% decyl-maltose neopentyl glycol. For the more stable CaMep2 protein, we obtained crystals in, for example, nonyl-glucoside, decyl-maltoside and octyl-glucose neopentyl glycol. Proteins were concentrated to 7–15 mg ml−1 using 100 kDa cutoff centrifugal devices (Millipore), flash-frozen and stored at −80 °C before use. Crystallization and structure determination Crystallization screening trials by sitting drop vapour diffusion were set up at 4 and 20 °C using in-house screens and the MemGold 1 and 2 screens (Molecular Dimensions) with a Mosquito crystallization robot. Crystals were harvested directly from the initial trials or optimized by sitting or hanging drop vapour diffusion using larger drops (typically 2–3 μl total volume). Bar-shaped crystals for ScMep2 diffracting to 3.2 Å resolution were obtained from 50 mM 2-(N-morpholino)ethanesulfonic acid (MES)/0.2 M di-ammonium hydrogen phosphate/30% PEG 400, pH 6. They belong to space group P212121 and have nine molecules (three trimers) in the asymmetric unit (AU). Well-diffracting crystals for CaMep2 were obtained in space group P3 from 0.1 M MES/0.2 M lithium sulphate/20% PEG400, pH 6 (two molecules per AU). An additional crystal form in space group R3 was grown in 0.04 M Tris/0.04 M NaCl/27% PEG350 MME, pH 8 (one molecule per AU). Diffracting crystals for the phosporylation-mimicking CaMep2 DD mutant were obtained in space group P6322 from 0.1 M sodium acetate/15–20% PEG400, pH 5 (using decyl-maltoside as detergent; one molecule per AU), while S453D mutant crystals grew in 24% PEG400/0.05 M sodium acetate, pH 5.4/0.05 M magnesium acetate tetrahydrate/10 mM NH4Cl (space group R32; one molecule per AU). Finally, the 442Δ truncation mutant gave crystals under many different conditions, but most of these diffracted poorly or not at all. A reasonable low-resolution data set (3.4 Å resolution) was eventually obtained from a crystal grown in 24% PEG400/0.05 M sodium acetate/0.05 M magnesium acetate, pH 6.1 (space group R32). Diffraction data were collected at the Diamond Light Source and processed with XDS or HKL2000 (ref. ). For MR, a search model was constructed with Sculptor within Phenix, using a sequence alignment of ScMep2 with Archaeoglobus fulgidus Amt-1 (PDB ID 2B2H; ∼40% sequence identity to ScMep2). A clear solution with nine molecules (three trimers) in the AU was obtained using Phaser. The model was subsequently completed by iterative rounds of manual building within Coot followed by refinement within Phenix. The structures for WT CaMep2 were solved using the best-defined monomer of ScMep2 (60% sequence identity with CaMep2) in MR with Phaser, followed by automated model building within Phenix. Finally, the structures of the three mutant CaMep2 proteins were solved using WT CaMep2 as the search model. The data collection and refinement statistics for all six solved structures have been summarized in Supplementary Tables 1 and 2. Growth assays The S. cerevisiae haploid triple mepΔ strain (Σ1278b MATα mep1::LEU2 mep2::LEU2 mep3::G418 ura3-52) and triple mepΔ npr1Δ strain (Σ1278b MATα mep1::LEU2 mep2::LEU2 mep3::G418 npr1::NAT1 ura3-52) were generated by integrating the NAT1 resistance gene at one NPR1 locus in the diploid strain MLY131 (ref.), followed by isolation of individual haploid strains. Cells were grown in synthetic minimal medium with glucose (2%) as the carbon source and ammonium sulphate (1 mM) or glutamate (0.1%) as the nitrogen source. Yeast cells were transformed as described. All DNA sequences encoding epitope-tagged ScMep2 and its mutant derivatives were generated by PCR and homologous recombination using the vector pRS316 (ref. ). In each case, the ScMEP2 sequences included the ScMEP2 promoter (1 kb), the ScMEP2 terminator and sequences coding for a His-6 epitope at the C-terminal end of the protein. All Mep2-His fusions contain the N4Q mutation to prevent glycosylation of Mep2 (ref.). All newly generated plasmid inserts were verified by DNA sequencing. For growth assays, S. cerevisiae cells containing plasmids expressing ScMep2 or mutant derivatives were grown overnight in synthetic minimal glutamate medium, washed, spotted by robot onto solid agar plates and culture growth followed by time course photography. Images were then processed to quantify the growth of each strain over 3 days as described. Protein modelling The MODELLER (version 9.15) software package was used to build protein structures for MD simulations. This method was required to construct two complete protein models, the double mutant R452D/S453D (with the four missing residues from the X-ray structure added) and also the construct in which the mutation at position 452 is reverted to R, and D453 is replaced with a phosphoserine. The quality of these models was assessed using normalized Discrete Optimized Protein Energy (DOPE) values and the molpdf assessment function within the MODELLER package. The model R452D/S453D mutant has a molpdf assessment score of 1854.05, and a DOPE assessment score of -60920.55. The model of the S453J mutant has a molpdf assessment score of 1857.01 and a DOPE assessment score of −61032.15. MD simulations WT and model structures were embedded into a pre-equilibrated lipid bilayer composed of 512 dipalmitoylphosphatidylcholine lipids using the InflateGRO2 computer programme. The bilayers were then solvated with the SPC water model and counterions were added to achieve a charge neutral state. All simulations were performed with the GROMACS package (version 4.5.5), and the GROMOS96 43a1p force field. During simulation time, the temperature was maintained at 310 K using the Nosé-Hoover thermostat with a coupling constant of 0.5 ps. Pressure was maintained at 1 bar using semi-isotropic coupling with the Parrinello-Rahman barostat and a time constant of 5 ps. Electrostatic interactions were treated using the smooth particle mesh Ewald algorithm with a short-range cutoff of 0.9 nm. Van der Waals interactions were truncated at 1.4 nm with a long-range dispersion correction applied to energy and pressure. The neighbour list was updated every five steps. All bonds were constrained with the LINCS algorithm, so that a 2-fs time step could be applied throughout. The phospholipid parameters for the dipalmitoylphosphatidylcholine lipids were based on the work of Berger. The embedded proteins were simulated for 200 ns each; a repeat simulation was performed for each system with different initial velocities to ensure reproducibility. To keep the c.p.u. times within reasonable limits, all simulations were performed on Mep2 monomers. This is also consistent with previous simulations for E. coli AmtB. Additional information Accession codes: The atomic coordinates and the associated structure factors have been deposited in the Protein Data Bank (http:// www.pdbe.org) with accession codes 5AEX (ScMep2), 5AEZ(CaMep2; R3), 5AF1(CaMep2; P3), 5AID(CaMep2; 442D), 5AH3 (CaMep2; R452D/S453D) and 5FUF (CaMep2; S453D). How to cite this article: van den Berg, B. et al. Structural basis for Mep2 ammonium transceptor activation by phosphorylation. Nat. Commun. 7:11337 doi: 10.1038/ncomms11337 (2016). Supplementary Material The eukaryotic plasma membrane as a nutrient-sensing device Nutrient sensing and signaling in the yeast Saccharomyces cerevisiae The transporter classification database A family of ammonium transporters in Saccharomyces cerevisiae A split-ubiquitin two-hybrid screen for proteins physically interacting with the yeast amino acid transceptor Gap1 and ammonium transceptor Mep2 The MEP2 ammonium permease regulates pseudohyphal differentiation in Saccharomyces cerevisiae Differential activation of ammonium transporters during the accumulation of ammonia by Colletotrichum gloeosporioides and its effect on appressoria formation andpathogenicity A Mep2-dependent transcriptional profile links permease function to gene expression during pseudohyphal growth in Saccharomyces cerevisiae The Mep2p ammonium permease controls nitrogen starvation-induced filamentous growth in Candida albicans Distinct transport mechanisms in yeast ammonium transport/sensor proteins of the Mep/Amt/Rh family and impact on filamentation The yeast ammonium transport protein Mep2 and its positive regulator, the Npr1 kinase, play an important role in normal and pseudohyphal growth on various nitrogen media through retrieval of excreted ammonium Function of human Rh based on structure of RhCG at 2.1A Mechanism of ammonia transport by Amt/MEP/Rh: structure of AmtB at 1.35A Crystal structure of the archaeal ammonium transporter Amt-1 from Archaeoglobus fulgidus An unusual twin-his arrangement in the pore of ammonia channels is essential for substrate conductance Ammonium transporters achieve charge transfer by fragmenting their substrate Ammonium transport proteins with changes in one of the conserved pore histidines have different performance in ammonia and methylamine conduction Molecular dynamics simulations on the Escherichia coli ammonia channel protein AmtB: mechanism of ammonia/ammonium transport Periplasmic vestibule plays an important role for solute recruitment, selectivity, and gating in the Rh/Amt/MEP superfamily The mechanism of ammonia transport based on the crystal structure of AmtB of Escherichia coli Different hydration patterns in the pores of AmtB and RhCG could determine their transport mechanisms Nitrogen assimilation in Escherichia coli: putting molecular data into a systems perspective Direct observation of electrogenic NH4(+) transport in ammonium transport (Amt) proteins Uncoupling of ionic currents from substrate transport in the plant ammonium transporter AtAMT1;2 Molecular mechanism of acute ammonia toxicity: role of NMDA receptors The glnKamtB operon. A conserved gene pair in prokaryotes Inhibitory complex of the transmembrane ammonia channel, AmtB, and the cytosolic regulatory protein, GlnK, at 1.96A In vitro analysis of the Escherichia coli AmtB-GlnK complex reveals a stoichiometric interaction and sensitivity to ATP and 2-oxoglutarate Adjusting ammonium uptake via phosphorylation The TORC1 effector kinase Npr1 fine tunes the inherent activity of the Mep2 ammonium transport protein N- terminal cysteines affect oligomer stability of the allosterically regulated ammonium transporter LeAMT1;1 The conserved carboxy-terminal region of the ammonia channel AmtB plays a critical role in channel function A cytosolic trans-activation domain essential for ammonium uptake Allosteric regulation of transport activity by heterotrimerization of Arabidopsis ammonium transporter complexes in vivo Ammonium toxicity and potassium limitation in yeast Isolation and characterization from pathogenic fungi of genes encoding ammonium permeases and their roles in dimorphism Impact of ammonium permeases mepA, mepB, and mepC on nitrogen-regulated secondary metabolism in Fusarium fujikuroi Molecular characterization, function and regulation of ammonium transporters (Amt) and ammonium-metabolizing enzymes (GS, NADP-GDH) in the ectomycorrhizal fungus Hebeloma cylindrosporum Structural insights into eukaryotic aquaporin regulation Expression of urea transporters and their regulation Molecular basis of nitrate uptake by the plant nitrate transporter NRT1.1 Overexpression and purification of integral membrane proteins in yeast XDS Processing of X-ray diffraction data collected in oscillation mode Towards automated crystallographic structure refinement with phenix.refine Likelihood-enhanced fast translation functions Coot: model-building tools for molecular graphics High efficiency transformation of intact yeast cells using single stranded nucleic acids as a carrier Plasmid construction by homologous recombination in yeast In vivo N-glycosylation of the Mep2 high-affinity ammonium transporter of Saccharomyces cerevisiae reveals an extracytosolic N-terminus Quantitative fitness analysis shows that NMD proteins and many other protein complexes suppress or enhance distinct telomere cap defects Comparative protein structure modeling of genes and genomes LAMBADA and InflateGRO2: efficient membrane alignment and insertion of membrane proteins for molecular dynamics simulations  GROMACS: fast, flexible, and free A unified formulation of the constant temperature molecular dynamics methods Canonical dynamics: equilibrium phase-space distributions Polymorphic transitions in single crystals: a new molecular dynamics method A smooth particle mesh Ewald method LINCS: a linear constraint solver for molecular simulations Molecular dynamics simulations of a fluid bilayer of dipalmitoylphosphatidylcholine at full hydration, constant pressure, and constant temperature  The PyMOL Molecular Graphics System. version 1.7.4 (Schrödinger, LLC). Author contributions B.v.d.B. performed the experiments related to Mep2 structure determination, designed research and wrote the paper. A.C. performed ammonium growth experiments of Mep variants. D.J. and S.K. performed modelling studies and MD simulations. A.B. collected the X-ray synchrotron data and maintained the Newcastle Structural Biology Laboratory. J.C.R. designed research related to the S. cerevisiae growth assays. X-ray crystal structures of Mep2 transceptors. (a) Monomer cartoon models viewed from the side for (left) A.\nfulgidus Amt-1 (PDB ID 2B2H), S. cerevisiae Mep2 (middle) and\nC. albicans Mep2 (right). The cartoons are in rainbow\nrepresentation. The region showing ICL1 (blue), ICL3 (green) and the CTR\n(red) is boxed for comparison. (b) CaMep2 trimer viewed from the\nintracellular side (right). One monomer is coloured as in a and one\nmonomer is coloured by B-factor (blue, low; red; high). The CTR is boxed.\n(c) Overlay of ScMep2 (grey) and CaMep2 (rainbow), illustrating\nthe differences in the CTRs. All structure figures were generated with\nPymol. Sequence conservation in ammonium transporters. ClustalW alignment of CaMep2, ScMep2, A. fulgidus Amt-1, E.\ncoli AmtB and A. thaliana Amt-1;1. The secondary structure\nelements observed for CaMep2 are indicated, with the numbers corresponding\nto the centre of the TM segment. Important regions are labelled. The\nconserved RxK motif in ICL1 is boxed in blue, the ER motif in ICL2 in cyan,\nthe conserved ExxGxD motif of the CTR in red and the AI region in yellow.\nColoured residues are functionally important and correspond to those of the\nPhe gate (blue), the binding site Trp residue (magenta) and the twin-His\nmotif (red). The Npr1 kinase site in the AI region is highlighted pink. The\ngrey sequences at the C termini of CaMep2 and ScMep2 are not visible in the\nstructures and are likely disordered. Growth of ScMep2 variants on low ammonium medium. (a) The triple mepΔ strain (black) and triple\nmepΔ npr1Δ strain (grey) containing plasmids\nexpressing WT and variant ScMep2 were grown on minimal medium containing\n1 mM ammonium sulphate. The quantified cell density reflects\nlogarithmic growth after 24 h. Error bars are the s.d. for three\nreplicates of each strain (b) The strains used in a were also\nserially diluted and spotted onto minimal agar plates containing glutamate\n(0.1%) or ammonium sulphate (1 mM), and grown for 3 days at\n30 °C. Structural differences between Mep2 and bacterial ammonium\ntransporters. (a) ICL1 in AfAmt-1 (light blue) and CaMep2 (dark blue), showing\nunwinding and inward movement in the fungal protein. (b) Stereo\ndiagram viewed from the cytosol of ICL1, ICL3 (green) and the CTR (red) in\nAfAmt-1 (light colours) and CaMep2 (dark colours). The side chains of\nresidues in the RxK motif as well as those of Tyr49 and His342 are labelled.\nThe numbering is for CaMep2. (c) Conserved residues in ICL1-3 and the\nCTR. Views from the cytosol for CaMep2 (left) and AfAmt-1, highlighting the\nlarge differences in conformation of the conserved residues in ICL1 (RxK\nmotif; blue), ICL2 (ER motif; cyan), ICL3 (green) and the CTR (red). The\nlabelled residues are analogous within both structures. In b and\nc, the centre of the trimer is at top. Channel closures in Mep2. (a) Stereo superposition of AfAmt-1 and CaMep2 showing the residues of\nthe Phe gate, His2 of the twin-His motif and the tyrosine residue Y49 in TM1\nthat forms a hydrogen bond with His2 in CaMep2. (b) Surface views\nfrom the side in rainbow colouring, showing the two-tier channel block\n(indicated by the arrows) in CaMep2. The Npr1 kinase target Ser453 is dephosphorylated and located in an\nelectronegative pocket. (a) Stereoviews of CaMep2 showing 2Fo–Fc\nelectron density (contoured at 1.0 σ) for CTR residues\nAsp419-Met422 and for Tyr446-Thr455 of the AI region. For clarity, the\nresidues shown are coloured white, with oxygen atoms in red and nitrogen\natoms in blue. The phosphorylation target residue Ser453 is labelled in\nbold. (b) Overlay of the CTRs of ScMep2 (grey) and CaMep2 (green),\nshowing the similar electronegative environment surrounding the\nphosphorylation site (P). The AI regions are coloured magenta. (c)\nCytoplasmic view of the Mep2 trimer indicating the large distance between\nSer453 and the channel exits (circles; Ile52 lining the channel exit is\nshown). Effect of removal of the AI region on Mep2 structure. (a) Side views for WT CaMep2 (left) and the truncation mutant\n442Δ (right). The latter is shown as a putty model according to\nB-factors to illustrate the disorder in the protein on the cytoplasmic side.\nMissing regions are labelled. (b) Stereo superpositions of WT CaMep2\nand the truncation mutant. 2Fo–Fc electron\ndensity (contoured at 1.0 σ) for residues Tyr49 and His342 is\nshown for the truncation mutant. Phosphorylation causes conformational changes in the CTR. (a) Cytoplasmic view of the DD mutant trimer, with WT CaMep2\nsuperposed in grey for one of the monomers. The arrow indicates the\nphosphorylation site. The AI region is coloured magenta. (b) Monomer\nside-view superposition of WT CaMep2 and the DD mutant, showing the\nconformational change and disorder around the ExxGxD motif. Side chains for\nresidues 452 and 453 are shown as stick models. Schematic model for phosphorylation-based regulation of Mep2 ammonium\ntransporters. (a) In the closed, non-phosphorylated state (i), the CTR (magenta) and\nICL3 (green) are far apart with the latter blocking the intracellular\nchannel exit (indicated with a hatched circle). Upon phosphorylation and\nmimicked by the CaMep2 S453D and DD mutants (ii), the region around the\nExxGxD motif undergoes a conformational change that results in the CTR\ninteracting with the inward-moving ICL3, opening the channel (full circle)\n(iii). The arrows depict the movements of important structural elements. The\nopen-channel Mep2 structure is represented by archaebacterial Amt-1 and\nshown in lighter colours consistent with Fig. 4. As\ndiscussed in the text, similar structural arrangements may occur in plant\nAMTs. In this case however, the open channel corresponds to the\nnon-phosphorylated state; phosphorylation breaks the CTR–ICL3\ninteractions leading to channel closure. (b) Model based on AMT\ntransporter analogy showing how phosphorylation of a\nMep2 monomer might allosterically open channels in the entire trimer via\ndisruption of the interactions between the CTR and ICL3 of a neighbouring\nmonomer 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+[{"sourceid":"4854314","sourcedb":"","project":"","target":"","text":"RNA protects a nucleoprotein complex against radiation damage Systematic analysis of radiation damage within a protein–RNA complex over a large dose range (1.3–25 MGy) reveals significant differential susceptibility of RNA and protein. A new method of difference electron-density quantification is presented. Radiation damage during macromolecular X-ray crystallographic data collection is still the main impediment for many macromolecular structure determinations. Even when an eventual model results from the crystallographic pipeline, the manifestations of radiation-induced structural and conformation changes, the so-called specific damage, within crystalline macromolecules can lead to false interpretations of biological mechanisms. Although this has been well characterized within protein crystals, far less is known about specific damage effects within the larger class of nucleoprotein complexes. Here, a methodology has been developed whereby per-atom density changes could be quantified with increasing dose over a wide (1.3–25.0 MGy) range and at higher resolution (1.98 Å) than the previous systematic specific damage study on a protein–DNA complex. Specific damage manifestations were determined within the large trp RNA-binding attenuation protein (TRAP) bound to a single-stranded RNA that forms a belt around the protein. Over a large dose range, the RNA was found to be far less susceptible to radiation-induced chemical changes than the protein. The availability of two TRAP molecules in the asymmetric unit, of which only one contained bound RNA, allowed a controlled investigation into the exact role of RNA binding in protein specific damage susceptibility. The 11-fold symmetry within each TRAP ring permitted statistically significant analysis of the Glu and Asp damage patterns, with RNA binding unexpectedly being observed to protect these otherwise highly sensitive residues within the 11 RNA-binding pockets distributed around the outside of the protein molecule. Additionally, the method enabled a quantification of the reduction in radiation-induced Lys and Phe disordering upon RNA binding directly from the electron density. Introduction    With the wide use of high-flux third-generation synchrotron sources, radiation damage (RD) has once again become a dominant reason for the failure of structure determination using macromolecular crystallography (MX) in experiments conducted both at room temperature and under cryocooled conditions (100 K). Significant progress has been made in recent years in understanding the inevitable manifestations of X-ray-induced RD within protein crystals, and there is now a body of literature on possible strategies to mitigate the effects of RD (e.g. Zeldin, Brockhauser et al., 2013; Bourenkov \u0026 Popov, 2010). However, there is still no general consensus within the field on how to minimize RD during MX data collection, and debates on the dependence of RD progression on incident X-ray energy (Shimizu et al., 2007; Liebschner et al., 2015) and the efficacy of radical scavengers (Allan et al., 2013) have yet to be resolved. RD manifests in two forms. Global radiation damage is observed within reciprocal space as the overall decay of the summed intensity of reflections detected within the diffraction pattern as dose increases (Garman, 2010; Murray \u0026 Garman, 2002). Dose is defined as the absorbed energy per unit mass of crystal in grays (Gy; 1 Gy = 1 J kg−1), and is the metric against which damage progression should be monitored during MX data collection, as opposed to time. At 100 K, an experimental dose limit of 30 MGy has been recommended as an upper limit beyond which the biological information derived from any macromolecular crystal may be compromised (Owen et al., 2006).  Specific radiation damage (SRD) is observed in the real-space electron density, and has been detected at much lower doses than any observable decay in the intensity of reflections. Indeed, the C—Se bond in selenomethionine, the stability of which is key for the success of experimental phasing methods, can be cleaved at a dose as low as 2 MGy for a crystal maintained at 100 K (Holton, 2007). SRD has been well characterized in a large range of proteins, and is seen to follow a reproducible order: metallo-centre reduction, disulfide-bond cleavage, acidic residue decarboxylation and methionine methylthio cleavage (Ravelli \u0026 McSweeney, 2000; Burmeister, 2000; Weik et al., 2000; Yano et al., 2005). Furthermore, damage susceptibility within each residue type follows a preferential ordering influenced by a combination of local environment factors (solvent accessibility, conformational strain, proximity to active sites/high X-ray cross-section atoms; Holton, 2009). Deconvoluting the individual roles of these parameters has been surprisingly challenging, with factors such as solvent accessibility currently under active investigation (Weik et al., 2000; Fioravanti et al., 2007; Gerstel et al., 2015). There are a number of cases where SRD manifestations have compromised the biological information extracted from MX-determined structures at much lower doses than the recommended 30 MGy limit, leading to false structural interpretations of protein mechanisms. Active-site residues appear to be particularly susceptible, particularly for photosensitive proteins and in instances where chemical strain is an intrinsic feature of the reaction mechanism. For instance, structure determination of the purple membrane protein bacterio­rhodopsin required careful corrections for radiation-induced structural changes before the correct photosensitive intermediate states could be isolated (Matsui et al., 2002). The significant chemical strain required for catalysis within the active site of phosphoserine aminotransferase has been observed to diminish during X-ray exposure (Dubnovitsky et al., 2005). Since the majority of SRD studies to date have focused on proteins, much less is known about the effects of X-ray irradiation on the wider class of crystalline nucleoprotein complexes or how to correct for such radiation-induced structural changes. Understanding RD to such complexes is crucial, since DNA is rarely naked within a cell, instead dynamically interacting with proteins, facilitating replication, transcription, modification and DNA repair. As of early 2016, \u003e5400 nucleoprotein complex structures have been deposited within the PDB, with 91% solved by MX. It is essential to understand how these increasingly complex macromolecular structures are affected by the radiation used to solve them. Nucleoproteins also represent one of the main targets of radiotherapy, and an insight into the damage mechanisms induced by X-ray irradiation could inform innovative treatments. When a typical macromolecular crystal is irradiated with ionizing X-rays, each photoelectron produced via interactions with both the macromolecule (direct damage) and solvent (indirect damage) can induce cascades of up to 500 secondary low-energy electrons (LEEs) that are capable of inducing further ionizations. Investigations on sub-ionization-level LEEs (0–15 eV) interacting with both dried and aqueous oligonucleotides (Alizadeh \u0026 Sanche, 2014; Simons, 2006) concluded that resonant electron attachment to DNA bases and the sugar-phosphate backbone could lead to the preferential cleavage of strong (∼4 eV, 385 kJ mol−1) sugar-phosphate C—O covalent bonds within the DNA backbone and then base-sugar N1—C bonds, eventually leading to single-strand breakages (SSBs; Ptasińska \u0026 Sanche, 2007). Electrons have been shown to be mobile at 77 K by electron spin resonance spectroscopy studies (Symons, 1997; Jones et al., 1987), with rapid electron quantum tunnelling and positive hole migration along the protein backbone and through stacked DNA bases indicated as a dominant mechanism by which oxidative and reductive damage localizes at distances from initial ionization sites at 100 K (O’Neill et al., 2002). The investigation of naturally forming nucleoprotein complexes circumvents the inherent challenges in making controlled comparisons of damage mechanisms between protein and nucleic acids crystallized separately. Recently, for a well characterized bacterial protein–DNA complex (C.Esp1396I; PDB entry 3clc; resolution 2.8 Å; McGeehan et al., 2008) it was concluded that over a wide dose range (2.1–44.6 MGy) the protein was far more susceptible to SRD than the DNA within the crystal (Bury et al., 2015). Only at doses above 20 MGy were precursors of phosphodiester-bond cleavage observed within AT-rich regions of the 35-mer DNA. For crystalline complexes such as C.Esp1396I, whether the protein is intrinsically more susceptible to X-ray-induced damage or whether the protein scavenges electrons to protect the DNA remains unclear in the absence of a non-nucleic acid-bound protein control obtained under exactly the same crystallization and data-collection conditions. To monitor the effects of nucleic acid binding on protein damage susceptibility, a crystal containing two protein molecules per asymmetric unit, only one of which was bound to RNA, is reported here (Fig. 1 ▸). Using newly developed methodology, we present a controlled SRD investigation at 1.98 Å resolution using a large (∼91 kDa) crystalline protein–RNA complex: trp RNA-binding attenuation protein (TRAP) bound to a 53 bp RNA sequence (GAGUU)10GAG (PDB entry 1gtf; Hopcroft et al., 2002). TRAP consists of 11 identical subunits assembled into a ring with 11-fold rotational symmetry. It binds with high affinity (K d ≃ 1.0 nM) to RNA segments containing 11 GAG/UAG triplets separated by two or three spacer nucleotides (Elliott et al., 2001) to regulate the transcription of tryptophan biosynthetic genes in Bacillus subtilis (Antson et al., 1999). In this structure, the bases of the G1-A2-G3 nucleotides form direct hydrogen bonds to the protein, unlike the U4-U5 nucleotides, which appear to be more flexible. Ten successive 1.98 Å resolution MX data sets were collected from the same TRAP–RNA crystal to analyse X-ray-induced structural changes over a large dose range (d 1 = 1.3 MGy to d 10 = 25.0 MGy). To avoid the previous necessity for visual inspection of electron-density maps to detect SRD sites, a computational approach was designed to quantify the electron-density change for each refined atom with increasing dose, thus providing a rapid systematic method for SRD study on such large multimeric complexes. By employing the high 11-fold structural symmetry within each TRAP macromolecule, this approach permitted a thorough statistical quantification of the RD effects of RNA binding to TRAP. Materials and methods    RNA synthesis and protein preparation    As previously described (Hopcroft et al., 2002), the 53-base RNA (GAGUU)10GAG was synthesized by in vitro transcription with T7 RNA polymerase and gel-purified. TRAP from B. stearothermophilus was overexpressed in Escherichia coli and purified. Crystallization    TRAP–RNA crystals were prepared using a previously established hanging-drop crystallization protocol (Antson et al., 1999). By using a 2:1 molar ratio of TRAP to RNA, crystals successfully formed from the protein–RNA complex (∼15 mg ml−1) in a solution containing 70 mM potassium phosphate pH 7.8 and 10 mM l-tryptophan. The reservoir consisted of 0.2 M potassium glutamate, 50 mM triethanol­amine pH 8.0, 10 mM MgCl2, 8–11% monomethyl ether PEG 2000. In order to accelerate crystallization, a further gradient was induced by adding 0.4 M KCl to the reservoir after 1.5 µl protein solution had been mixed with an equal volume of the reservoir solution. Wedge-shaped crystals of approximate length 70 µm (longest dimension) grew within 3 d and were vitrified and stored in liquid nitrogen immediately after growth. The cryosolution consisted of 12% monomethyl ether PEG 2000, 30 mM triethanolamine pH 8.0, 6 mM l-tryptophan, 0.1 M potassium glutamate, 35 mM potassium phosphate pH 7.8, 5 mM MgCl2 with 25% 2-methyl-2,4-pentanediol (MPD) included as a cryoprotectant. X-ray data collection    Data were collected at 100 K from a wedge-shaped TRAP–RNA crystal of approximate dimensions 70 × 20 × 40 µm (see Supplementary Fig. S2) on beamline ID14-4 at the ESRF using an incident wavelength of 0.940 Å (13.2 keV) and an ADSC Q315R mosaic CCD detector at 304.5 mm from the crystal throughout the data collection. The beam size was slitted to 0.100 mm (vertical) × 0.160 mm (horizontal), with a uniformly distributed profile, such that the crystal was completely bathed within the beam throughout data collection. Ten successive (1.98 Å resolution) 180° data sets (with Δφ = 1°) were collected over the same angular range from a TRAP–RNA crystal at 28.9% beam transmission. The TRAP–RNA macromolecule crystallized in space group C2, with unit-cell parameters a = 140.9, b = 110.9, c = 137.8 Å, α = γ = 90, β = 137.8° (the values quoted are for the first data set; see Supplementary Table S1 for subsequent values). For the first nine data sets the attenuated flux was recorded to be ∼5 × 1011 photons s−1. A beam refill took place immediately before data set 10, requiring a flux-scale factor increase of 1.42 to be applied, based on the ratio of observed relative intensity I D/I 1 at data set 10 to that extrapolated from data set 9. Dose calculation     RADDOSE-3D (Zeldin, Gerstel et al., 2013) was used to calculate the absorbed dose distribution during each data set (see input file; Supplementary Figs. S1 and S2). The crystal composition was calculated from the deposited TRAP–RNA structure (PDB entry 1gtf; Hopcroft et al., 2002). Crystal absorption coefficients were calculated in RADDOSE-3D using the concentration (mmol l−1) of solvent heavy elements from the crystallization conditions. The beam-intensity profile was modelled as a uniform (‘top-hat’) distribution. The diffraction-weighted dose (DWD) values (Zeldin, Brock­hauser et al., 2013) are given in Supplementary Table S1. Data processing and model refinement    Each data set was integrated using iMosflm (Leslie \u0026 Powell, 2007) and was scaled using AIMLESS (Evans \u0026 Murshudov, 2013; Winn et al., 2011) using the same 5% R free set of test reflections for each data set. To phase the structure obtained from the first data set, molecular replacement was carried out with Phaser (McCoy et al., 2007), using an identical TRAP–RNA structure (PDB entry 1gtf; resolution 1.75 Å; Hopcroft et al., 2002) as a search model. The resulting TRAP–RNA structure (TR1) was refined using REFMAC5 (Murshudov et al., 2011), initially using rigid-body refinement, followed by repeated cycles of restrained, TLS and isotropic B-factor refinement, coupled with visual inspection in Coot (Emsley et al., 2010). TR1 was refined to 1.98 Å resolution, with a dimeric assembly of non-RNA-bound and RNA-bound TRAP rings within the asymmetric unit. Consistent with previous structures of the TRAP–RNA complex, the RNA sequence termini were not observed within the 2F o − F c map; the first spacer (U4) was then modelled at all 11 repeats around the TRAP ring and the second spacer (U5) was omitted from the final refined structure. For the later data sets, the observed structure-factor amplitudes from each separately scaled data set (output from AIMLESS) were combined with the phases of TR1 and the resulting higher-dose model was refined with phenix.refine (Adams et al., 2010) using only rigid-body and isotropic B-factor refinement. During this refinement, the TRAP–RNA complex and nonbound TRAP ring were treated as two separate rigid bodies within the asymmetric unit. Supplementary Table S1 shows the relevant summary statistics.  D loss metric calculation    The CCP4 program CAD was used to create a series of nine merged .mtz files combining observed structure-factor amplitudes for the first data set F obs(d 1) with each later data set F obs(d n) (for n = 2, …, 10). All later data sets were scaled against the initial low-dose data set in SCALEIT. For each data set an atom-tagged .map file was generated using the ATMMAP mode in SFALL (Winn et al., 2011). A full set of nine Fourier difference maps F obs(d n) − F obs(d 1) were calculated using FFT (Ten Eyck, 1973) over the full TRAP–RNA unit-cell dimensions, with the same grid-sampling dimensions as the atom-tagged .map file. All maps were cropped to the TRAP asymmetric unit in MAPMASK. Comparing the atom-tagged .map file and F obs(d n) − F obs(d 1) difference map at each dose, each refined atom was assigned a set of density-change values X. The maximum density-loss metric, D loss (units of e Å−3), was calculated to quantify the per-atom electron-density decay at each dose, assigned as the absolute magnitude of the most negative Fourier difference map voxel value in a local volume around each atom as defined by the set X. Model system calculation    Model calculations were run for the simple amino acids glutamate and aspartate. In order to avoid decarboxylation at the C-terminus instead of the side chain on the Cα atom, the C-terminus of each amino acid was methylated. While the structures of the closed shell acids are well known, the same is not true of those in the oxidized state. The quantum-chemical calculations employed were chosen to provide a satisfactory description of the structure of such radical species and also provide a reliable estimation of the relative C—C(O2) bond strengths, which are otherwise not available. Structures of methyl-terminated (at the N- and C-termini) carboxylates were determined using analytic energy gradients with density functional theory (B3LYP functional; Becke, 1993) and a flexible basis set of polarized valence triple-zeta size with diffuse functions on the non-H atoms [6-311+G(d,p)] in the Gaussian 09 computational chemistry package (Frisch et al., 2009). The stationary points obtained were characterized as at least local minima by examination of the associated analytic Hessian. Effects of the medium were modelled using a dielectric cavity approach (Tomasi et al., 1999) parameterized for water. Results    Per-atom quantification of electron density    To quantify the exact effects of nucleic acid binding to a protein on SRD susceptibility, a high-throughput and automated pipeline was created to systematically calculate the electron-density change for every refined atom within the TRAP–RNA structure as a function of dose. This provides an atom-specific quantification of density–dose dynamics, which was previously lacking within the field. Previous studies have characterized SRD sites by reporting magnitudes of F obs(d n) − F obs(d 1) Fourier difference map peaks in terms of the sigma (σ) contour level (the number of standard deviations from the mean map electron-density value) at which peaks become visible. However, these σ levels depend on the standard deviation values of the map, which can deviate between data sets, and are thus unsuitable for quantitative comparison of density between different dose data sets. Instead, we use here a maximum density-loss metric (D loss), which is the per-atom equivalent of the magnitude of these negative Fourier difference map peaks in units of e Å−3. Large positive D loss values indicate radiation-induced atomic disordering reproducibly throughout the unit cells with respect to the initial low-dose data set. For each TRAP–RNA data set, the D loss metric successfully identified the recognized forms of protein SRD (Fig. 2 ▸ a), with clear Glu and Asp side-chain decarboxylation even in the first difference map calculated (3.9 MGy; Fig. 3 ▸ a). The main sequence of TRAP does not contain any Trp and Cys residues (and thus contains no disulfide bonds). The substrate Trp amino-acid ligands also exhibited disordering of the free terminal carboxyl groups at higher doses (Fig. 2 ▸ a); however, no clear Fourier difference peaks could be observed visually. Even for radiation-insensitive residues (e.g. Gly) the average D loss increases with dose: this is the effect of global radiation damage, since as dose increases the electron density associated with each refined atom becomes weaker as the atomic occupancy decreases (Fig. 2 ▸ b). Only Glu and Asp residues exhibit a rate of D loss increase that consistently exceeds the average decay (Fig. 2 ▸ b, dashed line) at each dose. Additionally, the density surrounding ordered solvent molecules was determined to significantly diminish with increasing dose (Fig. 2 ▸ b). The rate of D loss (attributed to side-chain decarboxylation) was consistently larger for Glu compared with Asp residues over the large dose range (Fig. 2 ▸ b and Supplementary Fig. S3); this observation is consistent with our calculations on model systems (see above) that suggest that, without considering differential hydrogen-bonding environments, CO2 loss is more exothermic by around 8 kJ mol−1 from oxidized Glu residues than from their Asp counterparts. RNA is less susceptible to electron-density loss than protein within the TRAP–RNA complex    Visual inspection of Fourier difference maps illustrated the clear lack of RNA electron-density degradation with increasing dose compared with the obvious protein damage manifestations (Figs. 3 ▸ b and 3 ▸ c). Only at the highest doses investigated (\u003e20 MGy) was density loss observed at the RNA phosphate and C—O bonds of the phosphodiester backbone. However, the median D loss was lower by a factor of \u003e2 for RNA P atoms than for Glu and Asp side-chain groups at 25.0 MGy (Supplementary Fig. S4), and furthermore could not be numerically distinguished from Gly Cα atoms within TRAP, which are not radiation-sensitive at the doses tested here (Supplementary Fig. S3). RNA binding protects radiation-sensitive residues    For the large number of acidic residues per TRAP ring (four Asp and six Glu residues per protein monomer), a strong dependence of decarboxylation susceptibility on local environment was observed (Fig. 4 ▸). For each Glu Cδ or Asp Cγ atom, D loss provided a direct measure of the rate of side-chain carboxyl-group disordering and subsequent decarboxylation. For acidic residues with no differing interactions between nonbound and bound TRAP (Fig. 4 ▸ a), similar damage was apparent between the two rings within the asymmetric unit, as expected. However, TRAP residues directly on the RNA-binding interfaces exhibited greater damage accumulation in nonbound TRAP (Fig. 4 ▸ b), and for residues at the ring–ring interfaces (where crystal contacts were detected) bound TRAP exhibited enhanced SRD accumulation (Fig. 4 ▸ c). Three acidic residues (Glu36, Asp39 and Glu42) are involved in RNA interactions within each of the 11 TRAP ring subunits, and Fig. 5 ▸ shows their density changes with increasing dose. Hotelling’s T-squared test (the multivariate counterpart of Student’s t-test) was used to reject the null hypothesis that the means of the D loss metric were equal for the bound and nonbound groups in Fig. 5 ▸. A significant reduction in D loss is seen for Glu36 in RNA-bound compared with nonbound TRAP, indicative of a lower rate of side-chain decarboxylation (Fig. 5 ▸ a; p = 6.06 × 10−5). For each TRAP ring subunit, the Glu36 side-chain carboxyl group accepts a pair of hydrogen bonds from the two N atoms of the G3 RNA base. In our analysis, Asp39 in the TRAP–(GAGUU)10GAG structure appears to exhibit two distinct hydrogen bonds to the G1 base within each of the 11 TRAP–RNA interfaces, as does Glu36 to G3; however, the reduction in density disordering upon RNA binding is far less significant for Asp39 than for Glu36 (Fig. 5 ▸ b, p = 0.0925). RNA binding reduces radiation-induced disorder on the atomic scale    One oxygen (O∊1) of Glu42 appears to form a hydrogen bond to a nearby water within each TRAP RNA-binding pocket, with the other (O∊2) being involved in a salt-bridge interaction with Arg58 (Hopcroft et al., 2002; Antson et al., 1999). Salt-bridge interactions have previously been suggested to reduce the glutamate decarboxylation rate within the large (∼62.4 kDa) myrosinase protein structure (Burmeister, 2000). A significant difference was observed between the D loss dynamics for the nonbound/bound Glu42 O∊1 atoms (Fig. 5 ▸ c; p = 0.007) but not for the Glu42 O∊2 atoms (Fig. 5 ▸ d; p = 0.239), indicating that the stabilizing strength of this salt-bridge interaction was conserved upon RNA binding and that the water-mediated hydrogen bond had a greater relative susceptibility to atomic disordering in the absence of RNA. The density-change dynamics were statistically indistinguishable between bound and nonbound TRAP for each Glu42 carboxyl group Cδ atom (p = 0.435), indicating that upon RNA binding the conserved salt-bridge interaction ultimately dictated the overall Glu42 decarboxylation rate. The RNA-stabilizing effect was not restricted to radiation-sensitive acidic residues. The side chain of Phe32 stacks against the G3 base within the 11 TRAP RNA-binding interfaces (Antson et al., 1999). With increasing dose, the D loss associated with the Phe32 side chain was significantly reduced upon RNA binding (Fig. 5 ▸ e; Phe32 Cζ; p = 0.0014), an indication that radiation-induced conformation disordering of Phe32 had been reduced. The extended aliphatic Lys37 side chain stacks against the nearby G1 base, making a series of nonpolar contacts within each RNA-binding interface. The D loss for Lys37 side-chain atoms was also reduced when stacked against the G1 base (Fig. 5 ▸ f; p = 0.0243 for Lys37 C∊ atoms). Representative Phe32 and Lys37 atoms were selected to illustrate these trends. Discussion    Here, MX radiation-induced specific structural changes within the large TRAP–RNA assembly over a large dose range (1.3–25.0 MGy) have been analysed using a high-throughput quantitative approach, providing a measure of the electron-density distribution for each refined atom with increasing dose, D loss. Compared with previous studies, the results provide a further step in the detailed characterization of SRD effects in MX. Our method­ology, which eliminated tedious and error-prone visual inspection, permitted the determination on a per-atom basis of the most damaged sites, as characterized by F obs(d n) − F obs(d 1) Fourier difference map peaks between successive data sets collected from the same crystal. Here, it provided the precision required to quantify the role of RNA in the damage susceptibilities of equivalent atoms between RNA-bound and nonbound TRAP, but it is applicable to any MX SRD study. The RNA was found to be substantially more radiation-resistant than the protein, even at the highest doses investigated (∼25.0 MGy), which is in strong concurrence with our previous SRD investigation of the C.Esp1396I protein–DNA complex (Bury et al., 2015). Consistent with that study, at high doses of above ∼20 MGy, F obs(d n) − F obs(d 1) map density was detected around P, O3′ and O5′ atoms of the RNA backbone, with no significant difference density localized to RNA ribose and basic subunits. RNA backbone disordering thus appears to be the main radiation-induced effect in RNA, with the protein–base interactions maintained even at high doses (\u003e20 MGy). The U4 phosphate exhibited marginally larger D loss values above 20 MGy than G1, A2 and G3 (Supplementary Fig. S4). Since U4 is the only refined nucleotide not to exhibit significant base–protein interactions around TRAP (with a water-mediated hydrogen bond detected in only three of the 11 subunits and a single Arg58 hydrogen bond suggested in a further four subunits), this increased U4 D loss can be explained owing to its greater flexibility. At 25.0 MGy, the magnitude of the RNA backbone D loss was of the same order as for the radiation-insensitive Gly Cα atoms and on average less than half that of the acidic residues of the protein (Supplementary Fig. S3). Consequently, no clear single-strand breaks could be located, and since RNA-binding within the current TRAP–(GAGUU)10GAG complex is mediated predominantly through base–protein interactions, the biological integrity of the RNA complex was dictated by the rate at which protein decarboxylation occurred. RNA interacting with TRAP was shown to offer significant protection against radiation-induced structural changes. Both Glu36 and Asp39 bind directly to RNA, each through two hydrogen bonds to guanine bases (G3 and G1, respectively). However, compared with Asp39, Glu36 is strikingly less decarboxylated when bound to RNA (Fig. 4 ▸). This is in good agreement with previous mutagenesis and nucleoside analogue studies (Elliott et al., 2001), which indicated that the G1 nucleotide does not bind to TRAP as strongly as do A2 and G3, and plays little role in the high RNA-binding affinity of TRAP (K d ≃ 1.1 ± 0.4 nM). For Glu36 and Asp39, no direct quantitative correlation could be established between hydrogen-bond length and D loss (linear R 2 of \u003c0.23 for all doses; Supplementary Fig. S5). Thus, another factor must be responsible for this clear reduction in Glu36 CO2 decarboxyl­ation in RNA-bound TRAP. The Glu36 carboxyl side chain also potentially forms hydrogen bonds to His34 and Lys56, but since these interactions are conserved irrespective of G3 nucleotide binding, this cannot directly account for the stabilization effect on Glu36 in RNA-bound TRAP. Radiation-induced decarboxylation has been proposed to be mediated by preferential positive-hole migration to the side-chain carboxyl group, with rapid proton transfer trapping the hole at the carboxyl group (Burmeister, 2000; Symons, 1997):where the forward rate is K 1 and the backward rate is K −1, where the forward rate is K 2. When bound to RNA, the average solvent-accessible area of the Glu36 side-chain O atoms is reduced from ∼15 to 0 Å2. We propose that with no solvent accessibility Glu36 decarboxylation is inhibited, since the CO2-formation rate K 2 is greatly reduced, and suggest that steric hindrance prevents each radicalized Glu36 CO2 group from achieving the planar conformation required for complete dissociation from TRAP. The electron-recombination rate K −1 remains high, however, owing to rapid electron migration through the protein–RNA complex to refill the Glu36 positive hole (the precursor for Glu decarboxylation). Upon RNA binding, the Asp39 side-chain carboxyl group solvent-accessible area changes from ∼75 to 35 Å2, still allowing a high CO2-formation rate K 2. Previous studies have reported inconsistent results concerning the dependence of the acidic residue decarboxylation rate on solvent accessibility (Weik et al., 2000; Fioravanti et al., 2007; Gerstel et al., 2015). The prevalence of radical attack from solvent channels surrounding the protein in the crystal is a questionable cause, considering previous observations indicating that the strongly oxidizing hydroxyl radical is immobile at 100 K (Allan et al., 2013; Owen et al., 2012). Furthermore, the suggested electron hole-trapping mechanism which induces decarboxylation within proteins at 100 K has no clear mechanistic dependence on the solvent-accessible area of each carboxyl group. By comparing equivalent acidic residues with and without RNA, we have now deconvoluted the role of solvent accessibility from other local protein environment factors, and thus propose a suitable mechanism by which exceptionally low solvent accessibility can reduce the rate of decarboxylation. Overall, no direct correlation between solvent accessibility and decarboxylation susceptibility was observed, but it is very clear that inaccessible residues are protected. Apart from these RNA-binding interfaces, RNA binding was seen to enhance decarboxylation for residues Glu50, Glu71 and Glu73, all of which are involved in crystal contacts between TRAP rings (Fig. 4 ▸ c). However, for each of these residues the exact crystal contacts are not preserved between bound and nonbound TRAP or even between monomers within one TRAP ring. For example, in bound TRAP, Glu73 hydrogen-bonds to a nearby lysine on each of the 11 subunits, whereas in nonbound TRAP no such interaction exists and Glu73 interacts with a variable number of refined waters in each subunit. Thus, the dependence of decarboxylation rates on these interactions could not be established. Radiation-induced side-chain conformational changes have been poorly characterized in previous SRD investigations owing to their strong dependence on packing density and geometric strain. Such structural changes are known to have significant roles within enzymatic pathways, and experimenters must be aware of these possible confounding factors when assigning true functional mechanisms using MX. Our results show that RNA binding to TRAP physically stabilizes non-acidic residues within the TRAP macromolecule, most notably Lys37 and Phe32, which stack against the G1 and G3 bases, respectively. It has been suggested (Burmeister, 2000) that Tyr residues can lose their aromatic –OH group owing to radiation-induced effects; however, no energetically favourable pathway for –OH cleavage exists and this has not been detected in aqueous radiation-chemistry studies. In TRAP, D loss increased at a similar rate for both the Tyr O atoms and aromatic ring atoms, suggesting that full ring conformational disordering is more likely. Indeed, no convincing reproducible Fourier difference peaks above the background map noise were observed around any Tyr terminal –OH groups. The RNA-stabilization effects on protein are observed at short ranges and are restricted to within the RNA-binding interfaces around the TRAP ring. For example, Asp17 is located ∼6.8 Å from the G1 base, outside the RNA-binding interfaces, and has indistinguishable Cγ atom D loss dose-dynamics between RNA-bound and nonbound TRAP (p \u003e 0.9). An increase in the dose at which functionally important residues remain intact has biological ramifications for understanding the mechanisms at which ionizing radiation damage is mitigated within naturally forming DNA–protein and RNA–protein complexes. Observations of lower protein radiation-sensitivity in DNA-bound forms have been recorded in solution at RT at much lower doses (∼1 kGy) than those used for typical MX experiments [e.g. an oestrogen response element–receptor complex (Stísová et al., 2006) and a DNA glycosylase and its abasic DNA target site (Gillard et al., 2004)]. In these studies, the main damaging species is predicted to be the oxidizing hydroxyl radical produced through solvent irradiation, which is known to add to double covalent bonds within both DNA and RNA bases to induce strand breaks and base modification (Spotheim-Maurizot \u0026 Davídková, 2011; Chance et al., 1997). It was suggested that physical screening of DNA by protein shielded the DNA–protein interaction sites from radical damage, yielding an extended life-dose for the nucleoprotein complex compared with separate protein and DNA constituents at RT. However, in the current MX study at 100 K, the main damaging species are believed to be migrating LEEs and holes produced directly within the protein–RNA components or in closely associated solvent. The results presented here suggest that biologically relevant nucleoprotein complexes also exhibit prolonged life-doses under the effect of LEE-induced structural changes, involving direct physical protection of key RNA-binding residues. Such reduced radiation-sensitivity in this case ensures that the interacting protein remains bound long enough to the RNA to complete its function, even whilst exposed to ionizing radiation. Within the nonbound TRAP macromolecule, the acidic residues within the unoccupied RNA-binding interfaces (Asp39, Glu36, Glu42) are notably amongst the most susceptible residues within the asymmetric unit (Fig. 4 ▸). When exposed to X-rays, these residues will be preferentially damaged by X-rays and subsequently reduce the affinity with which TRAP binds to RNA. Within the cellular environment, this mechanism could reduce the risk that radiation-damaged proteins might bind to RNA, thus avoiding the detrimental introduction of incorrect DNA-repair, transcriptional and base-modification pathways. The Python scripts written to calculate the per atom D loss metric are available from the authors on request. 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The TRAP–(GAGUU)10GAG complex asymmetric unit (PDB entry 1gtf; Hopcroft et al., 2002). Bound tryptophan ligands are represented as coloured spheres. RNA is shown is yellow. (a) Electron-density loss sites as indicated by D\nloss in the TRAP–RNA complex crystal by residue/nucleotide type for five doses [sites determined above the 4× average D\nloss threshold, calculated over the TRAP–RNA structure for the first difference map: F\nobs(d\n2) − F\nobs(d\n1)]. Cumulative frequencies are normalized to both the total number of non-H atoms per residue/nucleotide and the total number of that residue/nucleotide type present. (b) Average D\nloss for each residue/nucleotide type with respect to the DWD (diffraction-weighted dose; Zeldin, Brock­hauser et al., 2013). 95% confidence intervals (CI) are shown. Only a subset of key TRAP residue types are included. The average D\nloss (calculated over the whole TRAP asymmetric unit) is shown at each dose (dashed line). \nF\nobs(d\nn) − F\nobs(d\n1) Fourier difference maps for (a) n = 2 (3.9 MGy), (b) n = 3 (6.5 MGy) and (c) n = 7 (16.7 MGy) contoured at ±4σ (a) and ±3.5σ (b, c). In (a) clear difference density is observed around the Glu42 carboxyl side chain in chain H, within the lowest dose difference map at d\n2 = 3.9 MGy. Radiation-induced protein disordering is evident across the large dose range (b, c); in comparison, no clear deterioration of the RNA density was observed. \nD\nloss calculated for all side-chain carboxyl group Glu Cδ and Asp Cγ atoms within the TRAP–RNA complex for a dose of 19.3 MGy (d\n8). Residues have been grouped by amino-acid number, and split into bound and nonbound groupings, with each bar representing the mean calculated over 11 equivalent atoms around a TRAP ring. Whiskers indicate 95% CI. The D\nloss behaviour shown here was consistently exhibited across the entire investigated dose range. \nD\nloss against dose for (a) Glu36 Cδ, (b) Asp39 Cγ, (c) Glu42 O∊1, (d) Glu42 O∊2, (e) Phe32 Cζ and (f) Lys37 C∊ atoms. 95% CI are included for each set of 11 equivalent atoms grouped as bound/nonbound. RNA-binding interface interactions are shown for TRAP chain N, with the F\nobs(d\n7) − F\nobs(d\n1) Fourier difference map (dose 16.7 MGy) overlaid and contoured at a ±4σ 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diff --git a/annotated_BioC_JSON/PMC4855620_ann.json b/annotated_BioC_JSON/PMC4855620_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..007da7b8bfde79cb162673f4321825d6f24a6783
--- /dev/null
+++ b/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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+[{"sourceid":"4857006","sourcedb":"","project":"","target":"","text":"Reversal of DNA damage induced Topoisomerase 2 DNA–protein crosslinks by Tdp2 Mammalian Tyrosyl-DNA phosphodiesterase 2 (Tdp2) reverses Topoisomerase 2 (Top2) DNA–protein crosslinks triggered by Top2 engagement of DNA damage or poisoning by anticancer drugs. Tdp2 deficiencies are linked to neurological disease and cellular sensitivity to Top2 poisons. Herein, we report X-ray crystal structures of ligand-free Tdp2 and Tdp2-DNA complexes with alkylated and abasic DNA that unveil a dynamic Tdp2 active site lid and deep substrate binding trench well-suited for engaging the diverse DNA damage triggers of abortive Top2 reactions. Modeling of a proposed Tdp2 reaction coordinate, combined with mutagenesis and biochemical studies support a single Mg2+-ion mechanism assisted by a phosphotyrosyl-arginine cation-π interface. We further identify a Tdp2 active site SNP that ablates Tdp2 Mg2+ binding and catalytic activity, impairs Tdp2 mediated NHEJ of tyrosine blocked termini, and renders cells sensitive to the anticancer agent etoposide. Collectively, our results provide a structural mechanism for Tdp2 engagement of heterogeneous DNA damage that causes Top2 poisoning, and indicate that evaluation of Tdp2 status may be an important personalized medicine biomarker informing on individual sensitivities to chemotherapeutic Top2 poisons. INTRODUCTION Nuclear DNA compaction and the action of DNA and RNA polymerases create positive and negative DNA supercoiling—over- and under-winding of DNA strands, respectively—and the linking together (catenation) of DNA strands. Topoisomerases relieve topological DNA strain and entanglement to facilitate critical nuclear DNA transactions including DNA replication, transcription and cell division. The mammalian type II topoisomerases Top2α and Top2β enzymes generate transient, reversible DNA double strand breaks (DSBs) to drive topological transactions. Reversibility of Top2 DNA cleavage reactions is facilitated by formation of covalent enzyme phosphotyrosyl linkages between the 5′-phosphate ends of the incised duplex and an active site Top2 tyrosine, resulting in Top2 cleavage complexes (Top2cc). The Top2cc protein–DNA adduct is a unique threat to genomic integrity which must be resolved to prevent catastrophic Top2cc collisions with the cellular replication and transcription machineries. To promote cancer cell death, Top2 reactions are ‘poisoned’ by keystone pharmacological anticancer agents like etoposide, teniposide and doxorubicin. Importantly, Top2 is also poisoned when it engages abundant endogenous DNA damage not limited to but including ribonucleotides, abasic sites and alkylation damage such as exocyclic DNA adducts arising from bioactivation of the vinyl chloride carcinogen (Figure 1A). In the case of DNA damage-triggered Top2cc, compound DNA lesions arise that consist of the instigating lesion, and a DNA DSB bearing a bulky terminal 5′-linked Top2 DNA–protein crosslink. The chemical complexity of DNA damage-derived Top2cc necessitates that DNA repair machinery dedicated to resolving these lesions recognizes both DNA and protein, whilst accommodating diverse chemical structures that trap Top2cc. Precisely how the cellular DNA repair machinery navigates these complex lesions is an important aspect of Top2cc repair that has not yet been explored. Tdp2 processes phosphotyrosyl linkages in diverse DNA damage contexts. (A) Unrepaired DNA damage and repair intermediates such as bulky DNA adducts, ribonucleotides or abasic sites can poison Top2 and trap Top2 cleavage complex (Top2cc), resulting in a DSB with a 5′–Top2 protein adduct linked by a phosphotyrosine bond. Tdp2 hydrolyzes the 5′–phosphotyrosine adduct derived from poisoned Top2 leaving DNA ends with a 5′-phosphate, which facilitates DNA end joining through the NHEJ pathway. (B) DNA oligonucleotide substrates synthesized by EDC-imidazole coupling and used in Tdp2 enzyme assays contain deoxyadenine (dA), Ethenoadenine (ϵA) or an abasic site (THF) and a 5′–nitrophenol moiety. Phosphotyrosyl bond hydrolysis catalyzed by mTdp2cat releases p-nitrophenol, which is detected by measuring absorbance at 415 nm. (C) mTdp2cat reaction rates on p–nitrophenol modified DNA substrates shown in panel B. Rates are reported as molecules of PNP s−1 produced by mTdp2cat. P-values calculated using two-tailed t-test; error bars, s.d. n = 4, n.s. = not statistically significant. (D) Structure of mTdp2cat bound to 5′-phosphate DNA (product complex) containing ϵA (yellow). DNA binding β2Hβ–grasp (tan) and cap elements engage the 5′-nucleotide as well as the +2 and +3 nucleotides (blue) of substrate DNA. PDB entry 5HT2 is displayed, also see Table 1. (E) Structure of mTdp2cat bound to 5′-phosphate DNA (product complex) containing THF (yellow). DNA binding β2Hβ–grasp (tan) and cap elements engage the 5′-nucleotide as well as the +2 and +3 nucleotides (blue) of substrate DNA. PDB entry 5INK is displayed, also see Table 1. (F) Structure of mTdp2cat in the absence of DNA showing the extended 3-helix loop (tan) open-conformation of the DNA-binding grasp as seen in monomer E of the apo structure. PDB entry 5INM is displayed, also see Table 1. Tyrosyl DNA phosphodiesterase 2 (Tdp2) directly hydrolyzes 5′-phosphotyrosyl (5′-Y) linkages, and is a key modulator of cellular resistance to chemotherapeutic Top2 poisons. Tdp2 knockdown sensitizes A549 lung cancer cells to etoposide, and increases formation of nuclear γH2AX foci, a marker of DSBs, underlining the importance of Tdp2 in cellular Top2cc repair. Tdp2 is overexpressed in lung cancers, is transcriptionally up-regulated in mutant p53 cells and mediates mutant p53 gain of function phenotypes, which can lead to acquisition of therapy resistance during cancer progression. The importance of Tdp2 in mediating topoisomerase biology is further underlined by the facts that human TDP2 inactivating mutations are found in individuals with intellectual disabilities, seizures and ataxia, and at the cellular level, loss of Tdp2 inhibits Top2β-dependent transcription. It is possible that TDP2 single nucleotide polymorphisms (SNPs) encode mutations that impact Tdp2 function, but the molecular underpinnings for such Tdp2 deficiencies are not understood. Previously we reported high-resolution X-ray crystal structures of the minimal catalytically active endonuclease/exonuclease/phosphatase (EEP) domain of mouse Tdp2 (mTdp2cat) bound to a DNA substrate mimic, and a 5′-phosphorylated reaction product. However, important questions regarding the mechanism of Tdp2 engagement and processing of DNA damage remain. First, it is unclear if Tdp2 processes phosphotyrosyl linkages in the context of DNA damage that triggers Top2cc, and if so, how the enzyme can accommodate such complex DNA damage within its active site. Based on metal-bound Tdp2 structures, we also proposed a single Mg2+ mediated catalytic mechanism, but this mechanism requires further scrutiny and characterization. Herein, we report an integrated structure-function study of the Tdp2 reaction mechanism, including a description of new X-ray structures of ligand-free Tdp2, and Tdp2 bound to abasic and alkylated (1-N6-etheno-adenine) DNA damage. Our integrated results from structural analysis, mutagenesis, functional assays and quanyum mechanics/molecular mechanics (QM/MM) modeling of the Tdp2 reaction coordinate describe in detail how Tdp2 mediates a single-metal ion tyrosyl DNA phosphodiesterase reaction capable of acting on diverse DNA end damage. We further establish that DNA damage binding in the Tdp2 active site is linked to conformational change and binding of metal cofactor. Finally, we characterize a Tdp2 SNP that ablates the Tdp2 single metal binding site and Tdp2 substrate induced conformational changes, and confers Top2 drug sensitivity in mammalian cells. MATERIALS AND METHODS Generation of PNP modified oligonucleotide substrates Oligonucleotides with a 5′-phosphate modification were obtained from IDT and diluted to a concentration of 2 mmol l−1 in water. 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC; Pierce) was used to couple p-nitrophenol to the oligonucleotide with a modified version of the manufacturer's instructions. Briefly, 25 mg EDC were dissolved in 150 μl oligonucleotide solution. After the EDC dissolved, 400 μl DDW, 60 μl 100 mmol l−1 Imidazole pH 6 and 100 μl of PNP from a saturated water/PNP mixture preheated to 55°C were added. Reactions were heated at 55°C for 30 min to ensure the PNP dissolved, then incubated at 37°C overnight. Reactions were quenched with 500 μl 2 mol l−1 acetic acid and heated to 55°C for 1 h, followed by neutralization with 500 μl 2 mol l−1 Tris base. Reactions were twice diluted to 15 ml in DDW then concentrated to 300 μl with a 3K MWT cutoff spin concentrator (Amicon), then run on 20% (w/v) 19:1 8M urea-TBE PAGE to resolve reaction products. Bands were visualized by UV shadow, excised, soaked in 15 ml DDW for 16 h at 15°C, and purified on a C18 reverse-phase Sep-Pac (Waters). Protein expression, purification and crystallization Mouse Tdp2 catalytic domain (mTdp2cat, residues 118 to 370), human Tdp2 catalytic domain (hTdp2cat, residues 108–362) or full-length human Tdp2 (hTdp2FL, residues 1–362) were expressed and purified as previously described. Plasmids containing mutant Tdp2 sequences were generated using the Quickchange kit (Stratagene). Crystals containing mTdp2cat and 5′-phosphorylated DNA (product) with modified 5′ nucleotides (ϵA, dA, THF) were grown and cryoprotected as described. Sequences, modifications and synthesis sources for oligonucleotides used for co-crystallization are indicated in Supplementary Table S1. For alternate divalent metal complexes, a 5′-phosphorylated DNA substrate (substrate dC) was co-crystallized in the presence of Mg2+, and divalent metals were swapped by soaking the crystals in crystallization buffer containing 5 mmol l−1 MnCl2 or 10 mmol l−1 Ca(OAc)2 for 1 h prior to cryoprotection. Cryoprotectant solutions contained mother liquor plus 25% PEG-3350, 8% glycerol and 5% glucose, and either 5 mmol l−1 MnCl2 or 10 mmol l−1 Ca(OAc)2. Crystals of hTdp2cat bound to 5′-phosphate DNA (substrate dC) were prepared by mixing a 1.2-fold molar excess of DNA, and grown in 90 mmol l−1 TRIS pH 7.0, 27% (w/v) PEG600, 9% (v/v) glycerol and 450 mmol l−1 (NH4)2SO4. Crystals of apo-mTdp2cat were grown in 14–18% (w/v) PEG3350, 100 mmol l−1 HEPES pH 7.5, 200 mmol l−1 Li2SO4 and 10 mmol l−1 Mg(OAc)2, and soaked into the same buffer containing 25% PEG3350 and 12% (v/v) glycerol prior to flash-freezing in liquid nitrogen for data collection. X-ray diffraction data collection, phasing and refinement X-ray data (Table 1) for all structures except the manganese soak (PDB entry 5INP) were collected at 100 K on beamline 22-ID of the Advanced Photon Source at a wavelength of 1.000 Å. X-ray data for the manganese soak (PDB entry 5INP) were collected on a Rigaku HF007 Cu rotating anode X-ray source at a wavelength of 1.5418 Å. X-ray diffraction data were processed and scaled using the HKL2000 suite. The hTdp2cat-DNA and mTdp2cat-apo crystals were phased by molecular replacement in PHASER using chain A of PDB entry 4GZ1. Initial solutions were improved by iterative rounds of manual fitting in COOT and refinement in PHENIX. Data collection and refinement statistics \tmTdp2 118–370 5′-P +ϵA\tmTdp2 118–370 5′-P + THF\tmTdp2 118–370 5′-P + dA\tmTdp2 118–370 wt, apo\tmTdp2 118–370 D358N, apo\thTdp2 108–362 5′-P DNA\tmTdp2 118–370 5′-P DNA Mn2+ soak\tmTdp2 118–370 5′-P DNA Ca2+ soak\t \tPDB entry ID\t5HT2\t5INK\t5INL\t5INM\t5INN\t5INO\t5INP\t5INQ\t \tData collection\t \tSpace group\tP 21 21 21\tP 21 21 21\tP 21 21 21\tP 21 21 21\tP 21 21 21\tP 31\tP 21 21 21\tP 21 21 21\t \tCell dimensions\t \ta, b, c (Å)\t54.88, 68.60, 167.25\t54.62, 67.66, 166.74\t54.90, 69.05, 167.80\t95.85, 113.44, 114.96\t95.44, 114.88, 115.45\t69.66, 69.66, 120.68\t54.92, 68.52, 166.90\t54.70, 67.50, 167.52\t \tα, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 120\t90, 90, 90\t90, 90, 90\t \tResolution (Å)\t50–1.43 (1.48–1.43)\t50–2.15 (2.23–2.15)\t50–1.55 (1.61–1.55)\t50–2.40 (2.49–2.40)\t50–2.80 (2.90–2.80)\t50–3.20 (3.31–3.20)\t50–1.95 (2.02–1.95)\t50–1.85 (1.92–1.85)\t \tRsym or Rmerge\t0.051 (0.599)\t0.125 (0.650)\t0.067 (0.588)\t0.124 (0.518)\t0.113 (0.459)\t0.082 (0.574)\t0.054 (0.181)\t0.068 (0.564)\t \tI / σI\t21.0 (2.1)\t17.5 (2.7)\t19.4 (2.4)\t9.8 (2.1)\t10.6 (2.0)\t15.5 (2.2)\t25.5 (4.8)\t21.1 (3.1)\t \tCompleteness (%)\t97.4 (86.6)\t98.4 (95.4)\t99.0 (91.5)\t98.0 (94.2)\t95.7 (93.0)\t100 (100)\t97.1 (80.7)\t99.9 (100)\t \tRedundancy\t3.9 (3.0)\t6.7 (6.5)\t5.4 (3.7)\t4.2 (3.6)\t4.2 (3.4)\t3.9 (3.9)\t5.5 (2.0)\t6.0 (6.0)\t \tRefinement\t \tResolution (Å)\t29.9–1.43\t45.7–2.15\t34.1–1.55\t48.8–2.4\t49.3–2.80\t34.8–3.205\t29.8–1.95\t39.1–1.85\t \tNo. reflections\t114 115\t34 369\t91 871\t49 905\t29 807\t10 757\t45 238\t53 889\t \tRwork / Rfree\t0.116/0.150\t0.167/0.201\t0.129/0.168\t0.183/0.233\t0.214/0.255\t0.214/0.264\t0.152/0.179\t0.164/0.200\t \tNon-H atoms\t \tProtein/DNA\t4552\t4372\t4600\t9621\t9411\t4363\t4512\t4452\t \tLigand/ion\t25\t22\t25\t25\t28\t5\t15\t21\t \tWater\t726\t449\t721\t365\t148\t5\t444\t374\t \tB-factors (Å2)\t \tProtein/DNA\t26.5\t35.8\t26.7\t47.7\t53.3\t147.9\t35.6\t50.4\t \tLigand/ion\t52.5\t50.2\t53.0\t73.5\t82.2\t225.8\t53.7\t54.7\t \tWater\t44.2\t43.0\t46.0\t38.2\t28.3\t83.3\t44.7\t52.4\t \tR.m.s. deviations\t \tBond lengths (Å)\t0.007\t0.007\t0.005\t0.004\t0.002\t0.003\t0.007\t0.008\t \tBond angles (°)\t1.18\t1.01\t0.85\t0.81\t0.58\t0.78\t1.03\t1.08\t \t Each data set was collected from a single crystal. Values in parentheses are for highest-resolution shell (10% of relections). Limited proteolysis assays For proteolysis experiments, 4 μl reactions containing 40 μmol l−1 mTdp2cat (aa 118 to 370) in reaction buffer (10 mmol l−1 HEPES pH 7.5, 200 mmol l−1 NaCl, 0.5 mmol L−1 TCEP) with 4 mmol l−1 Mg(OAc)2 (indicated +Mg) or 8 mmol l−1 NaOAc (indicated -Mg), with 0 or 60 μmol l−1 12 nt DNA were incubated with 0, 5, 1.7 or 0.6 μg l−1 Trypsin for 1 h at 22°C. Reactions were quenched by addition of Laemmli SDS-PAGE dye, heated at 70°C for 10 min, and analyzed by SDS-PAGE. For mass-spectrometry analysis of peptide masses, reactions were quenched with 1% (v/v) trifluoroacetic acid, purified on a C18 ZipTip (Millipore) and an ESI-MS mass measurement made on a Q-ToF Ultima/Global (Micromass/Waters) using flow injection from a pressurized bomb. The instrument was operated in the positive ion, V-mode and calibrated using the multiply-charged ion envelope of horse heart cytochrome C. The molecular ion mass was determined using the Max Ent 1 routine from the MassLynx software. Intrinsic tryptophan fluorescence Reactions contained 50 μl with 1 μmol l−1 mTdp2cat (residues 118 to 370) in buffer (6 mmol l−1 HEPES pH 7.5, 300 mmol l−1 NaCl, 0.3 mmol L−1 TCEP and 0.01% (v/v) TWEEN-20) with 0–20 mmol l−1 Mg(OAc)2 or ultrapure Ca(OAc)2 (99.9965%, Alfa Aesar) titrated against 0–40 mmol l−1 NaOAc to maintain a constant concentration of acetate, with 0 or 1.5 μmol l−1 12 nt DNA. Reactions were incubated at room temperature for 20 minutes in a 96-well black plastic plate (Corning), then tryptophan fluorescence was measured in a Polarstar Omega platereader (BMG Labtech) with 4 readings of 150 pulses per well using 280/10 excitation and 350/10 emission filters. The increase in fluorescent intensity was calculated by subtracting the fluorescent intensity from samples with no divalent metal ions and plotted as a function of divalent metal ion concentration. Kd values and Hill coefficients were calculated using PRISM6. Tdp2 enzyme assays Assays on MBP-fusion proteins of the human catalytic domain (MBP-hTdp2cat) mutant proteins with 5′ -tyrosylated DNA substrates (Figures 5E and 6C and Supplementary Figure S5C) were performed as described. For experiments measuring the effect of divalent metal ions on reaction rates, 50 μl reactions contained 1 μmol l−1 mTdp2cat (residues 118 to 370, Figure 4B), hTdp2cat (residues 108–362) or hTdp2FL (residues 108–362) in buffer (10 mmol l−1 HEPES pH 7.5, 100 mmol l−1 NaCl, 0.5 mmol l−1 TCEP, 0.4 mg ml−1 BSA, 0.02% (v/v) TWEEN-20, 1 mmol l−1 Thymidine 5′–p-nitrophenyl phosphate) with 0–100 mmol l−1 Mg(OAc)2 titrated against 20–220 mmol l−1 NaOAc to maintain a constant concentration of acetate. Reactions contained 0, 1 or 10 mmol l−1 ultrapure Ca(OAc)2 (99.9965%, Alfa Aesar), with 0, 2 or 20 mmol l−1 less NaOAc to maintain a constant acetate concentration. PNP formation was monitored by the absorbance at 415 nm with a background correction at 515 nm. The change in absorbance at 415 nm at 10 min as a function of Mg2+ concentration was plotted. Preparation of NHEJ substrates Oligonucleotide 300 bp substrates with 5′ phosphorylated GATC overhangs were generated as previously described. Comparable substrate with 5′-phophotyrosine adducted GATC overhangs were generated by annealing the modified strand to complementary strands that generate caps for substrate head and tail ends. Head and tail caps have a 5′-phosphotyrosine-GATC overhang terminus on one end; the 5′-phosphorylated, non-adducted overhangs on the other end are made complementary to the head and tail ends of a 270 bp double stranded core fragment generated by polymerase chain reaction. Ligation of an excess of these caps to the 270 bp fragment generates a 300 bp substrate with 5′-phosphotyrosine end structures as described in Figure 7. Unligated caps are removed using a Qiaquick PCR cleanup kit. In vitro NHEJ reactions Purified NHEJ proteins (Ku, XRCC4-ligase IV, XLF) were prepared as previously described. End joining reactions were performed using 5 nM DNA substrate, 25 nM Ku, 25 nM XRCC4/LigaseIV complex, 50 nM XLF and hTdp2FL proteins as indicated. Reactions contained 25 mM Tris-HCl pH 7.5, 0.1 mM ethylenediaminetetraacetic acid (EDTA), 2 mM dithiothreitol (DTT), 125 mM KCl, 5 mM MgCl2, 100 μM ATP, 8% (w/v) polyethyleneglycol, 0.05% (w/v) Triton X-100, 50 μg ml−1 bovine serum albumin (BSA) and 50 ng supercoiled plasmid DNA. Reactions were carried out in a final volume of 10 μl and incubated at 37°C for 5 min. Reactions were stopped by the addition of 0.1% (w/v) SDS and 5 mM EDTA and analyzed by 5% native PAGE. Cellular NHEJ assays Mouse embryo fibroblast (MEF) cells from matched Tdp2+/+ and Tdp2−/− mice were a gift from F. Cortes-Ledesma, and were immortalized by transformation with a construct that expresses SV40 T-antigen (Addgene #1779). HCT-116 cells and a ligase IV deficient variant were the gift of E. A. Hendrickson. The cDNAs with wild-type human Tdp2 and polymorphic variants generated by mutagenesis were introduced into pLX302 (Addgene#25896) to prepare lentivirus. Tdp2−/− cells were infected with lentivirus and bulk cell cultures expressing lentiviral delivered construct purified by treatment with puromycin. Expression of human Tdp2 (hTdp2FL) was validated by Western analysis (12203-1-AP; Proteintech). Fifty nanograms of the 300 bp substrates used in in vitro experiments and 2 μg of carrier supercoiled plasmid DNA were introduced into 2 × 105 MEF or HCT 116 cells by electroporation (Neon, Invitrogen) using a 10 μl chamber and a single 1350 V, 30 ms pulse. Cells were recovered 1 h later, washed with phosphate buffered saline and DNA purified using a Qiamp DNA mini kit on a Qiacube. NHEJ products were quantified by qPCR and characterized by amplicon sequencing as previously described. Etoposide Clonogenic survival assays Clonogenic survival assays were carried out by treating log phase cells with Etoposide as described in the legend to Figure 7 before seeding treated and mock treated cells in 10 cm dishes. Colonies formed after 10 days post-treatment were strained using a crystal violet (0.5% w/v) solution. Plates containing a minimum of 50 colonies were counted by hand, and at least three plates were counted for each dose. QM/MM In the QM/MM calculation, in addition to the water nucleophile and the putative catalytic Lewis base (Asp272), the side chains of proximal residues (Asn130, Asn274, His236, His326, His359, Ser239 and Glu162), the bound phosphate moiety, the Mg2+ and three waters in its coordinate shell are all included in the QM sub-system. In addition, we modeled the position of the Top2 peptide tyrosine based on conformations of the substrate analog. The QM sub-system of this study consisted of 110 atoms with a zero net charge on the sub-system. QM/MM calculations are performed using Gaussian09.D01. Since the QM sub-system contains a large number of buried atoms and commonly used electrostatic potential fitting schemes to obtain the charge distribution at atomic positions become unreliable for such systems with buried atoms, we have selected the CM5 charge model in the current study to calculate the charges on atomic positions at each step of the QM/MM calculation. This CM5 scheme is an extension to the Hirshfeld population analysis and is adapted to handle buried charges properly. This charge distribution of the QM sub-system was used to evaluate contributions of residues in the MM region to the net stability of the transition and product states as compared to that of the initial reactive state. As reported previously, this residue analysis is solely based on the electrostatic energy contributions to the initial, transition and product states. According to this estimation, the residues Arg142, Lys213, Ser235, Asp277, Glu279, Asp292, Glu295, Asp308, Asp343, Arg354 and Trp360 contribute toward the stability of both the transition and product states as compared with the reactant. However, the electrostatic contributions from residues Asp132, Asp135, Glu186, Glu242, Arg247, Thr273, Arg276, Lys299, Lys322, Arg324, Arg327 and Asp358 have the opposite effect toward the stabilities of the transition and product states. RESULTS Tdp2 processing of compound DNA damage Two potent Top2 poisons include bulky alkylated DNA helix-distorting DNA base adducts (e.g. 1-N6-ethenoadenine, ϵA) and abundant abasic sites (Figure 1A). Whether Tdp2 processes phosphotyrosyl linkages within these diverse structural contexts is not known. To test this, we adapted an EDC coupling method to generate 5′-terminal p-nitrophenol (PNP) modified oligonucleotides that also harbored DNA damage at the 5′-nucleotide position (see Materials and Methods). We then evaluated the ability of a recombinant purified mouse Tdp2 catalytic domain (mTdp2cat) to release PNP (a structural mimic of a topoisomerase tyrosine) from the 5′-terminus of compound damaged DNA substrates using a colorimetric assay (Figure 1B). We observe robust Tdp2-dependent release of PNP from 5′-modified oligonucleotides in the context of dA-PNP, ϵA-PNP or the abasic-site analog tetrahydrofuran spacer (THF) (Figure 1C). Thus, Tdp2 efficiently cleaves phosphotyrosyl linkages in the context of a compound 5′ lesions composed of abasic or bulky DNA base adduct DNA damage. To understand the molecular basis for Tdp2 processing of Top2cc in the context of DNA damage, we crystallized and determined X-ray crystal structures of mTdp2cat bound to 5′-phosphate DNA (product complex) with a 5′-ϵA at 1.43 Å resolution (PDB entry 5HT2) and the abasic DNA damage mimic 5′-THF at 2.15 Å resolution (PDB entry 5INK; Figure 1D and E, Table 1). In these Tdp2-DNA complex structures, mTdp2cat adopts a mixed α-β fold typified by a central 12-stranded anti-parallel β-sandwich enveloped by several helical elements that mold the Tdp2 active site. One half of the molecule contributes to formation of the walls of the DNA-binding cleft that embraces the terminal position of the damaged DNA substrate. In the DNA lesion-bound state, two key DNA binding elements, the β-2-helix-β (β2Hβ) ‘grasp’, and ‘helical cap’ mold the substrate binding trench and direct the ssDNA of a 5′-overhang substrate into the active site. A comparison to an additional new structure of DNA-free Tdp2 (apo state, Figure 1F) shows that this loop is conformationally mobile and important for engaging DNA substrates. The mode of engagement of the 5′-nucleobase of the bulky ϵA adduct describes a mechanism for Tdp2 to bind 5′-tyrosylated substrates that contain diverse forms of DNA damage. The 5′-ϵA nucleobase is recognized by an extended Tdp2 van Der Waals interaction surface, referred to here as the ‘hydrophobic wall’ that is assembled with the sidechains of residues Leu315 and Ile317 (Figure 2A and B). Structures of mTdp2cat bound to DNA damage that triggers Top2 poisoning. (A) Structure of mTdp2cat bound to 5′-phosphate DNA (product complex) containing ϵA (yellow), Mg2+ (magenta) and its inner-sphere waters (gray). mTdp2cat is colored by electrostatic surface potential (red = negative, blue = positive, gray = neutral/hydrophobic). PDB entry 5HT2. (B) σ-A weighted 2Fo-Fc electron density map (at 1.43 Å resolution, contoured at 2.0 σ) for the ϵA DNA complex. The ϵA nucleotide is shown in yellow and a hydrogen bond from the ϵA O4′ to inner-sphere water is shown as gray dashes. (C) Structure of mTdp2cat bound to 5′-phosphate DNA (product complex) containing THF (yellow), Mg2+ (magenta) and its inner-sphere waters (gray). mTdp2cat is colored with red (electronegative), blue (electropositive) and gray (hydrophobic) electrostatic surface potential displayed. PDB entry 5INK is displayed. (D) σ-A weighted 2Fo-Fc electron density map (at 2.15 Å resolution, contoured at 2.0 σ) for THF-DNA complex. The THF is shown in yellow and a hydrogen bond from the THF O4′ to inner-sphere water is shown as gray dashes. For comparison, we also determined a structure of an undamaged 5′-adenine (5′-dA) bound to Tdp2 at 1.55 Å (PDB entry 5INL). A structural overlay of damaged and undamaged nucleotides shows no major distortions to nucleotide planarity between different bound sequences and DNA damage (compare ϵA, dA and dC, Supplementary Figure S1A–D). Therefore, structurally diverse undamaged or alkylated bases (e.g. ϵG, ϵT) could likely be accommodated in the Tdp2 active site via planar base stacking with the active site facing hydrophobic wall of the β2Hβ motif. Likewise, the abasic deoxyribose analog THF substrate binds similar to the alkylated and non-alkylated substrates, but with a slight alteration in the approach of the 5′-terminus (Figure 2C). Interestingly, in the absence of a nucleobase, O4′ of the THF ring adopts a close approach (2.8 Å) to a water molecule that directly participates in the outer sphere single Mg2+ ion coordination shell (Figure 2D). This shift is coincident with a small adjustment in the position of the +2 and +3 nucleotides (Supplementary Figure S1E). These collective differences may explain the slight, but statistically significant elevated activity on the THF substrate (Figure 1C). Structural plasticity in the Tdp2 DNA binding trench An intriguing feature of the DNA-damage bound conformation of the Tdp2 active site is an underlying network of protein–water–protein contacts that span a gap between the catalytic core and the DNA binding β2Hβ-grasp (Supplementary Figure S2). In this arrangement, six solvent molecules form a channel under the β2Hβ-grasp, ending with hydrogen bonds to the peptide backbone of the Mg2+ ligand Asp358. The paucity of hydrophobic interactions stabilizing the β2Hβ DNA-bound conformation suggests that conformational plasticity in the β2Hβ might be a feature of DNA damage and metal cofactor engagement. To test this hypothesis, we crystallized Tdp2 in the absence of DNA and determined a DNA free Tdp2 structure to 2.4 Å resolution (PDB entry 5INM; Figures 1F and 3A). Conformational plasticity in the Tdp2 active site. (A) The open, 3-helix conformation (tan) of flexible active-site loop observed in monomer E of the DNA-free mTdp2cat structure (PDB entry 5INM) is supported by T309 (green), which packs against the EEP core. The β2Hβ docking pocket (circled) is unoccupied and residues N312, N314 and L315 (orange) are solvent-exposed. Wall-eyed stereo view is displayed. (B) The closed β2Hβ conformation in the mTdp2cat–DNA product structure containing 5′-ϵA (yellow, PDB entry 5HT2). T309 (green) is an integral part of the β2Hβ DNA-binding grasp (tan) and hydrogen bonds to the backbone of Y321, while N314 (orange) occupies the β2Hβ docking pocket. Wall-eyed stereo view is displayed. (C) Alignment of active site loop conformers observed in the 5 promoters of the DNA-free mTdp2cat (PDB entry 5INM, see Table 1) crystallographic asymmetric unit (left) and sequence alignment showing residues not observed in the electron density as ‘∼’ (right). (D) Limited trypsin proteolysis probes the solvent accessibility of the flexible active-site loop. mTdp2cat WT (lanes 1–13) or mTdp2cat D358N (lanes 14–26) were incubated in the presence or absence of Mg2+ and/or a 12 nt self annealing, 5′-phosphorylated DNA (substrate ‘12 nt’ in Supplementary Table S1), then reacted with 0.6, 1.7 or 5 ng μl−1 of trypsin. Reactions were separated by SDS-PAGE and proteins visualized by staining with coomassie blue. (E) Limited chymotrypsin proteolysis probes the solvent accessibility of the flexible active-site loop. Experiments performed as in panel D for mTdp2cat WT (lanes 27–39) or mTdp2cat D358N (lanes 40–52), but with chymotrypsin instead of trypsin. This crystal form contains 5 Tdp2 protein molecules in the asymmetric unit, with variations in active site Mg2+ occupancy and substrate binding loops observed for the individual protomers. The most striking feature of the DNA ligand-free state is that the active site β2Hβ-grasp can adopt alternative structures that are distinct from the DNA-bound, closed β2Hβ DNA binding grasp (Figure 3A and B). In one monomer (chain ‘E’), the grasp adopts an ‘open’ 3-helix loop conformation that projects away from the EEP catalytic core. Two monomers have variable disordered states for which much of the DNA binding loop is not visible in the electron density. The remaining two molecules in the DNA-free crystal form are closed β2Hβ conformers similar to the DNA bound structures (Figure 3C). Thus, we posit that Tdp2 DNA binding conformationally selects the closed form of the β2Hβ grasp, rather than inducing closure upon binding. A detailed analysis of the extended 3-helix conformation shows that the substrate-binding loop is able to undergo metamorphic structural changes. In this open form, residues Asn312-Leu315 are distal from the active site and solvent-exposed (orange sticks, Figure 3A), while Thr309 (green surface, Figure 3A) packs into a shallow pocket of the EEP core to anchor the loop. Burial of Thr309 is enabled by an unusual main chain cis–peptide bond between Asp308-Thr309 and disassembly of the short antiparallel beta-strand of the β2Hβ fold. By comparison, the closed β2Hβ grasp conformer is stabilized by Asn312 and Asn314 binding into two β2Hβ docking pockets, and Leu315 engagement of the 5′-terminal nucleobase (Figure 3B). To transition into the closed β2Hβ conformation, Thr309 disengages from the EEP domain pocket, flips peptide backbone conformation cis to trans, and is integral to the β2Hβ antiparallel β-sheet. Stabilization of the closed β2Hβ-grasp conformation is linked to the active site through a hydrogen bond between Trp307 and the Mg2+ coordinating residue Asp358. Accordingly, in the DNA free structure, we observe a trend where the 2 closed monomers have an ordered Mg2+ ion in their active sites, while the monomers with open conformations have a poorly ordered or vacant metal binding site. Overall, these observations suggest that engagement of diverse damaged DNA ends is enabled by an elaborate substrate selected stabilization of the β2Hβ DNA binding grasp, and these rearrangements are coordinated with Mg2+ binding in the Tdp2 active site. To evaluate Mg2+ and DNA-dependent Tdp2 structural states in solution, we probed mTdp2cat conformations using limited trypsin and chymotrypsin proteolysis (Figure 3C–E). In the absence of DNA or Mg2+, mTdp2cat is efficiently cleaved in the metamorphic DNA binding grasp at one site by trypsin (Arg316), or at two positions by chymotrypsin (Trp307 and Leu315). By comparison, Mg2+, and to a greater extent Mg2+/DNA mixtures (compare Figure 3, lanes 4, 7 and 13) protect mTdp2cat from proteolytic cleavage. Interestingly, addition of Mg2+ alone protects against proteolysis as well. This is consistent with Mg2+ stabilizing the closed conformation of the β2Hβ-grasp through an extended hydrogen-bonding network with Asp358 and the indole ring of the β2Hβ-grasp residue Trp307 (also discussion below on Tdp2 active site SNPs). To assess structural conservation of Tdp2 conformational changes between human and mouse Tdp2, we also determined a 3.2 Å resolution structure of the human Tdp2cat domain bound to a DNA 5′-PO4 terminus product complex (PDB entry 5INO). Comparisons of the human hTdp2cat-DNA complex structure to the mTdp2cat DNA bound state show a high level of conservation of the DNA-bound conformations (Supplementary Figure S3A). Moreover, similar to mTdp2cat, proteolytic protection of the hTdp2cat substrate binding loop occurs with addition of Mg2+ and DNA (Supplementary Figure S3B). Thus, X-ray structures and limited proteolysis analysis indicate that DNA- and metal-induced conformational changes are a conserved feature of the vertebrate Tdp2-substrate interaction. Tdp2 metal ion dependence Consistently in high-resolution X-ray structural analyses we, and others observe a single Mg2+ metal bound in the Tdp2 active site. This includes the DNA-free (Figure 3A), DNA damage bound (Figure 3B) and reaction product-bound crystal forms of mouse, (PDB entry 4GZ1), D. rerio (PDB entry 4FPV) and C. elegans Tdp2 (PDB entry 4FVA). However, previous biochemical analysis has suggested an alternative two-metal ion mechanism for the Tdp2-phosphotyrosyl phosphodiesterase reaction. In these experiments, at limiting Mg2+ concentrations, Ca2+ addition to Tdp2 reactions stimulated activity. While this work was suggestive of a two metal ion mechanism for phosphotyrosyl bond cleavage by Tdp2, we note that second metal ion titrations can be influenced by metal ion binding sites outside of the active site. In fact, divalent metals have been observed in the Tdp2 protein–DNA complexes (PDB entry 4GZ2) distal to the active center, and we propose this might account for varied results in different studies. To further probe the metal ion dependence of the Tdp2 phosphodiesterase reaction, we performed metal ion binding assays, determined crystal structures in the presence of varied divalent metals (Mn2+ and Ca2+), and analyzed metal ion dependence of the Tdp2 phosphotyrosyl phosphodiesterase reaction (Figure 4). Metal cofactor interactions with Tdp2. (A) Intrinsic tryptophan fluorescence of mTdp2cat was used to monitor a conformational response to divalent metal ion binding. Either Mg2+ or Ca2+ were titrated in the presence or absence of 5′-P DNA, and the tryptophan fluorescence was monitored with an excitation wavelength of 280 nm and emission wavelength of 350 nm using 10 nm band pass filters. Both Mg2+ and Ca2+ induce a conformational change which elicits an increase in tryptophan fluorescence of mTdp2cat in the presence and absence of DNA, while D358N active site mutant of mTdp2cat is unresponsive to Mg2+. (B) mTdp2cat activity assayed on a T5PNP substrate as a function of Mg2+ and Ca2+ concentration. PNP release (monitored by absorbance at 415 nm) as a function of Mg2+ concentration and in the absence or presence of 1 or 10 mM Ca2+ is shown; error bars, s.d. n = 4. (C) σ-A weighted 2Fo-Fc electron density map (blue) and model-phased anomalous difference Fourier (magenta) maps for the mTdp2cat–DNA–Mn2+ complex (PDB entry 5INP) show a single Mn2+ (cyan) is bound with expected octahedral coordination geometry. A 53σ peak in the anomalous difference Fourier map (data collected at λ = 1.5418 Å) supports Mn2+ as the identity of this atom. (D) Comparison of Ca2+ (green Ca2+ ion, orange DNA) (PDB entry 5INQ), and Mg2+ (magenta Mg2+ ion, yellow DNA) (PDB entry 4GZ1) mTdp2cat–DNA structures shows that Ca2+ distorts the 5′-phosphate binding mode. Our proteolysis results indicate a Mg2+-dependent Tdp2 conformational response to metal binding. The Tdp2 active site has three tryptophan residues within 10 Å of the metal binding center, so we assayed intrinsic tryptophan fluorescence to detect metal-induced conformational changes in mTdp2cat. These data were an excellent fit to a single-site binding model both in the presence and absence of DNA (Figure 4A). This analysis revealed Mg2+ Kd values in the sub-millimolar range and Hill coefficients which were consistent with a single metal binding site both in the presence and absence of DNA (Supplementary Table S2). We then measured effects of metal ion concentrations on Tdp2 cleavage of p-nitrophenyl-thymidine-5′-phosphate by mTdp2cat. This small molecule substrate is not expected to be influenced by metal–DNA coordination outside of the active site. Inclusion of ultrapure Ca2+ (1 mM or 10 mM) results in a dose-dependent inhibition but not stimulation Tdp2 activity, even in conditions of limiting Mg2+ (Figure 4B). We performed the same titrations with human hTdp2FL and hTdp2cat (Supplementary Figure S4), and find similar stimulation of activity by Mg2+ and inhibition by Ca2+. Overall, these metal binding analyses are consistent with a single metal ion mediated reaction. To further evaluate the structural influence of divalent cations on the Tdp2 active site, we determined crystal structures by soaking crystals with metal cofactors that either support (Mn2+) or inhibit (Ca2+, Figure 4B) the Tdp2 reaction (PDB entries 5INP and 5INQ). Anomalous difference Fourier maps of the Tdp2–DNA–Mn2+ complex show a single binding site for Mn2+ in each Tdp2 active site (Figure 4C), with octahedral coordination and bond lengths typical for Mn2+ ligands (Supplementary Table S3). The Mn2+ ion is positioned in the Tdp2 active site similar to the Mg2+-bound complex (Figure 2C), which is consistent with the ability of Mn2+ to support robust Tdp2 catalytic activity. In contrast, while co-complex structures with Ca2+ also show a single metal ion, Ca2+ binds in a slightly different position, shifted ∼1 Å from the Mg2+ site. Although Ca2+ is also octahedrally coordinated, longer bond lengths for the Ca2+ ligands (Supplementary Table S3) shift the Ca2+ ion relative to the Mg2+ ion site. Interestingly, bi-dentate inner sphere metal contacts from the Ca2+ ion to Glu162 distort the active site phosphate-binding mode, and dislodge the 5′-PO4 out of the Tdp2 active site (Figure 4D). Together with results showing that under the conditions examined here, Ca2+ inhibits rather than stimulates the Tdp2 reaction, the divalent metal bound Tdp2 structures provide a mechanism for Ca2+-mediated inhibition of the Tdp2 reaction. Modeling the Tdp2 reaction coordinate Next, to examine the feasibility of our proposed single Mg2+ mechanism, we simulated the Tdp2 reaction coordinate with hybrid QM/MM modeling using Tdp2 substrate analog- and product-bound structures as guides. Previous structural analyses showed that the superposition of a DNA substrate mimic (5′-aminohexanol) and product (5′-PO4) complexes delineates a probable Tdp2 reaction trajectory characterized by inversion of stereochemistry about the adducted 5′-phosphorus. In this scheme (Figure 5A), a candidate nucleophilic water that is strongly hydrogen bonded to Asp272 and Asn274, is well positioned for the in-line nucleophilic attack ∼180° opposite of the P–O bond of the 5′-Tyr adduct. Structure-function analysis of the Tdp2 reaction mechanism. (A) Proposed mechanism for hydrolysis of phosphotyrosine bond by Tdp2. Residues in green form the binding-site for the 5′-tyrosine (red) and phosphate, yellow bind the 5′ nucleotide and blue bind nucleotides 2–3. Residue numbers shown are for the mTdp2 homolog. (B) Free energy during the QM/MM simulation as a function of distance between the nucleophilic water and 5′-phosphorus atom. Reaction proceeds from right to left. (C) Models for the mTdp2cat-DNA complex during the QM/MM reaction path simulation showing the substrate (left, tan), transition state intermediate (center, cyan) and product (right, pink) states. Residue numbers shown are for the mTdp2 homolog. (D) Electrostatic surface potential calculated for 5′-phosphotyrosine in isolation (upper panel) and in the presence of a cation–π interaction with the guanidinium group of Arg216 (lower panel) shows electron-withdrawing effect of this interaction. Electrostatic potential color gradient extends from positive (red) through neutral (gray), to negative (blue). (E) Bar graph displaying the relative activity of wild-type and mutant human MBP-hTdp2cat fusion proteins on the three substrates. Release of PNP from PNP phosphate and T5PNP was detected as an increase in absorbance at 415 nm. Reaction rates are expressed as the percent of activity relative to wildtype MBP-hTdp2cat; error bars, s.d. n = 3. Mutants of hTdp2 (black) and the equivalent residue in mTdp2 (tan) are indicated. We examined the energy profile of the nucleophilic attack of the water molecule by using the distance between the water oxygen and the P atom on the phosphate moiety as the sole reaction coordinate in the present calculation (Figure 5B and C). A starting model was generated from atomic coordinates of the mTdp2cat 5′–aminohexanol substrate analog structure (PDB 4GZ0) with a tyrosine replacing the 5′-aminohexanol then adding the Mg2+ and inner-sphere waters from the mTdp2-DNA product structure (PDB, 4GZ1), and running an initial round of molecular dynamics simulation (10 ns) to allow the system to reach an equilibrium. After QM/MM optimization of this model (Figure 5C, ‘i-substrate’), the O–P distance is 3.4 Å, which is in agreement with the range of distances observed in the mTdp2cat 5′-aminohexanol substrate analog structure (3.2–3.4 Å). No appreciable energy penalty is observed during the first 0.5 Å of the reaction coordinate. When the reaction reaches an O–P distance of 2.18 Å, formation of a transition state with an energy maximum of +7.4 kcal mol−1 is observed. Here, the water proton and the neighboring O of Asp272 participates in a strong hydrogen bond (distance of 1.58 Å) and the phosphotyrosyl O–P distance is stretched to 1.77 Å, which is 0.1 Å beyond an equilibrium bond length. In the subsequent two steps of the simulation, as the water-phosphate O–P distance reduces to 1.98 Å, a key hydrogen bond between the nucleophilic water and Asp272 shortens to 1.38 Å as the water H–O bond approaches the point of dissociation. The second proton on the water nucleophile maintains a strong hydrogen bond with Asn274 throughout the reaction, implicating this residue in orienting the water nucleophile during the reaction. Concomitant with this, the phosphotyrosyl O–P bond weakens (d = 1.89 Å), and the formation of the penta-covalent transition state (Figure 5C ‘ii-transition state’) is observed. The final steps show inversion of stereochemistry at the phosphate, along with lengthening and breaking of the phosphotyrosyl O–P bond. Product formation is coupled to a transfer of a proton from the nucleophillic water to Asp272, consistent with the proposed function for this residue as the catalytic base. Of note, both nitrogens of the imidazole side chain of His 359 require protonation for stability of the simulation. Asp 326 makes a hydrogen bond to N∂1 of His359, suggesting that this salt bridge could stabilize the protonated form of His359 as has been demonstrated for the analogous Asp-His pair in the EEP domain of APE1, which elevates the pKa of this His above 8.0. In our model, the transition state contains a hydrogen bond between the doubly protonated His359 and the phosphate oxygen that also coordinates with the single catalytic Mg2+, while the second His359 imidazole proton maintains a H-bond with the Asp326 residue throughout the reaction. In the final optimized structure, the observed product state (Figure 5C, ‘iii-product’) is found in a conformation that is 7.4 kcal mol−1 more stable than the initial reactive state (Figure 5B). The tyrosine oxy-anion product is coordinated to the Mg2+ ion with a 2.0 Å distance, which is the shortest of the six Mg2+ ligands (including three water molecules, one of the free oxygens on the phosphate group and the Glu162 residue), indicating the single Mg2+ greatly stabilizes the product oxy-anion. An additional striking feature gleaned from the QM/MM modeling is the putative binding mode of the Top2 tyrosine-leaving group. A trio of conserved residues (Tyr 188, Arg 216 and Ser 239) forms the walls of a conserved Top2 tyrosine binding pocket. We propose this cation–π interaction further contributes to tuned stabilization of the negatively charged phenolate reaction product. Consistent with this, analysis of electrostatic potential of the phosphotyrosyl moiety using Gaussian 09.D01 in the presence and absence of the Arg216 guanidinium reveals Arg216 is strongly electron withdrawing (Figure 5D). We further examined the contribution of this cation–π interaction to the reaction chemistry by moving the guanidinium group of Arg216 from the QM system to the MM system as either a +1 or ∼0 charge species, and re-computed energy penalties for each step in the reaction coordinate (Supplementary Figure S5A). Removing Arg216 from the quantum subsystem incurs an ∼2 kcal mol−1 penalty in the transition state and product complex. Removing the +1 charge on the Arg216 has a minimal impact on the transition state, but incurs an additional ∼2 kcal mol−1 penalty in the product complex. Altogether, QM/MM modeling identifies new determinants of the Tdp2 reaction, and demonstrates our proposed single Mg2+ catalyzed reaction model is a viable mechanism for Tdp2-catalyzed 5′-phosphotyrosine bond hydrolysis. Tdp2 mutational analysis To test the aspects of the Tdp2 reaction mechanism described here derived from high-resolution mouse Tdp2 crystal structures (denoted with superscript numbering ‘m’ for numbering of the mouse protein), we engineered and purified thirteen human MBP-hTdp2cat mutant proteins (denoted with superscript numbering and ‘h’ for the human protein) and assayed the impacts of mutations on Tdp2 catalytic activity using three in vitro reporter substrates including a tyrosylated DNA substrate (5′-Y), p-nitrophenyl phosphate (PNPP) and thymidine 5′-monophosphate p-nitrophenyl ester (T5PNP) (Figure 5E, Supplementary Figures S5B and S5C). By analyzing activities on this nested set of chemically related substrates we aimed to dissect structure-activity relationships of Tdp2 catalysis. For example, mutations impacting Tdp2 active site chemistry and phosphotyrosyl bond cleavage should similarly affect catalysis on all three substrates, but mutants impacting DNA damage binding might only impair catalysis on 5′-Y and T5PNP but not PNPP that lacks a nucleobase. Structural results and QM/MM modeling indicate mAsp272 activates a water molecule for in-line nucleophilic attack of the scissile phosphotyrosyl linkage. To test if this proposed Lewis base is critical for reaction chemistry we mutated it to a His, which could alternatively support metal binding, as well as bulky hydrophobic residues (Leu and Met) that we predict would block the water-binding site. Similar to a previously characterized hD262N mutation, all three substitutions ablate activity, supporting essential roles for hAsp262 (mAsp272) in catalysis. Next, we mutated key elements of the mobile loop (β2Hβ hydrophobic wall, Figure 2A and C). Mutations hI307A, hL305A, hL305F and hL305W all impaired catalysis on both nucleotide-containing substrates (\u003c50% activity). The hL305W substitution that we expect to have the most distorting impact on conformation of the β2Hβ hydrophobic wall also has the largest impact on catalysis of the DNA substrate 5′-Y. By comparison, as predicted by our model where β2Hβ dictates key interactions with undamaged and damaged nucleobases, all of these substitutions have little impact on PNPP (\u003e90% activity). Third, we altered properties of the proposed enzyme substrate cation–π interface. No activity was detected for a mutant that removes the positive charge at this position (hR206A). The precise geometry of this pocket is also critical for catalysis as replacement of hArg206 (mArg216) with a lysine also results in a profound decrease in catalysis (\u003c5% activity on 5′-Y, no detectable activity on T5PNP or PNPP). Similarly, mutation of hTyr178 that structurally scaffolds the hArg206 (mArg216) guanidinium also significantly impacts activity, with Y178F and Y178W having \u003c25% activity on all substrates. Fourth, we evaluated roles for the hHis351–hAsp316 (mAsp326–mHis359) transition state stabilization charge pair. We found that mutations that removed the charge yet retained the ability to hydrogen bond (hH351Q) or should abrogate the elevated pKa of the Histidine (hD316N) had severe impacts on catalysis. Thus altogether, our mutational data support key roles for the active site Lewis base aspartate, mobile substrate engagement loops, enzyme–substrate cation–π interactions, and active site transition state stabilizing charge interaction in supporting Tdp2 catalysis. A Tdp2 active site single nucleotide polymorphism impairs Tdp2 function Recently, it was found that inactivation of TDP2 by a splice-site mutation is associated with neurological disease and confers hypersensitivity to Top2 poisons. We considered whether human SNPs causing missense mutations might also impact Tdp2 DNA–protein crosslink repair functions established here as well as Tdp2-mediated NHEJ of blocked DNA termini. We identified two SNPs in human TDP2 curated in the NCBI SNP database that result in missense mutations within the DNA processing active site: rs199602263 (minor allele frequency 0.0002), which substitutes hAsp350 for Asn, and rs77273535 (minor allele frequency 0.004, which substitutes hIle307 for Val) (Figure 6A). We show the hD350N substitution severely impairs activity on all substrates tested in vitro, whereas hI307V only has a mild impact on catalysis (Figure 6B–D). To better understand the basis for the D350N catalytic defect, we analyzed the structural environment of this substitution based on the high-resolution structures of mTdp2cat (Figure 6A). Interestingly, the Tdp2 single Mg2+ ion octahedral coordination shell also involves an extended hydrogen-bonding network mediated by hAsp350 (mAsp358) that stabilizes the DNA-bound conformation of the β2Hβ substrate-binding loop through hydrogen bonding to mTrp307. Here, hAsp350 (mAsp358) serves as a structural nexus linking active site metal binding to substrate binding loop conformations. Tdp2 SNPs impair function. (A) Active site residues mutated by TDP2 SNPs. D350N (mTdp2 D358N) and I307V (mTdp2 I317V) substitutions are mapped onto the Tdp2 active site of the high-resolution mTdp2cat structure (4GZ1). (B) Coomassie blue stained SDS-PAGE gel of purified WT and mutant MBP-hTdp2cat proteins used for assays in panels C and D. (C) Activity of WT and mutant MBP-hTdp2cat proteins on a 5′–phosphotyrosyl–DNA oligonucleotides with 3′-fluorescein label. Samples were withdrawn from reactions, neutralized with TBE-urea loading dye at the indicated timepoints, and electrophoresed on a 20% TBE-urea PAGE. (D) Relative activity of WT and indicated mutant human MBP-hTdp2cat fusion proteins on three model Tdp2 substrates. Quantification of percent MBP-hTdp2cat activity relative to WT protein for the 5′-Y DNA oligonucleotide substrate (blue bars), T5PNP (red bars) and PNPP (green bars) is displayed. Release of PNP from PNP phosphate (PNPP) and was detected as an increase in absorbance at 415 nm, whereas the 5′-Y substrate is quantification of activity in a gel based assay shown in Figure 6C. Error bars, s.d. n = 3. To define the molecular basis for the hD350N (mD358N) defect, we crystallized and determined the structure of the DNA-free form of the mD358N protein to 2.8Å resolution (PDB entry 5INN). This structure shows the D358N mutation disrupts the hydrogen bond between Asp358 and Trp307, shifts the position of Asn358 and destabilizes Trp307. Consequently, poor electron density is visible for the β2Hβ loop which is mostly disordered (Supplementary Figure S6). Although Mg2+ is present at the same concentration as the WT-mTdpcat crystals (10 mM), we find the metal site is unoccupied in the mD358N crystals. Therefore, metal-regulated opening/closure of the active site may modulate Tdp2 activity, and D350N is sufficient to block both metal binding and conformational change. In support of this, we also find that hD350N (mD358N) impairs Mg2+ binding as measured by intrinsic tryptophan fluorescence (Figure 4A), and abrogates Mg2+-stimulated active site conformational changes detected by trypsin and chymotrypsin sensitivity of the Tdp2 metamorphic loop (Figure 3D). Tdp2 facilitates NHEJ repair of 5′-phosphotyrosine adducted DSBs Overall, our Tdp2 structure/activity studies reveal a tuned, 5′-detyrosylation DNA end processing activity and it has been demonstrated that Tdp2 could enable repair of Top2 damage by the non-homologous end-joining (NHEJ) pathway. Accordingly, we demonstrate here that 5′-tyrosylated ends are sufficient to severely impair an in vitro reconstituted mammalian NHEJ reaction (Figure 7A, lanes 3 and 6), unless supplemented with catalytic quantities of hTdp2FL (Figure 7A, lane 8). Interestingly, hTdp2cat is slightly more effective than hTdp2FL in promoting NHEJ of adducted ends, while a catalytically deficient E152Q mutant was inactive in this assay, supporting the notion that Tdp2 catalytic activity is required to support NHEJ of phosphotyrosyl blocked DSBs (Supplementary Figure S7A). We confirmed that efficient joining of the same tyrosine-adducted substrate in cells (Figure 7B) was dependent on both NHEJ (reduced over 10-fold in ligase IV deficient HCT 116 cells; Supplementary Figure S7B), and Tdp2 (reduced 5-fold in Tdp2 deficient MEFs; Figure 7C). Moreover, products with error (i.e. junctions have missing sequence flanking the adducted terminus) are twice as frequent in cells deficient in Tdp2 (Figure 7D). Therefore, in accord with previous work, joining of tyrosine adducted ends after Tdp2-mediated detyrosylation is both more efficient and more accurate than joining after endonucleolytic excision (e.g. mediated by Artemis or the Mre11/Rad50/Nbs1 complex). Effects of Tdp2 active site SNP-encoded mutants on cellular Tdp2 functions. (A) Cy5 labeled substrates with 5′-phosphate termini (Lanes 1–4) or 5′-tyrosylated termini (Lanes 5–9) were incubated with Ku, the NHEJ ligase (XRCC4, ligase IV and XLF; X-L-X) and 1 nM hTdp2FL as indicated (+) for 5 min at 37°C. Concatemer ligation products were detected by 5% native PAGE. (B) Workflow diagram of cellular end joining assays. DNA substrates with 5′-phosphotyrosine adducts and 4 nucleotide 5′ overhangs were electroporated into cultured mammalian cells. After 1 h, DNA was recovered from cells and repair efficiency by qPCR or sequencing as indicated. (C) qPCR assessment of cellular end joining efficiency of the tyrosylated substrate comparing results from wildtype MEF cells to Tdp2−/− cells and Tdp2−/− cells complemented with wildtype or the noted hTDP2FL variants; Joining efficiency shown is the ratio of junctions recovered relative to WT cells. Error bars, s.d, n = 3. (D) Junctions recovered from cellular end-joining assays in the noted cell types were characterized by sequencing to assess the end-joining error rate. Error bars, s.d, n = 3. (E) Clonogenic survival assay of WT, Tdp2 knockout and complemented MEF cells after treatment with indicated concentrations of etoposide for 3 h; error bars, s.d, n = 3. We next compared the ability of wild-type and mutant hTdp2FL variants to complement Tdp2 deficient mouse embryonic fibroblasts (Supplementary Figure S7C). Joining of extrachromosomal DNA with phosphotyrosine blocked ends, both in terms of efficiency (Figure 7C) and fidelity (Figure 7D), was indistinguishable comparing MEFs from a wild-type mouse, MEFs from a Tdp2-/- mouse overexpressing wild-type human Tdp2, and Tdp2 -/- MEFs overexpressing the I307V variant human Tdp2. In contrast, joining of 5′ phosphotyrosine-blocked ends was reduced 5-fold in Tdp2-/- MEFs, and an equivalent defect was observed in Tdp2-/- MEFs overexpressing Tdp2 D350N. Moreover, the frequency of inaccurate repair was 2-fold higher in both Tdp2 deficient cells and Tdp2 deficient cells overexpressing D350N, relative to cells expressing wild type Tdp2 or hTdp2 I307V (Figure 7D). Expression of wild type or I307V human Tdp2 in Tdp2-/- MEFs was also sufficient to confer levels of resistance to etoposide comparable to the matched wild-type MEF line, while overexpression of human D350N Tdp2 had no apparent complementation activity (Figure 7E). The rare D350N variant is thus inactive by all metrics analyzed. By comparison the more frequent I307V has only mild effects on in vitro activity, and no detectable impact on cellular assays. DISCUSSION Top2 chemotherapeutic agents remain frontline treatments, and exposure to the chemical and damaged DNA triggers of Top2-DNA protein crosslink formation are unavoidable. Understanding how cells cope with complex DNA breaks bearing topoisomerase–DNA protein crosslinks is key to deciphering individual responses to chemotherapeutic outcomes and genotoxic agents that poison Top2. Together with mutagenesis and functional assays, our new Tdp2 structures in the absence of ligands and in complex with DNA damage reveal four novel facets of Tdp2 DNA-protein conjugate processing: (i) The Tdp2 active site is well-suited for accommodating a variety of DNA structures including abasic and bulky alkylated DNA lesions that trigger Top2 poisoning, (ii) High-resolution structural analysis coupled with mutational studies and QM/MM molecular modeling of the Tdp2 reaction coordinate support a single metal-ion mechanism for the diverse clade of EEP domain catalyzed phosphoryl hydrolase reactions, (iii) The Tdp2 active site is conformationally plastic, and undergoes intricate rearrangements upon DNA and Mg2+ cofactor binding and (iv) Naturally occurring Tdp2 variants undermine Tdp2 active site chemistry, cellular and biochemical activities. This mechanistic dissection of Tdp2 interactions with damaged DNA and metal cofactor provides a detailed molecular understanding of the mechanism of Tdp2 DNA protein crosslink processing. Tdp2 was originally identified as a protein conferring resistance to both Top1 and Top2 anti-cancer drugs, however it is hypothesized that the predominant natural source of substrates for Tdp2 are likely the potent DNA damage triggers of Top2 poisoning and Top2 DNA protein crosslinks encountered during transcription. The properties of complex DNA strand breaks bearing Top2-DNA protein crosslinks necessitate that Tdp2 accommodates both damaged nucleic acid as well as the topoisomerase protein in its active site for catalysis. The Tdp2 substrate interaction groove facilitates DNA-protein conjugate recognition in two important ways. First, the nucleic acid binding trench is assembled by a dynamic β2Hβ DNA damage-binding loop that is capable of recognizing and processing diverse phosphotyrosyl linkages even in the context of bulky adducts such as ϵA. This is achieved by binding of nucleic acid ‘bases out’ by an extended base-stacking hydrophobic wall of the β2Hβ-loop. Secondly, our QM/MM analysis further highlights an enzyme–substrate cation–π interaction as an additional key feature of the Tdp2 protein–DNA crosslink binding and reversal. The strictly conserved active site Arg216 appears optimally positioned to stabilize a delocalized charge on the phenolate product of the phosphotyrosyl cleavage reaction through molecular orbital overlap and polarization of the leaving group. To our knowledge, this is the first proposed example of a substrate cation–π interface exploited to promote a phosphoryl-transfer reaction. This unique feature likely provides an additional level of substrate-specificity for Tdp2 by restricting activity to hydrolysis of aromatic adducts characteristic of Top2cc, picornaviral protein–RNA and Hepatitis B Virus (HBV) protein–DNA processing intermediates. By comparison, other EEP nucleases such as Ape1 and Ape2 have evolved robust DNA damage specific endonucleolytic and exonucleolytic activities not shared with Tdp2. The dynamic nature of the Tdp2 active site presents opportunities for enzyme regulation. However, whether additional protein factors can bind to Tdp2 and modulate assembly/disassembly of the Tdp2 β2Hβ-loop is unknown. We hypothesize that binding of the Top2 protein component of a DNA–protein crosslink and/or other protein-regulated assembly of the Tdp2 active site might also serve to regulate Tdp2 activity to restrict it from misplaced Top2 processing events, such that it cleaves only topologically trapped or poisoned Top2 molecules when needed. Furthermore, high-resolution structures of mouse (Figures 3 and 4) and C. elegans Tdp2 show that a single metal ion typifies the Tdp2 active site from worms to man. Herein, we report five additional lines of evidence from metal binding detected by intrinsic tryptophan fluorescence, crystallographic analysis of varied metal cofactor complexes, mutagenesis, Ca2+ inhibition studies and QM/MM analysis that all support a feasible single Mg2+ mediated Tdp2 catalytic mechanism. The advent of personalized medical screening opens doors for assessment of individual vulnerabilities to commonly used chemotherapeutic drugs. It would be beneficial to employ this knowledge during the early decision making processes regarding treatment. Etoposide and other Top2 poisons remain front line anti-cancer drugs, and Tdp2 frameshift mutations in the human population confer hypersensitivity to Top2 poisons including etoposide and doxyrubicin. Given Tdp2 variation in the human population, links to neurological disease and viral pathogenesis, our finding that TDP2 SNPs ablate catalytic activity has probable implications for modulation of cancer chemotherapy, susceptibility to environmentally linked Top2 poisons, and viral infection. Lastly, Tdp2 inhibitors may synergize or potentiate cytotoxic effects of current anticancer treatments that target Tdp2. Thus, we anticipate this atomic-level and mechanistic definition of the molecular determinants of Tdp2 catalysis and conformational changes driven by DNA–protein and protein–protein interactions will foster unique strategies for the development of Tdp2 targeted small molecule interventions. ACCESSION NUMBERS Coordinates and structure factors have been deposited in the RCSB Protein Data Bank under accession code 5HT2 (mTdp2-Mg-ϵA-DNA complex), 5INK (mTdp2-Mg-THF complex), 5INL (mTdp2-Mg-dA-DNA-product complex), 5INM (mTdp2-apo structure), 5INN (mTdp2-D350N structure), 5INO (hTdp2-Mg-DNA product complex), 5INP (mTdp2-Mn-DNA product complex) and 5INQ (mTdp2-Ca-DNA product complex). Supplementary Material SUPPLEMENTARY DATA Supplementary Data are available at NAR Online. FUNDING US National Institute of Health Intramural Program; US National Institute of Environmental Health Sciences (NIEHS) [1Z01ES102765 to R.S.W.]; National Cancer Institute (NCI) [R01 CA084442 to D.A.R]. Use of the APS was supported by the U. S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. W-31-109-Eng-38. Funding for open access publication: NIEHS [1Z01ES102765 to R.S.W]. Conflict of interest statement. None declared. 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diff --git a/annotated_BioC_JSON/PMC4869123_ann.json b/annotated_BioC_JSON/PMC4869123_ann.json
new file mode 100644
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+[{"sourceid":"4869123","sourcedb":"","project":"","target":"","text":"Inhibiting complex IL-17A and IL-17RA interactions with a linear peptide IL-17A is a pro-inflammatory cytokine that has been implicated in autoimmune and inflammatory diseases. Monoclonal antibodies inhibiting IL-17A signaling have demonstrated remarkable efficacy, but an oral therapy is still lacking. A high affinity IL-17A peptide antagonist (HAP) of 15 residues was identified through phage-display screening followed by saturation mutagenesis optimization and amino acid substitutions. HAP binds specifically to IL-17A and inhibits the interaction of the cytokine with its receptor, IL-17RA. Tested in primary human cells, HAP blocked the production of multiple inflammatory cytokines. Crystal structure studies revealed that two HAP molecules bind to one IL-17A dimer symmetrically. The N-terminal portions of HAP form a β-strand that inserts between two IL-17A monomers while the C-terminal section forms an α helix that directly blocks IL-17RA from binding to the same region of IL-17A. This mode of inhibition suggests opportunities for developing peptide antagonists against this challenging target. The family of IL-17 cytokines and receptors consists of six polypeptides, IL-17A-F, and five receptors, IL-17RA-E. IL-17A is secreted from activated Th17 cells, and several innate immune T cell types including macrophages, neutrophils, natural killer cells, and dendritic cells. IL-17A signals through a specific cell surface receptor complex which consists of IL-17RA and IL-17RC. IL-17A’s downstream signaling leads to increased production of inflammatory cytokines such as IL-6, IL-8, CCL-20 and CXCL1 by various mechanisms including stimulation of transcription and stabilization of mRNA. Although various cell types have been reported to express IL-17RA, the highest responses to IL-17A come from epithelial cells, endothelial cells, keratinocytes and fibroblasts. IL-17A and its signaling is important in host defense against certain fungal and bacterial infections as demonstrated by patients with autoantibodies against IL-17A and IL-17F, or with inborn errors of IL-17 immunity. In addition to its physiological role, IL-17A is a key pathogenic factor in inflammatory and autoimmune diseases. In phase II and III clinical trials, neutralizing monoclonal antibodies against IL-17A (secukinumab and ixekizumab) or its receptor IL-17RA (brodalumab) are highly efficacious in treating moderate to severe plaque psoriasis and psoriatic arthritis. Secukinumab has been approved recently as a new psoriasis drug by the US Food and Drug Administration (Cosentyx™). In addition to psoriasis and psoriatic arthritis, IL-17A blockade has also shown preclinical and clinical efficacies in ankylosing spondylitis and rheumatoid arthritis. Among IL-17 cytokines, IL-17A and IL-17F share the highest homology. These polypeptides form covalent homodimers, and IL-17A and IL-17F also form an IL-17A/IL-17F hetereodimer. Structures are known for apo IL-17F and its complex with IL-17RA, for apo IL-17A, its complex with an antibody Fab, and its complex with IL-17RA. In these structures, both IL-17A and IL-17F adopt a cysteine-knot fold with two intramolecular disulfides and two interchain disulfide bonds that covalently link two monomers. There has been active research in identifying orally available chemical entities that would functionally antagonize IL-17A-mediated signaling. Developing small molecules targeting protein-protein interactions is difficult with particular challenges associated with the large, shallow IL-17A/IL-17RA interfaces. Since IL-17RA is a shared receptor for at least IL-17A, IL-17F, IL-17A/IL-17F and IL-17E, we chose to seek IL-17A-specific inhibitors that may have more defined pharmacological responses than IL-17RA inhibitors. Our initial approach was to identify peptide inhibitors which could serve as leads for the development of anti-inflammatory therapeutics that could be used alone or in combination with other agents. Our efforts resulted in discovery of a high affinity IL-17A peptide antagonist (HAP), which we attempted to increase the functional production and pharmacokinetics after fusing HAP to antibodies for evaluation as a bispecific therapeutic in animal studies. Unfortunately, this past work revealed stability issues of the uncapped HAP in cell culture Here, we provide the details of the discovery and optimization that led to HAP and report the complex structure of IL-17A with HAP, which provides structure based rationalization of peptide optimization and structure activity relationship (SAR). Results Identification of IL-17A peptide inhibitors Peptides specifically binding to human IL-17A were identified from phage panning using cyclic and linear peptide libraries (Supplementary Figure S1). Positive phage pools were then sub-cloned into a maltose-binding protein (MBP) fusion system. Single clones were isolated and sub-cultured in growth medium, and culture supernatants were used in an enzyme-linked immunosorbent assay (ELISA) to identify specific IL-17A-binding clones. The positive binding supernatants were tested for the ability to block biotinylated IL-17A signaling through IL-17RA in an IL-17A/IL-17RA competition ELISA assay where unlabeled IL-17A was used as positive control to inhibit biotinylated IL-17A binding. Approximately 10% of the clones that specifically bound to IL-17A also prevented the cytokine from binding to IL-17RA. Sequences identified from phage clones were chemically synthesized (Supplementary Table 1) and tested for inhibition of IL-17A binding to IL-17RA (Table 1). A 15-mer linear peptide 1 was shown to block IL-17A/IL-17RA binding with an IC50 of 80 nM in the competition ELISA assay (Table 1). This peptide was then tested in a cell-based functional assay wherein production of GRO-α in BJ human fibroblast cells was measured as a function of IL-17A stimulation using 1 ng/ml IL-17A. Peptide 1 was found to be active in this functional assay with an IC50 of 370 nM. Optimization of IL-17A peptide inhibitors A SAR campaign was undertaken to improve the potency of peptide 1. An alanine scan of peptide 2, an analogue of 1 with a lysine to arginine substitution at position 14, was initiated. When alanine was already present (positions 7 and 15), substitution was made with lysine (Table 1, peptides 3–17). Positions 1, 2, 4, 5, 7, 14 and 15 were shown to be amenable to substitution without significant loss (less than 3-fold) of binding affinity as measured by the IL-17A/IL-17RA competition ELISA. In particular, at position 5 (13), substitution of methionine with alanine resulted in a seven fold improvement in potency (80 nM versus 11 nM respectively). In order to rapidly evaluate the effects of substitution of natural amino acids at tolerant positions identified by the alanine scan, the lead sequence was subjected to site-specific saturation mutagenesis using MBP. Each of the seven positions identified by the alanine scan was individually modified while keeping the rest of the sequence constant. Modifications at positions 2 and 14 were shown to display improvement in binding affinity (data not shown). Peptides with beneficial point mutations at positions 2, 5, and 14 were synthesized and evaluated in the competition ELISA (Table 1). Two of the changes, V2H (18) or V2T (21) displayed improved binding in the competition ELISA. Since the replacement of methionine at position 5 with alanine was beneficial, the additional hydrophobic amino acids isoleucine (24), leucine (25) and valine (26) were evaluated and an additional two-three fold improvement in binding was observed for the valine and isoleucine replacements in comparison with alanine. Introduction of a methionine (27) or a carboxamide (28 and 29) at position 14 was shown to improve the binding affinity of the lead peptide. In general, there was good agreement between the respective binding affinities of the synthesized peptides and their MBP fusion counterparts, except for substitution of valine at position 2 to a tryptophan (22), which resulted in a fivefold loss of affinity, for the free peptide when compared with the MBP fusion. Combining the key amino-acid residues identified by SAR into a single peptide sequence resulted in peptide 30, named high affinity peptide (HAP), that was found to inhibit IL-17A signaling in a BJ human fibroblast cell assay with an IC50 of 17 nM, a more than 20-fold improvement over the phage peptide 1 (Table 2 and Supplementary Figure S2). We also examined the effect of removing the acetyl group at the N-terminus of HAP (which is present in all the peptides made, see Supplementary Material). The un-capped peptide (31) had an IC50 of 420 nM in the cell-based assay. The loss of cellular activity of 31 was most likely due to the degradation of the N-terminus of 31, since peptide 31 was shown to be able to bind to IL-17A with similar affinity as HAP itself. Furthermore, our previous work had reported that in antibody fusions the uncapped peptide was degraded under cell assay conditions with removal of the first 1-3 residues to inactive products with the same N-terminal sequences as peptides 32–34. In this work, 32–34 are capped by protective acetyl group and reflect the same inactivity as reported. C-terminal truncations showed a more gradual reduction in activity (35–37; Table 2). After deletion of three amino acids from the C-terminal end (37), the peptide is no longer active. Dimerization of HAP can further increase its potency We reasoned that since the IL-17A protein is almost exclusively present in a dimeric form, dimerizing the IL-17A binding peptides could result in an improvement in binding affinity and inhibitory activity. Homodimers of HAP were made through attachment of polyethylene glycol (PEG) spacers of different lengths at amino acids 4, 7 and 14, as these positions were identified in the alanine scan analysis as not contributing significantly to the activity, and at each N-terminus (Supplementary Table S2). Due to the high reactivity of the pentafluoroester (PFP) group used as the activating group in the PEG, the histidine at position 2 and the lysine at position 15 were replaced with threonine and dimethyllysine respectively to prevent formation of side products, which resulted in peptide 38 that was comparable in activity with HAP. This exercise revealed that several dimeric peptides with the longer PEG21 spacer were significantly more potent than the monomer peptide in the cell-based assay (Supplementary Table S2). Peptide 45, dimerized via attachment of a PEG21 spacer at position 14 (Supplementary Scheme S1 and Figure S3), was the most potent with cellular IC50 of 0.1 nM. This significant improvement in antagonism was not seen in the peptide monomer functionalized with a PEG21 group at position 14 as peptide 48 had an IC50 of 21 nM (Supplementary Scheme S2). The species cross-reactivity of the dimeric peptide 45 and HAP were assessed in a murine functional cell assay using 15 ng/ml murine IL-17A. Peptide 45 blocked the receptor binding of murine IL-17A although with potency two orders of magnitude weaker than that observed against human IL-17A (IC50 = 41 nM vs IC50 = 0.1 nM, respectively). The monomer HAP was much weaker (IC50 \u003e1 μM) in inhibiting murine IL-17A signaling (Supplementary Figure S4). Although the dimeric peptide 45 is much more potent than HAP in the cell-based assay, in subsequent studies we decided to focus our efforts solely on characterizations of the monomeric peptide HAP in hopes to identify smaller peptide inhibitors containing the best minimal functional group. Orthogonal assays to confirm HAP antagonism To further characterize the interaction of HAP with IL-17A, we set out to determine its in vitro binding affinity, specificity and kinetic profile using Surface Plasmon Resonance (SPR) methods (Fig. 1A). HAP binds to immobilized human IL-17A homodimer tightly (Table 3). It has slightly weaker affinity for human IL-17A/F heterodimer and \u003e10 fold weaker affinity for mouse IL-17A (Table 3). HAP does not show significant binding to immobilized human IL-17F homodimer or IL-17RA at concentrations up to 100 nM. Additionally, we investigated the antagonism of the human IL-17A/IL-17RA interaction by HAP using orthogonal methods including SPR and Förster resonance energy transfer (FRET) competition assays (Fig. 1B,C). In both assays, incubation of IL-17A with HAP effectively blocks the binding of IL-17A to immobilized IL-17RA with similar sub-nM IC50 (Table 3). HAP blocks IL-17A signaling in a human primary cell assay While either IL-17A or TNF-α alone can stimulate the release of multiple inflammatory cytokines, when acting together they can synergistically enhance each other’s effects (Supplementary Figure S5). These integrative responses to IL-17A and TNF-α in human keratinocytes have been reported to account for key inflammatory pathogenic circuits in psoriasis. Thus, we chose to study HAP’s efficacy in blocking the production of IL-8, IL-6 and CCL-20 by primary human keratinocytes stimulated by IL-17A in the presence of TNF-α, an assay which may be more disease-relevant. HAP inhibits the production of all three cytokines in a dose-dependent fashion (Fig. 1D). Significantly, the baseline levels of IL-8, IL-6 and CCL-20 stimulated by TNF-α alone are not inhibited by HAP, further indicating the selectivity of HAP (Fig. 1D). Such pharmacological selectivity may be important to suppress inflammatory pathogenic circuits in psoriasis, while sparing the anti-infectious immune responses produced by TNF-α. The relatively high IC50 values in this assay (Table 3) are probably due to the high IL-17A concentration (100 ng/ml) needed for detection of IL-6. As a reference, a commercial anti-IL-17A antibody (R\u0026D Systems) inhibits the production of IL-8 with an IC50 of 13(±6) nM (N = 3). Indeed, the IC50 was 14(±9) nM (N = 12) for HAP inhibition of IL-8 production when only 5 ng/ml IL-17A was used in this assay. In patients, the concentration of IL-17A in psoriatic lesions is reported to be 0.01 ng/ml, well below the EC50 (5–10ng/ml) of IL-17A induced IL-8 production in vitro. Similar to keratinocytes assay results, while HAP inhibits IL-17A stimulated IL-6 production by BJ human fibroblast potently (IC50 of 17 nM), it does not inhibit TNF-α stimulated IL-6 production at concentrations up to 10 μM (Supplementary Figure S2). Crystallization and structure determination Extensive crystallization trials, either by co-crystallization or by soaking HAP into preformed apo IL-17A crystals, failed to lead to an IL-17A/HAP complex crystals. We theorized that HAP binding induced large conformational changes in IL-17A that led to the difficulty of getting an IL-17A/HAP binary complex crystal. It is known that an antibody antigen-binding fragment (Fab) can be used as crystallization chaperones in crystallizing difficult targets. We hypothesized that HAP may target the N-terminal of IL-17A which is known to be more flexible than its C-terminal and conformational changes needed for HAP binding may be more likely there. We designed an antibody Fab known to target the C-terminal half of IL-17A based on a published IL-17A/Fab complex crystal structure, and produced it in HEK293 cells. In an SPR assay HAP and this Fab were able to co-bind IL-17A without large changes in their binding affinities and kinetics, confirming our hypothesis (Supplementary Figure S6). Furthermore, since it binds to an area far away from that of HAP (see below), this Fab should have minimum effects on HAP binding conformation. Crystals of Fab/IL-17A/HAP ternary complex were obtained readily in crystallization screens. Crystallization of IL-17A and its binding partners was accomplished using two forms of IL-17A. These were, respectively, a presumably more homogeneous form of IL-17A that lacked the disordered N-terminal peptide and a full-length form of the cytokine with a full complement of disulfide bonds. (see Method). Crystals of the Fab/truncated IL-17A/HAP complex diffracted to 2.2 Å, and the Fab/full length IL-17A/HAP complex diffracted to 3.0 Å (Supplementary Table S3). Both structures were solved by molecular replacement. Both complexes crystallized in the space group of P321, with half the complex (1 Fab/1 IL-17A monomer/1 HAP) in the asymmetric unit. The intact complex can be generated by applying crystallographic 2-fold symmetry. Electron densities for HAP residues Ile1-Asn14 were readily interpretable with the exception of Lys15, which is disordered. When considering the protein, the complex structure containing the full length IL-17A is identical to that of the truncated IL-17A, with the exception of Cys106 (Ser106 in the truncated IL-17A), which is disordered. Cys106 is covalently linked to Cys10 that resides in the disordered N-terminal peptide in the full length IL-17A. Overall structure of Fab/IL-17A/HAP complex In a similar manner to the published structure of Fab/IL-17A complex, two Fab molecules bind symmetrically to the C-terminal of the cytokine dimer, interacting with epitopes from both monomers (Fig. 2A). Two copies of HAP bind to the N-terminal of the cytokine dimer, also symmetrically, and each HAP molecule also interacts with both IL-17A monomers (Fig. 2). Based on disclosed epitopes of Secukinumab and Ixekizumab, HAP binds to IL-17A at an area that is also different from those of those two antibodies. The N-terminal 5 residues of HAP, 1IHVTI, form an amphipathic β-strand that inserts between β-strand 4 of one IL-17A monomer and β-strand 0 (the first ordered peptide of IL-17A) of the second monomer. This β-strand is parallel to both strands 0 and 4 (Fig. 3B). Strands 0 of two IL-17A monomer are antiparallel, as appeared in other IL-17A structures. The C-terminal 8 residues of the HAP that are ordered in the structure, 7ADLWDWIN, form an amphipathic α-helix interacting with the second IL-17A monomer. Pro6 of HAP makes a transition between the N-terminal β-strand and the C-terminal α-helix of HAP. As a comparison, an IL-17A/IL-17RA complex structure (PDB code 4HSA) is also shown with IL-17A in the same orientation (Fig. 2C). Inhibition mechanism of IL-17A signaling by HAP IL-17RA binds IL-17A at three regions on the IL-17A homodimer. HAP binds IL-17A at region I. Region I is formed by residues at the ends of β strands 0 and 4, and from loops 1–2 and 3–4 of IL-17A (Fig. 2). Conformational changes in region I induced by HAP binding alone may allosterically affect IL-17RA binding, but more importantly, the α-helix of HAP directly competes with IL-17RA for binding to IL-17A (Fig. 3). The most significant interactions between the α helix of HAP and IL-17A involve Trp12 of HAP, which binds in a hydrophobic pocket in IL-17A formed by the side chains of Phe110, Tyr62, Pro59 and the hydrophobic portion of the Arg101 side chain (Fig. 3A). The Trp12 side chain of HAP donates a hydrogen bond to the main chain oxygen of Pro69 of IL-17A. The positively charged Arg101 side chain of the IL-17A engages in a charge-helix dipole interaction with the main chain oxygen of Trp12. Additionally, Leu9 and Ile13 of the HAP have hydrophobic interactions with IL-17A, and the Asp8 side chain has hydrogen bond and ion pair interactions with Tyr62 and Lys114 of IL-17A, respectively. In region I, an IL-17RA peptide interacts with IL-17A in a very similar fashion to the α-helix of HAP. The IL-17RA peptide has sequences of 27LDDSWI, and part of the peptide is also α-helical (Fig. 3B). Leu7, Trp31 and Ile32 of IL-17RA interact very similarly with the same residues of IL-17A as Leu9, Trp12 and Ile13 of HAP (Fig. 3B). In this sense, the α-helix of HAP with a sequence of 9LWDWI is a good mimetic of the 27LDDSWI peptide of IL-17RA. The β-strand of HAP has no equivalent in IL-17RA. However, it mimics the β-strand 0 of IL-17A. The amphipathic β-strand of HAP orients the hydrophilic side chains of His2 and Thr4 outwards, and the hydrophobic side chains of Ile1, Val3 and Ile5 inward (Fig. 3A). β-strand 0 in IL-17A is also amphipathic with the sequence of 21TVMVNLNI. In all IL-17A structures obtained to date, β-strand 0 orients the hydrophilic side chains of Thr21, Asn25 and Asn27 outward, and the hydrophobic side chains of Val22, Val24, Leu26 and Ile28 inward. The binding pocket occupied by either Trp12 of HAP or Trp31 of IL-17RA is not formed in the apo IL-17A structure (Fig. 3C). Conformational changes of IL-17A are needed for both HAP and IL-17RA to bind to that region. Particularly for HAP, β-strands 0 have to shift out of the hydrophobic cleft formed by the main body of the IL-17A by as much as 10 Å between Cα atoms (Fig. 3C). Disruptions of the apo IL-17A structure by HAP binding are apparently compensated for by formation of the new interactions that involve almost the entire HAP molecule (Fig. 3B). Structure basis for the observed SAR of peptides The IL-17A/HAP complex structure obtained is very consistent with the observed SAR of our identified peptide inhibitors, explaining well how the evolution of the initial phage peptide 1 to HAP and 45 improved its potency (Supplementary Figure S7). The important interactions involving Trp12 of HAP explain the \u003e90 times drop in potency of the W12A variant (6 vs 1, Table 1). The amphipathic nature of the HAP β-strand explains the preference of the hydrophilic residues at the 2 and 4 positions of peptides (14, 18, 19, 21 and 23 vs 1 and 22, Table 1). All N-terminal residues of HAP are part of the β-sheet with β-stands 0 and 4 of IL-17A, which explains why removal of the first 1–3 residues completely abolishes the ability of HAP to block IL-17A cell signaling (31,32 and 33, Table 2). The C-terminal Asn14 and Lys15 of HAP are not directly involved in interactions with IL-17A, and this is reflected in the gradual reduction in activity caused by C-terminal truncations (35 and 36, Table 2). Each peptide monomer in 45 may not necessarily be more potent than HAP, but two monomer peptides within the same molecule that can simultaneously bind to IL-17A can greatly improve its potency due to avidity effects. HAP targets region I of IL-17A, an area that has the least sequence conservation in IL-17 cytokines. This lack of sequence conservation in the HAP binding site explains the observed specificity of HAP binding to human IL-17A. For example, inspection of the published IL-17F crystal structure (PDB code 1JPY) revealed a pocket of IL-17F similar to that of IL-17A for W12 of HAP binding, but it is occupied by a Phe-Phe motif at the N-terminal peptide of IL-17F. This Phe-Phe motif is missing in IL-17A. Sequence alignments between human and mouse IL-17A indicated that among IL-17A residues that interacting with HAP, majority differences occur in strand 0 of IL-17A which interacts with the N-terminal β-strand of HAP. In human IL-17A the sequences are 21TVMVNLNI, and in mouse they are 21NVKVNLKV. Discussion Using a combination of phage display and SAR we have discovered novel peptides that are IL-17A antagonists. One of those peptides, HAP, also shows activity in inhibiting the production of multiple inflammatory cytokines by primary human keratinocytes stimulated by IL-17A and TNF-α, a disease relevant-model. We have also determined the complex structure of IL-17A/HAP, which provides the structural basis for HAP’s antagonism to IL-17A signaling. During IL-17A signaling, IL-17A binds to one copy of IL-17RA and one copy of IL-17RC. Since apo IL-17A is a homodimer with 2 fold symmetry, IL-17RA potentially can bind to either face of the IL-17A dimer. With two HAP molecules covering both faces of the IL-17A dimer, HAP can block IL-17RA approaching from either face. To form the 1:2 complex observed in crystal structure, it is important that there is no strong negative cooperativity in the binding of two HAP molecules. In fact, in native electrospray ionization mass spectrometry analysis only 1:2 IL-17A/HAP complex was observed even when IL-17A was in excess (Supplementary Figure S8), indicating a positive binding cooperativity that favors inhibition of IL-17RA binding by HAP. HAP, with only 15 residues, can achieve almost the same binding affinity as the much larger IL-17RA molecule, indicating a more efficient way of binding to IL-17A. The interaction of IL-17A with IL-17RA has an extensive interface, covering ~2,200 Å2 surface area of IL-17A. Due to the discontinuous nature of the IL-17A/IL-17RA binding interface, it is classified as having tertiary structural epitopes on both binding partners, and is therefore hard to target using small molecules. Our studies of HAP demonstrated an uncommon mode of action for a peptide in inhibiting such a difficult protein-protein interaction target, and suggest further possible improvements in its binding potency. One way of further improving HAP’s potency is by dimerization. Homo-dimerization of HAP (45) achieved sub-nanomolar potency against human IL-17A in cell assay. In the crystal structure, the distance between the carbonyl of Asn14 of one HAP molecule and the N-terminus of the second is only 15.7 Å, suggesting the potential for more potent dimeric peptides to be designed by using linkers of different lengths at different positions. Another direction of improving HAP is by reducing its size. As demonstrated by the crystal structure, binding of the α-helix of HAP should be sufficient for preventing IL-17RA binding to IL-17A. Theoretically, it is possible to design chemicals such as stapled α-helical peptides to block α-helix-mediated IL-17A/IL-17RA interactions. Such peptides may have smaller sizes with more favorable physical properties (proteolytic resistance, serum half-life, permeability, etc). In summary, these peptide-based anti-IL-17A modalities could be further developed as alternative therapeutic options to the reported monoclonal antibodies. We are also very interested in finding non-peptidic small molecule IL-17A antagonists, and HAP can be used as an excellent tool peptide. The strategy utilized in generating the complex structures of HAP may also be useful for enabling structure based design of some known small molecule IL-17A antagonists. Methods Peptide synthesis Peptides were synthesized either on a PTI Symphony or Biotage Syro II synthesizer employing standard Fmoc chemistry on Rink resin with N-terminal amine capped with acetyl group unless noted otherwise. Resin, amino acids and solvents were purchased from EMD Chemicals. Peptides were then removed from polymer support using the cleavage cocktail (88:5:5:2, TFA:PhOH:H2O:TIPS) and precipitated with cold diethyl ether. For the dimers, the native peptides incorporating a primary amine (N-terminal amine or lysine at specific positions in the peptide sequence) were mixed with the bis-pentafluoroesters of the desired PEG (purchased from Quanta BioDesign) in the presence of DIEA in DMF. Peptides were purified by reversed phase HPLC on Phenomenex preparative Luna C18 columns using water:acetonitrile gradients and lyophilized. HPLC-MS conditions for peptide purity analysis All peptides were assessed for purity by analytical C18 RP-HP-LCMS prior to use in biological assays. Buffers used were 0.1% trifluoroacetic acid in water (A) and 0.1% trifluoroacetic acid in acetonitrile (B). The standard method (1) consisted of a linear gradient of 5% to 95% B over 10 minutes on Agilent 1100 series HPLC-MSD and method (2) consisted of a linear gradient of 5% to 95% B over 8 minutes on Waters 2767 series HPLC-MSD. The C18 column (Phenomenex, Luna C18, 4.6 × 150mm) effluent was immediately mass analyzed in electrospray positive mode. Accurate mass measurements of final peptides were performed using C18 reversed-phase chromatography mass spectrometry (RPHPLCMS) and mass detected on a Waters Synapt G2 Q-Tof mass spectrometer tuned to a resolution (FWHM) of 25,000. Exact intact masses were calculated based on the monoisotopic m/z value of the base peak charge state. All peptides were analyzed using these methods. Supporting data of peptide 45 is shown (Supplementary Figure S2) as representative data sets of the all the molecules investigated (characterization of all peptides presented on Table 1S). ELISA assay rhIL-17 (#317-ILB), IL-17R (#2269-IL) were from R\u0026D Systems. ELISA wash buffer (50-63-01) and TMB SureBlue Microwell Peroxidase substrate (#50-63-01 or #52-00-00) were from KPL. Half-well high-binding ELISA plates (#3690) and full-well plates (#9018) were from Costar. Superblock (#2011 or AAA500) was from ScyTek. No-Weigh NSH-PEG4-Biotin (21329) and Zeba Desalt Spin Columns (89882) were from Pierce. ELISAs were as follows in duplicate unless otherwise specified. Plates were coated in PBS overnight at 4 °C and blocked using Superblock for one hour at RT. Subsequent incubations were for 1 hr at room temperature with dilutions in Superblock. Plates were washed 3X between steps using a Biotek ELx405 plate washer and developed using TMB substrate. Reactions were stopped in 2 M H2SO4 and OD450 values were read on a Molecular Devices SpectraMax Plus plate reader. IL-17 RA competition: half-well plates were coated with 0.5 μg/mL IL-17R-Fc and blocked. Peptides were titrated in Superblock containing 0.4 μg/mL Biotinylated IL-17A. Biotinylated IL-17A was detected by adding tetramethylbenzidine (TMB). Reactions were stopped in 2 M H2SO4 and OD450 values were read on a Molecular Devices SpectraMax Plus plate reader. Human BJ fibroblast and murine MLE-12 cell assays for inhibitor screen For inhibitor screen, human BJ fibroblast cells (ATCC CRL2522) (American Type Culture Collection, VA) were used. For mouse cell based assay MLE-12 mouse epithelial cells (ATCC CRL2110) were used. Both cell lines were maintained in ATCC recommended media. Cells were seeded at 5 × 103 cells/well into 96-well flat-bottom microtiter plates in which peptides that had been pre-diluted with cytokines (1 ng/mL for human IL-17A or 15 ng/mL for mouse IL-17A) in culture medium. Cells were incubated at 37 °C for 16–24 hrs, and supernatants were collected and analyzed by ELISA for either human CXCL1/GRO-α (R\u0026D Systems DY275) or mouse CXCL1/KC (R\u0026D Systems DY453). Primary human keratinocytes cell assay Primary human keratinocytes were cultured in Epilife medium with EDGS (Life Technology, Cascade Biologics) following product instructions. 5 days after establishing the culture from frozen vials, cells were plated at 10,000/well (80 μl) in culture media in 384 well plate. 4 hours after plating the cells, 10 μl of 10X peptide stocks were added. Final DMSO concentration was 1%. Immediately after peptide addition, 10 μl of a mixture of recombinant human IL-17A (endotoxin Level \u003c0.10 EU per 1 μg of the protein, E.coli expression, \u003e97% pure judged by SDS/page, R\u0026D System, Minneapolis, MN) 100 ng /ml (final) and TNF-α (Sigma-Aldrich, St Louis, MO) 10 ng/ ml (final), or TNF-α 10 ng/ml only were added to the cells. Cell assay plates were incubated for 48 hours at 37 °C in a tissue culture incubator before harvesting culture supernatants for analysis of IL-6, IL-8 and CCL-20 production using kits K211AKB-2, K211ANB-2, and K211BEB-2, respectively (Meso Scale Discovery, Rockville, MD). Protein production Details of non-commercial protein constructs used in this study and their purifications are in the Supplementary Material. SPR binding assays The IL-17 SPR binding assay was run on a Biacore 3000 SPR instrument (GE Healthcare). Biotinylated human IL-17A, or IL-17A/F heteromer, or IL-17F, or mouse IL-17A (Cell Signaling) was captured on a Biacore Streptavidin chip to achieve protein density of about 2500 to 3500 RUs on the surface. The SPR running buffer was 10 mM HEPES, pH 7.4, 150 mM NaCl, 0.01% P20 with 3% DMSO. Peptide samples were injected at a flow rate of 50 μl/min for 180 seconds association and at least 600 seconds dissociation using a 2–3 fold dilution series. Multiple blank injections were run before and after each peptide series for references. The data were processed and analyzed with Scrubber 2.0 (BioLogical Software) and Biaeval software (GE Healthcare) to calculate binding constants and on and off rates. The IL-17A SPR competition assay was run on a BioRAD ProteON instrument (BioRAD Laboratories). IL-17RA-Fc fusion protein (R\u0026D Systems) was captured on a GLM chip using the standard amine coupling reaction. The chip surface activated using a mixture of sulfo-NHS and EDC, was exposed to the receptor dissolved in acetate buffer pH 5.0 at 0.01 mg/ml concentration. The surface was deactivated using 1 M ethanolamine HCl. An adjacent flow cell on the chip was treated identically without the protein and was used as a reference surface for subsequent data analysis. Three-fold dilution series of peptides mixed with 5 nM IL-17A were injected at a flow rate of 50 μl/min for 120 seconds, followed by 300 seconds of dissociation. The receptor on the surface was regenerated using a 30 second injection of 3 M MgCl2. Observed signal between 180 and 200 seconds was averaged for each sample. Average response observed for multiple 5 nM IL-17A samples without any peptide was used as 100% signal for calculating % inhibition for each concentration of compound. The data were fit with Microcal Origin software (OriginLab, MA) to calculate IC50 for peptides. FRET Assay The FRET signal of a Eu3+-labeled IL-17A donor and an Alexa Fluor 647 labeled IL-17RA acceptor was measured to monitor the interaction of IL-17A and IL-17RA. Maximal FRET was observed when IL-17A was bound to IL-17RA and diminished FRET was observed when IL-17A was separated from IL-17RA. The excitation of the donor at a wavelength of 320 nm triggers fluorescence at 615 nM and this in turn serves to excite the acceptor, which then fluoresces at a wavelength of 665 nm. The fluorescence at both 615 nm and 665 nm were measured and the ratio of 665 nm/615 nm was used to monitor the IL-17A/IL-17RA binding. Final assay concentrations were 1 nM biotinylated IL-17A labeled with 0.67 nM Europium-Streptavidin (Invitrogen), 6 nM IL-17RA-Fc fusion protein (R\u0026D Systems) labeled with 1 nM Alexa Fluor 647 antibody (BioLegand) in a buffer containing 10 mM HEPES pH 7.4, 150 mM NaCl, 0.02% BSA, and 0.01% Tween 20. Peptides were tested using a half log dilution series of 11 concentrations. The IL-17A was incubated with the Europium-Streptavidin to 1.5X final assay concentration for one hour at room temperature. Peptides were prepared at 50X final concentration in 100% DMSO and 300 nl were added to a 384-well white assay plate (Greiner). 10 μl of the Eu3+-labeled IL-17A was added to the peptides and incubated at room temperature for one hour. During this pre-incubation of peptide and IL-17A, IL-17RA was incubated with Alexa Fluor 647 antibody to 3X final assay concentration at room temperature for one hour and then 5 μl of the 3X Alexa Fluor 647 labeled IL-17RA was added to the assay for a total volume of 15.3 μl and a final DMSO concentration of 2%. The plates were covered and incubated at room temperature for 3 hours. The FRET signal of the IL-17A/IL-17RA interaction was measured using an EnVision Multilabel plate reader (PerkinElmer). The peptide data was converted into % inhibition, using 0% (no HAP) and 100% inhibition (100 nM HAP) as controls. A four parameter logistic nonlinear regression model using the percent inhibition at each concentration was used to calculate an IC50 for each peptide. Protein crystallization and data collection To crystallize Fab/IL-17A/HAP complex, 30 mM HAP in DMSO stock was added to the Fab/IL-17A complex to a final concentration of 1 mM. The complex was then screened for crystals using commercial screen kits using a sitting drop vapor diffusion format. Crystals of Fab/truncated IL-17A/HAP complex were obtained under condition of 0.02 mM CdCl2, 0.02 M MgCl2, 0.02 M NiCl2, 0.1 M NaOAc pH = 4.2–4.9, and 24–28% PEG MME 2000. Crystals of Fab/full length IL-17A covalent dimer/HAP complex were obtained under conditions of 0.1 M NaOAc, pH = 4.5 and 30% PEG MME 5000. Crystals were soaked briefly in cryo solutions of the mother liquor supplemented with 25% glycerol before flash cooled in liquid nitrogen. Crystal data sets were collected at APS IMCA 17ID beamline (Chicago, IL), processed with autoPROC. Data collection statistics are listed in Supplementary Table 2. Structure determination and refinement Fab/IL-17A/HAP complex structures were solved with molecular replacement method using the published FAN/IL-17A crystal structure (pdb code 2VXS), using program Phaser. Structure refinements was carried out using program Buster and manual model building using program COOT. Final refinement statistics are listed in Supplementary Table 2. Additional Information How to cite this article: Liu, S. et al. Inhibiting complex IL-17A and IL-17RA interactions with a linear peptide. Sci. Rep. 6, 26071; doi: 10.1038/srep26071 (2016). Supplementary Material Structure and signalling in the IL-17 receptor family Innate IL-17-producing cells: the sentinels of the immune system Cutting Edge: Interleukin 17 Signals through a Heteromeric Receptor Complex Interleukin-17 and its target genes: mechanisms of interleukin-17 function in disease IL-17 induces production of IL-6 and IL-8 in rheumatoid arthritis synovial fibroblasts via NF-kappaB- and PI3-kinase/Akt-dependent pathways Induction, function and regulation of IL-17-producing T cells IL-17 as a future therapeutic target for rheumatoid arthritis Autoantibodies against IL-17A, IL-17F, and IL-22 in patients with chronic mucocutaneous candidiasis and autoimmune polyendocrine syndrome type I Chronic Mucocutaneous Candidiasis in Humans with Inborn Errors of Interleukin-17 Immunity Comparison of ixekizumab with etanercept or placebo in moderate-to-severe psoriasis (UNCOVER-2 and UNCOVER-3): results from two phase 3 randomised trials Secukinumab in Plaque Psoriasis — Results of Two Phase 3 Trials Secukinumab, a human anti-interleukin-17A monoclonal antibody, in patients with psoriatic arthritis (FUTURE 2): a randomised, double-blind, placebo-controlled, phase 3 trial Safety and efficacy of brodalumab for psoriasis after 120 weeks of treatment U.S. Food and Drug Administration. FDA approves new psoriasis drug Cosentyx. U.S. FDA press release (2015). IL-17 in synovial fluids from patients with rheumatoid arthritis is a potent stimulator of osteoclastogenesis Human interleukin-17: A T cell-derived proinflammatory cytokine produced by the rheumatoid synovium High levels of IL-17 in rheumatoid arthritis patients: IL-15 triggers in vitro IL-17 production via cyclosporin A-sensitive mechanism TH-17 cells in rheumatoid arthritis IL-1-independent role of IL-17 in synovial inflammation and joint destruction during collagen-induced arthritis Treatment with a neutralizing anti-murine interleukin-17 antibody after the onset of collagen-induced arthritis reduces joint inflammation, cartilage destruction, and bone erosion The human IL-17F/IL-17A heterodimeric cytokine signals through the IL-17RA/IL-17RC receptor complex IL‐17s adopt a cystine knot fold: structure and activity of a novel cytokine, IL‐17F, and implications for receptor binding Structural basis of receptor sharing by interleukin 17 cytokines Crystal structures of interleukin 17A and its complex with IL-17 receptor A Structure of IL-17A in Complex with a Potent, Fully Human Neutralizing Antibody Identification of Functional Roles for Both IL-17RB and IL-17RA in Mediating IL-25-Induced Activities Pyroglutamate and O-Linked Glycan Determine Functional Production of Anti-IL17A and Anti-IL22 Peptide-Antibody Bispecific Genetic Fusions Pharmacokinetics of anti-IL17A and anti-IL22 peptide–antibody bispecific genetic fusions in mice The maltose-binding protein as a scaffold for monovalent display of peptides derived from phage libraries A novel heterodimeric cytokine consisting of IL-17 and IL-17F regulates inflammatory responses Th17 cytokines and their emerging roles in inflammation and autoimmunity Integrative responses to IL-17 and TNF-alpha in human keratinocytes account for key inflammatory pathogenic circuits in psoriasis Novel microperfusion method confirms that psoriasis lesional skin contains higher protein levels of IL-17A and β-defensin-2 compared to nonlesional skin Antibody fragments as tools in crystallography   Small-Molecule Inhibitors of Protein-Protein Interactions: Progressing toward the Reality  Inhibition of [alpha]-helix-mediated protein-protein interactions using designed molecules Evaluation of Diverse α/β-Backbone Patterns for Functional α-Helix Mimicry: Analogues of the Bim BH3 Domain Identification and Characterization of Synthetic Small Molecule Macrocycle Antagonists of Human IL17A Data processing and analysis with the autoPROC toolbox Phaser crystallographic software Refinement of severely incomplete structures with maximum likelihood in BUSTER-TNT Features and development of Coot Presenting your structures: the CCP4mg molecular-graphics software Author Contributions P.J., L.D.W., J.J. and G.C. designed and performed phage display and mutagenesis optimization experiments. J.D. and B.O. designed and synthesized peptides. S.S., X.S. and M.G. created protein constructs and purified in house protein reagents. S.L. designed and performed crystallographic studies and analyzed the structures, made Figures 2 and 3, Supplementary Table S3 and Supplementary Figure S7. M.E.B., P.V.S. and W.L. performed FRET, SPR and cell assay, respectively, and made Table 3 and Figure 1. W.L. made Supplementary Figures S2 and S5. J.D., B.D.O., D.T., A.B. and C.W.B. contributed to peptide SAR. J.D., L.D.W. and P.J. prepared Tables 1 and 2, Supplementary Schemes S1 and S2, Supplementary Figures S1, S3 and S4 and Supplementary Tables S1 and S2. P.V.S. and B.A.C. performed Fab/HAP co-binding SPR and made Supplementary Figure S6. X.F. performed mass spectral analysis of the stoichiometry of HAP binding to IL-17A and made Supplementary Figure S8. L.D.W., G.C., D.T., A.B., C.W.B., G.W., R.W.L., A.T., J.M.W. and X.Q. supervised the study. S.L., J.D., J.M.W. and L.D.W. wrote the main text. All authors reviewed the manuscript. Binding of HAP to IL-17A and inhibition of IL-17A/IL-17RA are measured by SPR, FRET and cell-based assays. (A) Typical SPR sensorgrams (black) of HAP at indicated concentrations binding to biotinylated human IL-17A immobilized on a streptavidin chip surface, fitted with single site binding model curves (red). Kinetic parameters (ka, kd) were obtained by a global fit using three concentrations in triplicate. KD determined by the standard equation, KD = kd/ka. (B) HAP inhibits SPR signaling of IL-17A binding to immobilized IL-17RA. Data are mean and error bars of +/− standard deviation of three measurements. (C) Inhibition of IL-17A and IL-17RA binding by HAP measured by FRET assay. Data are mean and error bars of +/− standard deviation from 299 experiments, each performed in duplicate. (D) Example of HAP selective inhibition of the production of IL-8 (triangles), IL-6 (squares) and CCL-20 (circles) by primary human keratinocyte cells synergistically stimulated by 100 ng/ml IL-17A and 10 ng/ml TNF-α. HAP does not inhibit the baseline production of IL-6, IL-8 and CCL-20 stimulated by 10 ng/ml TNF-α alone (gray lines and symbols). Data are mean and error bars of +/− standard deviation of duplicated experiments. Overall structure of the Fab/IL-17A/HAP complex in ribbon presentation. For clarity, different molecules are colored differently. Two HAP molecules are colored blue and red, and IL-17A monomers are colored ice blue and pink, respectively. Picture prepared using program CCP4MG. (A) Overview of the distinct binding sites of Fab and HAP to IL-17A. (B) Close-in view of the IL-17A/HAP structure. IL-17A β-strands are labelled. Each of the two bound HAP interacts with both monomers of the IL-17A dimer. (C) As a comparison, the IL-17A/IL-17RA complex was shown with IL-17A in the same orientation. Three distinct areas IL-17A/IL-17RA interface are labeled. Mechanism of the inhibition of the IL-17A/IL-17RA interaction by HAP. (A) HAP binds at region I of IL-17A. IL-17A dimer is in surface presentation (β-strands 0 shown as ribbons for clarity). Polar interactions are shown in dashes. HAP residues as well as key IL-17A residues are labeled. For clarity, a few HAP residues are also shown in stick model with carbon atoms colored green, oxygen in red and nitrogen in blue. (B) I-17RA (ribbon in gold) peptide Leu27-Ile32 binds to the same area as the HAP α-helix. Trp31 of IL-17RA binds to the same pocket in IL-17A as Trp12 of HAP. (C) As illustrated by overlay a single HAP molecule and β-strands 0 (grey) of the IL-17A/HAP complex in the apo IL-17A structure, conformational changes in region I of IL-17A are needed for binding of both the β-stand and α-helix of the HAP. Notice that the Trp binding pocket for W12 of HAP or W31 of IL-17RA is missing in the apo structure. ELISA competition activity of peptide analogues of 1. Compound #\tCompetition IC50(nM)\t1\t2\t3\t4\t5\t6\t7\t8\t9\t10\t11\t12\t13\t14\t15\t \t1\t80\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tK\tA\t \t2\t80\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t3\t66\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tK\t \t4\t172\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tA\tA\t \t5\t402\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tA\tR\tA\t \t6\t7451\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tA\tI\tR\tA\t \t7\t474\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tA\tW\tI\tR\tA\t \t8\t524\tI\tV\tV\tT\tM\tP\tA\tD\tL\tA\tD\tW\tI\tR\tA\t \t9\t640\tI\tV\tV\tT\tM\tP\tA\tD\tA\tW\tD\tW\tI\tR\tA\t \t10\t4016\tI\tV\tV\tT\tM\tP\tA\tA\tL\tW\tD\tW\tI\tR\tA\t \t11\t84\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t12\t2651\tI\tV\tV\tT\tM\tA\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t13\t11\tI\tV\tV\tT\tA\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t14\t195\tI\tV\tV\tA\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t15\t5717\tI\tV\tA\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t16\t149\tI\tA\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t17\t167\tA\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t18\t45\tI\tH\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t19\t71\tI\tQ\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t20\t117\tI\tR\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t21\t31\tI\tT\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t22\t522\tI\tW\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t23\t64\tI\tY\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t24\t3\tI\tV\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t25\t33\tI\tV\tV\tT\tL\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t26\t6\tI\tV\tV\tT\tV\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t27\t31\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tM\tA\t \t28\t26\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tN\tA\t \t29\t33\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tQ\tA\t \t All peptides are acetylated at their N-termini. Cell-based activity of amino acid deletions of 30. Compound #\tCell IC50(nM)\t1\t2\t3\t4\t5\t6\t7\t8\t9\t10\t11\t12\t13\t14\t15\t \t1\t370\tI\tV\tV\tT\tM\tP\tA\tD\tL\tW\tD\tW\tI\tK\tA\t \t24\t18\tI\tV\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t26\t137\tI\tV\tV\tT\tL\tP\tA\tD\tL\tW\tD\tW\tI\tR\tA\t \t30\t17\tI\tH\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tN\tK\t \t32\t\u003e1000\t \tH\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tN\tK\t \t33\t\u003e1000\t \t \tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tN\tK\t \t34\t\u003e1000\t \t \t \tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tN\tK\t \t35\t12\tI\tH\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\tN\t \t \t36\t183\tI\tH\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\tI\t \t \t \t37\t\u003e1000\tI\tH\tV\tT\tI\tP\tA\tD\tL\tW\tD\tW\t \t \t \t \t Binding affinity/IC50 of HAP in orthogonal in vitro assays/primary keratinocyte cell assay. Assay\tKD/IC50 (nM)\t \tSPR binding (N = 3)*: hIL-17A, hIL-17A/F, mIL-17A\t2.6(±0.5), 4.6(±0.7), 30.1(±0.1)\t \tSPR competition\t0.9(±0.3)\t \tFRET IL-17RA competition (N = 598)\t0.8 ± 0.2\t \tHuman primary keratinocytes IC-8 (N = 9), IC-6 (N = 4), CCL-20 (N = 4)\t151(±49), 136(±37), 253(±64)\t \t *KD = kd/ka. ka (M−1s−1)and kd (s−1): 2(±1) × 105, 6(±5) × 10−4 (hIL-17A); 4.6(±0.7) × 105, 2.1(±0.2) × 10−3 (hIL-17A/F); 2.7(±0.2) × 105, 8.2(±0.8) × 10−3 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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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diff --git a/annotated_BioC_JSON/PMC4872110_ann.json b/annotated_BioC_JSON/PMC4872110_ann.json
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index 0000000000000000000000000000000000000000..79a899f3a4347860fbedd32ff0d04246cca7de7f
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+++ b/annotated_BioC_JSON/PMC4872110_ann.json
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+[{"sourceid":"4872110","sourcedb":"","project":"","target":"","text":"Ribosome biogenesis factor Tsr3 is the aminocarboxypropyl transferase responsible for 18S rRNA hypermodification in yeast and humans The chemically most complex modification in eukaryotic rRNA is the conserved hypermodified nucleotide N1-methyl-N3-aminocarboxypropyl-pseudouridine (m1acp3Ψ) located next to the P-site tRNA on the small subunit 18S rRNA. While S-adenosylmethionine was identified as the source of the aminocarboxypropyl (acp) group more than 40 years ago the enzyme catalyzing the acp transfer remained elusive. Here we identify the cytoplasmic ribosome biogenesis protein Tsr3 as the responsible enzyme in yeast and human cells. In functionally impaired Tsr3-mutants, a reduced level of acp modification directly correlates with increased 20S pre-rRNA accumulation. The crystal structure of archaeal Tsr3 homologs revealed the same fold as in SPOUT-class RNA-methyltransferases but a distinct SAM binding mode. This unique SAM binding mode explains why Tsr3 transfers the acp and not the methyl group of SAM to its substrate. Structurally, Tsr3 therefore represents a novel class of acp transferase enzymes. INTRODUCTION Eukaryotic ribosome biogenesis is highly complex and requires a large number of non-ribosomal proteins and small non-coding RNAs in addition to ribosomal RNAs (rRNAs) and proteins. An increasing number of diseases—so called ribosomopathies—are associated with disturbed ribosome biogenesis. During eukaryotic ribosome biogenesis several dozens of rRNA nucleotides become chemically modified. The most abundant rRNA modifications are methylations at the 2′-OH ribose moieties and isomerizations of uridine residues to pseudouridine, catalyzed by small nucleolar ribonucleoprotein particles (snoRNPs). In addition, 18S and 25S (yeast)/ 28S (humans) rRNAs contain several base modifications catalyzed by site-specific and snoRNA-independent enzymes. In Saccharomyces cerevisiae 18S rRNA contains four base methylations, two acetylations and a single 3-amino-3-carboxypropyl (acp) modification, whereas six base methylations are present in the 25S rRNA. While in humans the 18S rRNA base modifications are highly conserved, only three of the yeast base modifications catalyzed by ScRrp8/HsNML, ScRcm1/HsNSUN5 and ScNop2/HsNSUN1 are preserved in the corresponding human 28S rRNA. Ribosomal RNA modifications have been suggested to optimize ribosome function, although in most cases this remains to be clearly established. They might contribute to increased RNA stability by providing additional hydrogen bonds (pseudouridines), improved base stacking (pseudouridines and base methylations) or an increased resistance against hydrolysis (ribose methylations). Most modified rRNA nucleotides cluster in the vicinity of the decoding or the peptidyl transferase center, suggesting an influence on ribosome functionality and stability. Defects of rRNA modification enzymes often lead to disturbed ribosome biogenesis or functionally impaired ribosomes, although the lack of individual rRNA modifications often has no or only a slight influence on the cell. Importantly, malfunctions of several base modifying enzymes are linked to developmental diseases, aging or tumorigenesis. The chemically most complex modification is located in the loop capping helix 31 of 18S rRNA (Supplementary Figure S1B). There a uridine (U1191 in yeast) is modified to 1-methyl-3-(3-amino-3-carboxypropyl)-pseudouridine (m1acp3Ψ, Figure 1A). This base modification was first described in 1968 for hamster cells and is conserved in eukaryotes. This hypermodified nucleotide, which is located at the P-site tRNA, is synthesized in three steps beginning with the snR35 H/ACA snoRNP guided conversion of uridine into pseudouridine. In a second step, the essential SPOUT-class methyltransferase Nep1/Emg1 modifies the pseudouridine to N1-methylpseudouridine. Methylation can only occur once pseudouridylation has taken place, as the latter reaction generates the substrate for the former. The final acp modification leading to N1-methyl-N3-aminocarboxypropyl-pseudouridine occurs late during 40S biogenesis in the cytoplasm, while the two former reactions are taking place in the nucleolus and nucleus, and is independent from pseudouridylation or methylation. Both the methyl and the acp group are derived from S-adenosylmethionine (SAM), but the enzyme responsible for acp modification remained elusive for more than 40 years. Tsr3 is necessary for acp modification of 18S rRNA in yeast and human. (A) Hypermodified nucleotide m1acp3Ψ is synthesized in three steps: pseudouridylation catalyzed by snoRNP35, N1-methylation catalyzed by methyltransferase Nep1 and N3-acp modification catalyzed by Tsr3. The asterisk indicates the C1-atom labeled in the 14C-incorporation assay. (B) RP-HPLC elution profile of yeast 18S rRNA nucleosides. Hypermodified m1acp3Ψ elutes at 7.4 min (wild type, left profile) and is missing in Δtsr3 (middle profile) and Δnep1 Δnop6 mutants (right profile). (C) 14C-acp labeling of 18S rRNAs. Wild type (WT) and plasmid encoded 18S rRNA (U1191U) show the 14C-acp signal, whereas the 14C-acp signal is missing in the U1191A mutant plasmid encoded 18S rRNA (U1191A) and Δtsr3 mutants (Δtsr3). Upper lanes show the ethidium bromide staining of the 18S rRNAs for quantification. All samples were loaded on the gel with two different amounts of 5 and 10 μl. (D) Primer extension analysis of acp modification in yeast 18S rRNA (right gel) including a sequencing ladder (left gel). The primer extension stop at nucleotide 1191 is missing exclusively in Δtsr3 mutants and Δtsr3 Δsnr35 recombinants. (E) Primer extension analysis of human 18S rRNA after siRNA knockdown of HsNEP1/EMG1 (541, 542 and 543) and HsTSR3 (544 and 545) (right gel), including a sequencing ladder (left gel). The primer extension arrest is reduced in HTC116 cells transfected with siRNAs 544 and 545. The efficiency of siRNA mediated HsTSR3 repression correlates with the primer extension signals (see Supplementary Figure S2A). As a loading control, a structural stop is shown (asterisks). Only a few acp transferring enzymes have been characterized until now. During the biosynthesis of wybutosine, a tricyclic nucleoside present in eukaryotic and archaeal phenylalanine tRNA, Tyw2 (Trm12 in yeast) transfers an acp group from SAM to an acidic carbon atom. Archaeal Tyw2 has a structure very similar to Rossmann-fold (class I) RNA-methyltransferases, but its distinctive SAM-binding mode enables the transfer of the acp group instead of the methyl group of the cofactor. Another acp modification has been described in the diphtamide biosynthesis pathway, where an acp group is transferred from SAM to the carbon atom of a histidine residue of eukaryotic translation elongation factor 2 by use of a radical mechanism. In a recent bioinformatic study, the uncharacterized yeast gene YOR006c was predicted to be involved in ribosome biogenesis. It is highly conserved among eukaryotes and archaea (Supplementary Figure S1A) and its deletion leads to an accumulation of the 20S pre-rRNA precursor of 18S rRNA, suggesting an influence on D-site cleavage during the maturation of the small ribosomal subunit. On this basis, YOR006C was renamed ‘Twenty S rRNA accumulation 3′ (TSR3). However, its function remained unclear although recently a putative nuclease function during 18S rRNA maturation was predicted. Here, we identify Tsr3 as the long-sought acp transferase that catalyzes the last step in the biosynthesis of the hypermodified nucleotide m1acp3Ψ in yeast and human cells. Furthermore using catalytically defective mutants of yeast Tsr3 we demonstrated that the acp modification is required for 18S rRNA maturation. Surprisingly, the crystal structures of archaeal homologs revealed that Tsr3 is structurally similar to the SPOUT-class RNA methyltransferases. In contrast, the only other structurally characterized acp transferase enzyme Tyw2 belongs to the Rossmann-fold class of methyltransferase proteins. Interestingly, the two structurally very different enzymes use similar strategies in binding the SAM-cofactor in order to ensure that in contrast to methyltransferases the acp and not the methyl group of SAM is transferred to the substrate. MATERIALS AND METHODS Genetic constructions, growth conditions and yeast media Detailed descriptions are available in Supplementary Data. Cell culture HCT116(+/+) cells (CCL-247; ATCC) were grown at 37°C in a humidified incubator under 5% CO2 in the McCoy's 5a modified (Sigma-Aldrich)/10% FBS media. All media were supplemented with 50 U/ml penicillin and 50 μg/ml streptomycin (Life Technologies). DsiRNA inactivation and RT-qPCR Reverse transfection of HCT116 cells, DsiRNA inactivation and RT-qPCR using total human RNA are described in Supplementary Data. Sucrose gradient analysis Detailed descriptions for analytical or preparative separations of ribosomal subunits or polysome gradients are provided in Supplementary Data. HPLC analysis of 18S rRNA nucleosides 40S subunits from 200 ml yeast culture were isolated by sucrose gradient centrifugation in a SW28 rotor as described above, and precipitated with 2.5 vol of 100% ethanol (−20°C over night). Precipitated 40S subunits were dissolved in water and the 18S rRNA was purified via spin columns (Ambion PureLink RNA Mini Kit). RNA fragments were hydrolysed and dephosphorylated as described by Gehrke and Kuo. HPLC analysis of rRNA nucleoside composition was performed using a Supelcosil LC-18S column (Sigma; 250 × 4.6 mm, 5 μm) with a pre-column (4.6 × 20 mm) as previously described. 14C labeling of 18S rRNA nucleotide Ψ(U)1191 To enhance 14C-labeling, mutants of interest were recombined with a Δmet13 deletion. Resulting strains were cultivated with l-[1-14C]-methionine (Hartmann Analytic, 0.1 mCi/ml, 54 mCi/mmol) as described before. From isotope labeled cells total RNA was isolated with the PureLink RNA Mini Kit (Ambion) after enzymatic cell lysis with zymolyase. Ribosomal RNAs were separated on a 4% denaturing polyacrylamide gel. After ethidium bromide straining gels were dried and analyzed by autoradiography for 3–5 days using a storage phosphor screen. Signals were visualized with the Typhoon 9100 (GE Healthcare). Northern blot analysis 5 μg of total yeast RNAs extracted with phenol/chloroform were separated on 1.2% agarose gels in BPTE buffer for 16 h at 60V and afterwards transferred to a Biodyne B membrane by vacuum blotting. Oligonucleotides D/A2 or +1-A0 were radiolabeled using γ-[32P]-ATP and T4-polynucleotide kinase and hybridized to the membrane at 37°C. Signals were visualized by phosphoimaging with the Typhoon 9100 (GE Healthcare). RNA extraction from human cells, gel-electrophoresis and northern blotting were performed as described before. Primer extension 20 pmol of oligonucleotide PE-1191 complementary to yeast 18S rRNA nucleotides 1247–1228 were labeled with 50 μCi γ-[32P]-ATP using T4-polynucleotide kinase, purified via Sephadex G-25 and annealed to 500 ng of 18S rRNA. Primer annealing and reverse transcription were carried out as described by Sharma et al.. After precipitation with ethanol and 3 M NaAc pH 5.2 pellets were washed with 70% ethanol, dried and dissolved in 12 μl formamide loading dye. 2 μl of primer extension samples were separated on sequencing or mini gels which were dried after running and exposed on a storage phosphor screen. Signals were visualized with the Typhoon 9100 (GE Healthcare). Primer extension on human RNA was performed using 5 μg of total RNA with AMV Reverse Transcriptase (Promega) and oligonucleotide PE_1248. Following alkaline hydrolysis, cDNAs were precipitated with ethanol, resuspended in acrylamide loading buffer and separated on a 6% (v/v) denaturing acrylamide gel in 0.5× TBE at 80 W for 1.5 h. After migration, the gels were dried and exposed to Fuji Imaging plates (Fujifilm). The signal was acquired with a Phosphor imager (FLA-7000, Fujifilm). Protein detection and localization A description of the western blot detection of HA-fused Tsr3 in yeast crude extracts or sucrose gradients fractions is provided in Supplementary Data. For cellular localization Tsr3 was expressed as N-terminal fusion with yEGFP in a yeast strain encoding for ScNop56-mRFP. Protein localization in exponentially growing cells was visualized using a Leica TCS SP5. In vitro SAM binding Purified SsTsr3 protein in 25 mM Tris–HCl pH 7.8 250 mM NaCl was mixed with S-[methyl-14C]-adenosyl-l-methionine (PerkinElmer; 20 μCi/ml, 58 mCi/mmol) and 0–10 mM non-labeled SAM in a binding buffer (50 mM Tris–HCl pH 7.8, 250 mM NaCl) in a total volume of 50 μl and incubated at 30°C for 10 min. Samples were passed over HAWP02500 membrane filters (Millipore) and unbound 14C-SAM was removed by washing three times with 5 ml buffer using a vacuum filtering equipment. Filter bound 14C-SAM was measured by liquid scintillation spectrometry in a Wallac 1401 scintillation counter. Protein expression and purification Genes coding for archaeal Tsr3 homologs without any tags were obtained commercially (Genscript) in pET11a vectors and overexpressed in Escherichia coli BL21(DE3). Proteins were purified by a combination of heat shock and appropriate column chromatography steps as described in detail in the Supplementary Data. Crystallization, X-ray data collection, structure calculation and refinement Initial hits for VdTsr3 and SsTsr3 were obtained using the Morpheus Screen (Molecular dimensions) and further refined as described in the Supplementary Data. Diffraction data were collected at the Swiss Light Source (Paul Scherer Institut). The structure of VdTsr3 was determined at 1.6 Å by SAD using a selenomethionine derivative. The structure of SsTsr3 was determined at 2.25 Å by molecular replacement using VdTsr3 as the search model. A detailed description of the data collection, processing, structure calculation and refinement procedures can be found in the Supplementary Data and in Supplementary Table S1. Structures were deposited in the Protein Data Bank as entries 5APG (VdTsr3) and 5AP8 (SsTsr3). Analytical gel filtration For analytical gel filtration experiments a Sephadex S75 10/300 GL column (GE Healthcare) was used. 100 μl protein samples (25 mM Tris–HCl pH 7.8, 250 mM NaCl, 2 mM β-mercaptoethanol) with a protein concentration of 150 μM were used. The flow rate was 0.5 ml/min. The column was calibrated using the marker proteins of the LMW gel filtration calibration kit (GE Healthcare). Protein elution was followed by recording the adsorption at a wavelength of λ = 280 nm. Fluorescence quenching and fluorescence anisotropy measurements Fluorescence quenching and fluorescence anisotropy measurements were carried out in triplicates at 25°C on a Fluorolog 3 spectrometer (Horiba Jobin Yvon) equipped with polarizers. For fluorescence quenching with SAM, SAH and 5′-methylthioadenosine experiments the tryptophan fluorescence of SsTsr3 (200 nM in 25 mM Tris–HCl pH 7.8, 250 mM NaCl, 2 mM β-mercaptoethanol) was excited at 295 nm and emission spectra were recorded from 250 to 450 nm for each titration step. The fluorescence intensity at 351 nm for each titration step was normalized with regard to the fluorescence of the free protein and was used for deriving binding curves. KD's were derived by nonlinear regression with Origin 8.0 (Origin Labs) using Equation (1):  (F is the normalized fluorescence intensity, a is the change in fluorescence intensity, c is the ligand concentration and KD is the dissociation constant). 5′-Fluoresceine labeled RNAs for fluorescence anisotropy measurements were obtained commercially (Dharmacon), deprotected according to the manufacturer's protocol and the RNA concentration adjusted to 50 nM in 25 mM Tris–HCl pH 7.8, 250 mM NaCl. Fluoresceine fluorescence was excited at 492 nm and emission was recorded at 516 nm. The data were fitted to Equation (1) (F is the normalized fluorescence anisotropy, a is the change in fluorescence anisotropy). RESULTS Tsr3 is the enzyme responsible for 18S rRNA acp modification in yeast and humans The S. cerevisiae 18S rRNA acp transferase was identified in a systematic genetic screen where numerous deletion mutants from the EUROSCARF strain collection (www.euroscarf.de) were analyzed by HPLC for alterations in 18S rRNA base modifications. For the Δtsr3 deletion strain the HPLC elution profile of 18S rRNA nucleosides (Figure 1B) was very similar to that of the pseudouridine-N1 methyltransferase mutant Δnep1, where a shoulder at ∼ 7.4 min elution time was missing in the elution profile. As previously reported this shoulder was identified by ESI-MS as corresponding to m1acp3Ψ. In order to directly analyze the presence of the acp modification of nucleotide 1191 we used an in vivo14C incorporation assay with 1-14C-methionine. Whereas the acp labeling of 18S rRNA was clearly present in the wild type strain no radioactive labeling could be observed in a Δtsr3 strain (Figure 1C). No radioactive labeling was detected in the 18S U1191A mutant which served as a control for the specificity of the 14C-aminocarboxypropyl incorporation. As previously shown, only the acp but none of the other modifications at U1191 of yeast 18S rRNA blocks reverse transcriptase activity. Therefore the presence of the acp modification can be directly assessed by primer extension. Indeed, in wild-type yeast a strong primer extension stop signal occurred at position 1192. In contrast, in a Δtsr3 mutant no primer extension stop signal was present at this position. As expected, in a Δsnr35 deletion preventing pseudouridylation and N1-methylation (resulting in acp3U) as well as in a Δnep1 deletion strain where pseudouridine is not methylated (resulting in acp3Ψ) a primer extension stop signal of similar intensity as in the wild type was observed. In a Δtsr3 Δsnr35 double deletion strain the 18S rRNA contains an unmodified U and the primer extension stop signal was missing (Figure 1D). The Tsr3 protein is highly conserved in yeast and humans (50% identity). Human 18S rRNA has also been shown to contain m1acp3Ψ in the 18S rRNA at position 1248. After siRNA-mediated depletion of Tsr3 in human colon carcinoma HCT116(+/+) cells the acp primer extension arrest was reduced in comparison to cells transfected with a non-targeting scramble siRNA control (Figure 1E, compare lanes 544 and scramble). The efficiency of siRNA-mediated depletion was established by RT-qPCR and found to be very high with siRNA 544 (Supplementary Figure S2A, remaining TSR3 mRNA level of 2%). By comparison, treating cells with siRNA 545, which only reduced the TSR3 mRNA to 20%, did not markedly reduced the acp signal. This suggests that low residual levels of HsTsr3 are sufficient to modify the RNA. As a control for loading, a structural stop is shown (asterisk, Figure 1E). Thus, HsTsr3 is also responsible for the acp modification of 18S rRNA nucleotide Ψ1248 in helix 31. Similar to yeast, siRNA-mediated depletion of the Ψ1248 N1-methyltransferase Nep1/Emg1 had no influence on the primer extension arrest (Figure 1E). Phenotypic characterization of Δtsr3 mutants Although the acp modification of 18S rRNA is highly conserved in eukaryotes, yeast Δtsr3 mutants showed only a minor growth defect. However, the Δtsr3 deletion was synthetic sick with a Δsnr35 deletion preventing pseudouridylation and Nep1-catalyzed methylation of nucleotide 1191 (Figure 2A). Interestingly, no increased growth defect could be observed for Δtsr3 Δnep1 recombinants containing the nep1 suppressor mutation Δnop6 as well as for Δtsr3 Δsnr35 Δnep1 recombinants with unmodified U1191 (Supplementary Figure S2D and E). Phenotypic characterization of yeast TSR3 deletion (Δtrs3) and human TSR3 depletion (siRNAs 544 and 545) and cellular localization of yeast Tsr3. (A) Growth of yeast wild type, Δtsr3, Δsnr35 and Δtsr3 Δsnr35 segregants after meiosis and tetrad dissection of Δtsr3/TSR3 Δsnr35/SNR35 heterozygous diploids. The Δtsr3 deletion is synthetic sick with a Δsnr35 deletion preventing U1191 pseudouridylation. (B) In agar diffusion assays the yeast Δtsr3 deletion mutant shows a hypersensitivity against paromomycin and hygromycin B which is further increased by recombination with Δsnr35. (C) Northern blot analysis with an ITS1 hybridization probe after siRNA depletion of HsTSR3 (siRNAs 544 and 545) and a scrambled siRNA as control. The accumulation of 18SE and 47S and/or 45S pre-RNAs is enforced upon HsTSR3 depletion. Right gel: Ethidium bromide staining showing 18S and 28S rRNAs. (D) Cytoplasmic localization of yeast Tsr3 shown by fluorescence microscopy of GFP-fused Tsr3. From left to right: differential interference contrast (DIC), green fluorescence of GFP-Tsr3, red fluorescence of Nop56-mRFP as nucleolar marker, and merge of GFP-Tsr3/Nop56-mRFP with DIC. (E) Elution profile (A254) after sucrose gradient separation of yeast ribosomal subunits and polysomes (upper part) and western blot analysis of 3xHA tagged Tsr3 (Tsr3-3xHA) after SDS-PAGE separation of polysome profile fractions taken every 20 s (lower part). The TSR3 gene was genetically modified at its native locus, resulting in a C-terminal fusion of Tsr3 with a 3xHA epitope expressed by the native promotor in yeast strain CEN.BM258-5B. The influence of the acp modification of nucleotide 1191 on ribosome function was analyzed by treating Δtsr3 mutants with protein synthesis inhibitors. Similar to a temperature-sensitive nep1 mutant, the Δtsr3 deletion caused hypersensitivity to paromomycin and, to a lesser extent, to hygromycin B (Figure 2B), but not to G418 or cycloheximide (data not shown). In accordance with the synthetic sick growth phenotype the paromomycin and hygromycin B hypersensitivity further increased in a Δtsr3 Δsnr35 recombination strain (Figure 2B). In a yeast Δtsr3 strain as well as in the Δtsr3 Δsnr35 recombinant 20S pre-rRNA accumulated significantly and the level of mature 18S rRNA was reduced (Supplementary Figures S2C and S3D), as reported previously. A minor effect on 20S rRNA accumulation was also observed for Δsnr35, but - probably due to different strain backgrounds – to a weaker extent than described earlier. In human cells, the depletion of HsTsr3 in HCT116(+/+) cells caused an accumulation of the human 20S pre-rRNA equivalent 18S-E suggesting an evolutionary conserved role of Tsr3 in the late steps of 18S rRNA processing (Figure 2C and Supplementary Figure S2B). Surprisingly, early nucleolar processing reactions were also inhibited, and this was observed in both yeast Δtsr3 cells (see accumulation of 35S in Supplementary Figure S2C) and Tsr3 depleted human cells (see 47S/45S accumulation in Figure 2C and Northern blot quantification in Supplementary Figure S2B). Consistent with its role in late 18S rRNA processing, TSR3 deletion leads to a ribosomal subunit imbalance with a reduced 40S to 60S ratio of 0.81 (σ = 0.024) which was further increased in a Δtsr3 Δsnr35 recombinant to 0.73 (σ = 0.023) (Supplementary Figure S2F). In polysome profiles, a reduced level of 80S ribosomes and a strong signal for free 60S subunits was observed in line with the 40S subunit deficiency (Supplementary Figure S2G). Cellular localization of Tsr3 in S. cerevisiae Fluorescence microscopy of GFP-tagged Tsr3 localized the fusion protein in the cytoplasm of yeast cells and no co-localization with the nucleolar marker protein Nop56 could be observed (Figure 2D). This agrees with previous biochemical data suggesting that the acp modification of 18S rRNA occurs late during 40S subunit biogenesis in the cytoplasm, and makes an additional nuclear localization as reported in a previous large-scale analysis unlikely. After polysome gradient separation C-terminally epitope-labeled Tsr3-3xHA was exclusively detectable in the low-density fraction (Figure 2E). Such distribution on a density gradient suggests that Tsr3 only interacts transiently with pre-40S subunits, which presumably explains why it was not characterized in pre-ribosome affinity purifications. Structure of Tsr3 Searches for sequence homologs of S. cerevisiae Tsr3 (ScTsr3) by us and others revealed that the genomes of many archaea contain genes encoding Tsr3-like proteins. However, these archaeal homologs are significantly smaller than ScTsr3 (∼190 aa in archaea vs. 313 aa in yeast) due to shortened N- and C-termini (Supplementary Figure S1A). To locate the domains most important for Tsr3 activity, ScTsr3 fragments of different lengths containing the highly conserved central part were expressed in a Δtsr3 mutant (Figure 3A) and analyzed by primer extension (Figure 3B) and Northern blotting (Figure 3C). N-terminal truncations of up to 45 aa and C-terminal truncations of up to 76 aa mediated acp modification as efficiently as the full-length protein and no significant increased levels of 20S pre-RNA were detected. Even a Tsr3 fragment with a 90 aa C-terminal truncation showed a residual primer extension stop, whereas N-terminal truncations exceeding 46 aa almost completely abolished the primer extension arrest (Figure 3B). Domain characterization of yeast Tsr3 and correlation of acp modification with late 18S rRNA processing steps. (A) Scheme of the TSR3 gene with truncation positions in the open reading frame. TSR3 fragments of different length were expressed under the native promotor from multicopy plasmids in a Δtsr3 deletion strain. (B) Primer extension analysis of 18S rRNA acp modification in yeast cells expressing the indicated TSR3 fragments. N-terminal deletions of 36 or 45 amino acids and C-terminal deletions of 43 or 76 residues show a primer extension stop comparable to the wild type. Tsr3 fragments 37–223 or 46–223 cause a nearly complete loss of the arrest signal. The box highlights the shortest Tsr3 fragment (aa 46–270) with wild type activity (strong primer extension block). (C) Northern blot analysis of 20S pre-rRNA accumulation. A weak 20S rRNA signal, indicating normal processing, is observed for Tsr3 fragment 46–270 (highlighted in a box) showing its functionality. Strong 20S rRNA accumulation similar to that of the Δtsr3 deletion is observed for Tsr3 fragments 37–223 or 46–223. Thus, the archaeal homologs correspond to the functional core of Tsr3. In order to define the structural basis for Tsr3 function, homologs from thermophilic archaea were screened for crystallization. We focused on archaeal species containing a putative Nep1 homolog suggesting that these species are in principle capable of synthesizing N1-methyl-N3-acp-pseudouridine. Well diffracting crystals were obtained for Tsr3 homologs from the two crenarchaeal species Vulcanisaeta distributa (VdTsr3) and Sulfolobus solfataricus (SsTsr3) which share 36% (VdTsr3) and 38% (SsTsr3) identity with the ScTsr3 core region (ScTsr3 aa 46–223). While for S. solfataricus the existence of a modified nucleotide of unknown chemical composition in the loop capping helix 31 of its 16S rRNA has been demonstrated, no information regarding rRNA modifications is yet available for V. distributa. Crystals of VdTsr3 diffracted to a resolution of 1.6 Å whereas crystals of SsTsr3 diffracted to 2.25 Å. Serendipitously, VdTsr3 was purified and crystallized in complex with endogenous (E. coli) SAM (Supplementary Figure S4) while SsTsr3 crystals contained the protein in the apo state. The structure of VdTsr3 was solved ab initio, by single-wavelength anomalous diffraction phasing (Se-SAD) with Se containing derivatives (selenomethionine and seleno-substituted SAM). The structure of SsTsr3 was solved by molecular replacement using VdTsr3 as a search model (see Supplementary Table S1 for data collection and refinement statistics). The structure of VdTsr3 can be divided into two domains (Figure 4A). The N-terminal domain (aa 1–92) has a mixed α/β-structure centered around a five-stranded all-parallel β-sheet (Figure 4B) with the strand order β5↑-β3↑-β4↑-β1↑-β2↑. The loops connecting β1 and β2, β3 and β4 and β4 and β5 include α-helices α1, α2 and α3, respectively. The loop connecting β2 and β3 contains a single turn of a 310-helix. Helices α1 and α2 are located on one side of the five-stranded β-sheet while α3 packs against the opposite β-sheet surface. The C-terminal domain (aa 93–184) has a globular all α-helical structure comprising α-helices α4 to α9. Both domains are tightly packed against each other. Remarkably, the entire C-terminal domain (92 aa) of the protein is threaded through the loop which connects β-strand β3 and α-helix α2 of the N-terminal domain. Thus, the VdTsr3 structure contains a deep trefoil knot. The structure of SsTsr3 in the apo state is very similar to that of VdTsr3 (Figure 4C) with an RMSD for equivalent Cα atoms of 1.1 Å. The only significant difference in the global structure of the two proteins is the presence of an extended α-helix α8 and the absence of α-helix α9 in SsTsr3. Tsr3 has a fold similar to SPOUT-class RNA methyltransferases. (A) Cartoon representation of the X-ray structure of VdTsr3 in two orientations. β-strands are colored in crimson whereas α-helices in the N-terminal domain are colored light blue and α-helices in the C-terminal domain are colored dark blue. The bound S-adenosylmethionine is shown in a stick representation and colored by atom type. A red arrow marks the location of the topological knot in the structure. (B) Secondary structure representation of the VdTsr3 structure. The color coding is the same as in (A). (C) Structural superposition of the X-ray structures of VdTsr3 in the SAM-bound state (red) and SsTsr3 (blue) in the apo state. The locations of the α-helix α8 which is longer in SsTsr3 and of α-helix α9 which is only present in VdTsr3 are indicated. (D) Secondary structure cartoon (left) of S. pombe Trm10 (pdb4jwf)—the SPOUT-class RNA methyltransferase structurally most similar to Tsr3 and superposition of the VdTsr3 and Trm10 X-ray structures (right). (E) Analytical gel filtration profiles for VdTsr3 (red) and SsTsr3 (blue) show that both proteins are monomeric in solution. Vertical lines indicate the elution volumes of molecular weight markers. Vd, Vulcanisaeta distributa; Ss, Sulfolobus solfataricus. Structure predictions suggested that Tsr3 might contain a so-called RLI domain which contains a ‘bacterial like’ ferredoxin fold and binds two iron-sulfur clusters through eight conserved cysteine residues. However, no structural similarity to an RLI-domain was detectable. This is in accordance with the functional analysis of alanine replacement mutations of cysteine residues in ScTsr3 (Supplementary Figure S3). The β-strand topology and the deep C-terminal trefoil knot of archaeal Tsr3 are the structural hallmarks of the SPOUT-class RNA-methyltransferase fold. The closest structural homolog identified in a DALI search is the tRNA methyltransferase Trm10 (DALI Z-score 6.8) which methylates the N1 nitrogen of G9/A9 in many archaeal and eukaryotic tRNAs by using SAM as the methyl group donor. In comparison to Tsr3 the central β-sheet element of Trm10 is extended by one additional β-strand pairing to β2. Furthermore, the trefoil knot of Trm10 is not as deep as that of Tsr3 (Figure 4D). Interestingly, Nep1—the enzyme preceding Tsr3 in the biosynthetic pathway for the synthesis of m1acp3Ψ—also belongs to the SPOUT-class of RNA methyltransferases. However, the structural similarities between Nep1 and Tsr3 (DALI Z-score 4.4) are less pronounced than between Tsr3 and Trm10. Most SPOUT-class RNA-methyltransferases are homodimers. A notable exception is Trm10. Gel filtration experiments with both VdTsr3 and SsTsr3 (Figure 4E) showed that both proteins are monomeric in solution thereby extending the structural similarities to Trm10. So far, structural information is only available for one other enzyme that transfers the acp group from SAM to an RNA nucleotide. This enzyme, Tyw2, is part of the biosynthesis pathway of wybutosine nucleotides in tRNAs. However, there are no structural similarities between Tsr3 and Tyw2, which contains an all-parallel β-sheet of a different topology and no knot structure. Instead, Tyw2 has a fold typical for the class-I-or Rossmann-fold class of methyltransferases (Supplementary Figure S5B). Cofactor binding of Tsr3 The SAM-binding site of Tsr3 is located in a deep crevice between the N- and C-terminal domains in the vicinity of the trefoil knot as typical for SPOUT-class RNA-methyltransferases (Figure 4A). The adenine base of the cofactor is recognized by hydrogen bonds between its N1 nitrogen and the backbone amide of L93 directly preceding β5 as well as between its N6-amino group and the backbone carbonyl group of Y108 located in the loop connecting β5 in the N-terminal and α4 in the C-terminal domain (Figure 5A). Furthermore, the adenine base of SAM is involved in hydrophobic packing interactions with the side chains of L45 (β3), P47 and W73 (α3) in the N-terminal domain as well as with L93, L110 (both in the loop connecting β5 and α4) and A115 (α5) in the C-terminal domain. The ribose 2′ and 3′ hydroxyl groups of SAM are hydrogen bonded to the backbone carbonyl group of I69. The acp side chain of SAM is fixed in position by hydrogen bonding of its carboxylate group to the backbone amide and the side chain hydroxyl group of T19 in α1 as well as the backbone amide group of T112 in α4 (C-terminal domain). Most importantly, the methyl group of SAM is buried in a hydrophobic pocket formed by the sidechains of W73 and A76 both located in α3 (Figure 5A and B). W73 is highly conserved in all known Tsr3 proteins, whereas A76 can be replaced by other hydrophobic amino acids. Consequently, the accessibility of this methyl group for a nucleophilic attack is strongly reduced in comparison with RNA-methyltransferases such as Trm10 (Figure 5B, C). In contrast, the acp side chain of SAM is accessible for reactions in the Tsr3-bound state (Figure 5B). SAM-binding by Tsr3. (A) Close-up view of the SAM-binding pocket of VdTsr3. Nitrogen atoms are dark blue, oxygen atoms red, sulfur atoms orange, carbon atoms of the protein light blue and carbon atoms of SAM yellow. Hydrogen bonds are indicated by dashed lines. (B) Solvent accessibility of the acp group of SAM bound to VdTsr3. The solvent accessible surface of the protein is shown in semitransparent gray whereas SAM is show in a stick representation. Atoms are colored as in (A). A red arrow indicates the reactive CH2-moiety of the acp group. (C) Solvent accessibility of the SAM methyl group for SAM bound to the RNA methyltransferase Trm10. Bound SAM was modelled based on the X-ray structure of the Trm10/SAH-complex (pdb4jwf). A red arrow indicates the SAM methyl group. (D) Binding of SAM analogs to SsTsr3. Tryptophan fluorescence quenching curves upon addition of SAM (blue), 5′-methyl-thioadenosine (red) and SAH (black). (E) Binding of 14C-labeled SAM to SsTsr3. Radioactively labeled SAM is retained on a filter in the presence of SsTsr3. Addition of unlabeled SAM competes with the binding of labeled SAM. A W66A-mutant of SsTsr3 (W73 in VdTsr3) does not bind SAM. (F) Primer extension (upper left) shows a strongly reduced acp modification of yeast 18S rRNA in Δtsr3 cells expressing Tsr3-S62D, -E111A or –W114A. This correlates with a 20S pre-rRNA accumulation comparable to the Δtsr3 deletion (right: northern blot). 3xHA tagged Tsr3 mutants are expressed comparable to the wild type as shown by western blot (lower left). Binding affinities for SAM and its analogs 5′-methylthioadenosin and SAH to SsTsr3 were measured using tryptophan fluorescence quenching. VdTsr3 could not be used in these experiments since we could not purify it in a stable SAM-free form. SsTsr3 bound SAM with a KD of 6.5 μM, which is similar to SAM-KD's reported for several SPOUT-class methyltransferases. 5′-methylthioadenosin—the reaction product after the acp-transfer—binds only ∼2.5-fold weaker (KD = 16.7 μM) compared to SAM. S-adenosylhomocysteine which lacks the methyl group of SAM binds with significantly lower affinity (KD = 55.5 μM) (Figure 5D). This suggests that the hydrophobic interaction between SAM's methyl group and the hydrophobic pocket of Tsr3 is thermodynamically important for the interaction. On the other hand, the loss of hydrogen bonds between the acp sidechain carboxylate group and the protein appears to be thermodynamically less important but these hydrogen bonds might play a crucial role for the proper orientation of the cofactor side chain in the substrate binding pocket. Accordingly, a W66A-mutation (W73 in VdTsr3) of SsTsr3 significantly diminished SAM-binding in a filter binding assay compared to the wild type (Figure 5E). Furthermore, a W to A mutation at the equivalent position W114 in ScTsr3 strongly reduced the in vivo acp transferase activity (Figure 5F). The side chain hydroxyl group of T19 seems of minor importance for SAM binding since mutations of T17 (T19 in VdTsr3) to either A or D did not significantly influence the SAM-binding affinity of SsTsr3 (KD's = 3.9 or 11.2 mM, respectively). Nevertheless, a mutation of the equivalent position S62 of ScTsr3 to D, but not to A, resulted in reduced acp modification in vivo, as shown by primer extension analysis (Figure 5F). The acp-transfer reaction catalyzed by Tsr3 most likely requires the presence of a catalytic base in order to abstract a proton from the N3 imino group of the modified pseudouridine. The side chain of D70 (VdTsr3) located in β4 is ∼5 Å away from the SAM sulfur atom. This residue is conserved as D or E both in archaeal and eukaryotic Tsr3 homologs. Mutations of the corresponding residue in SsTsr3 to A (D63) does not significantly alter the SAM-binding affinity of the protein (KD = 11.0 μM). However, the mutation of the corresponding residue of ScTsr3 (E111A) leads to a significant decrease of the acp transferase activity in vivo (Figure 5F). RNA-binding of Tsr3 Analysis of the electrostatic surface properties of VdTsr3 clearly identified positively charged surface patches in the vicinity of the SAM-binding site suggesting a putative RNA-binding site (Figure 6A). Furthermore, a negatively charged MES-ion is found in the crystal structure of VdTsr3 complexed to the side chain of K22 in helix α1. Its negatively charged sulfate group might mimic an RNA backbone phosphate. Helix α1 contains two more positively charged amino acids K17 and R25 as does the loop preceding it (R9). A second cluster of positively charged residues is found in or near helix α3 (K74, R75, K82, R85 and K87). Some of these amino acids are conserved between archaeal and eukaryotic Tsr3 (Supplementary Figure S1A). In the C-terminal domain, the surface exposed α-helices α5 and α7 carry a significant amount of positively charged amino acids. A triple mutation of the conserved positively charged residues R60, K65 and R131 to A in ScTsr3 resulted in a protein with a significantly impaired acp transferase activity in vivo (Figure 6D) in line with an important functional role for these positively charged residues. RNA-binding of Tsr3. (A) Electrostatic charge distribution on the surface of VdTsr3. Surface areas colored in blue are positively charged whereas red areas are negatively charged. SAM is shown in a stick representation. Also shown in stick representation is a negatively charged MES ion. Conserved basic amino acids are labeled. (B) Comparison of the secondary structures of helix 31 from the small ribosomal subunit rRNAs in S. cerevisiae and S. solfataricus with the location of the hypermodified nucleotide indicated in red. For S. solfataricus the chemical identity of the hypermodified nucleotide is not known but the existence of NEP1 and TSR3 homologs suggest that it is indeed N1-methyl-N3-acp-pseudouridine. (C) Binding of SsTsr3 to RNA. 5′-fluoresceine labeled RNA oligonucleotides corresponding either to the native (20mer – see inset) or a stabilized (20mer_GC - inset) helix 31 of the small ribosomal subunit rRNA from S. solfataricus were titrated with increasing amounts of SsTsr3 and the changes in the fluoresceine fluorescence anisotropy were measured and fitted to a binding curve (20mer – red, 20mer_GC – blue). Oligo-U9-RNA was used for comparison (black). The 20mer_GC RNA was also titrated with SsTsr3 in the presence of 2 mM SAM (purple). (D) Mutants of ScTsr3 R60, K65 or R131 (equivalent to K17, K22 and R91 in VdTsr3) expressed in Δtsr3 yeast cells show a primer extension stop comparable to the wild type. Combination of the three point mutations (R60A/K65A/R131A) leads to a strongly reduced acp modification of 18S rRNA. In order to explore the RNA-ligand specificity of Tsr3 we titrated SsTsr3 prepared in RNase-free form with 5′-fluoresceine-labeled RNA and determined the affinity by fluorescence anisotropy measurements. SsTsr3 in the apo state bound a 20mer RNA corresponding to helix 31 of S. solfataricus 16S rRNA (Figure 6B) with a KD of 1.9 μM and to a version of this hairpin stabilized by additional GC base pairs (20mer-GC) with a KD of 0.6 μM (Figure 6C). A single stranded oligoU-RNA bound with a 10-fold-reduced affinity (6.0 μM). The presence of saturating amounts of SAM (2 mM) did not have a significant influence on the RNA-affinity of SsTsr3 (KD of 1.7 μM for the 20mer-GC-RNA) suggesting no cooperativity in substrate binding. DISCUSSION U1191 is the only hypermodified base in the yeast 18S rRNA and is strongly conserved in eukaryotes. The formation of 1-methyl-3-(3-amino-3-carboxypropyl)-pseudouridine (m1acp3Ψ) is very complex requiring three successive modification reactions involving one H/ACA snoRNP (snR35) and two protein enzymes (Nep1/Emg1 and Tsr3). This makes it unique in eukaryotic rRNA modification. The m1acp3Ψ base is located at the tip of helix 31 on the 18S rRNA (Supplementary Figure S1B) which, together with helices 18, 24, 34 and 44, contribute to building the decoding center of the small ribosomal subunit. A similar modification (acp3U) was identified in Haloferax volcanii and corresponding modified nucleotides were also shown to occur in other archaea. As shown here TSR3 encodes the transferase catalyzing the acp modification as the last step in the biosynthesis of m1acp3Ψ in yeast and human cells. Unexpectedly, archaeal Tsr3 has a structure similar to SPOUT-class RNA methyltransferases, and it is the first example for an enzyme of this class transferring an acp group, due to a modified SAM-binding pocket that exposes the acp instead of the methyl group of SAM to its RNA substrate. Similar to the structurally unrelated Rossmann-fold Tyw2 acp transferase, the SAM methyl group of Tsr3 is bound in an inaccessible hydrophobic pocket whereas the acp side chain becomes accessible for a nucleophilic attack by the N3 of pseudouridine. In contrast, in the structurally closely related RNA methyltransferase Trm10 the methyl group of the cofactor SAM is accessible whereas its acp side chain is buried inside the protein. This suggests that enzymes with a SAM-dependent acp transferase activity might have evolved from SAM-dependent methyltransferases by slight modifications of the SAM-binding pocket. Thus, additional examples for acp transferase enzymes might be found with similarities to other structural classes of methyltransferases. In contrast to Nep1, the enzyme preceding Tsr3 in the m1acp3Ψ biosynthesis pathway, Tsr3 binds rather weakly and with little specificity to its isolated substrate RNA. This suggests that Tsr3 is not stably incorporated into pre-ribosomal particles and that its binding to the nascent ribosomal subunit possibly requires additional interactions with other pre-ribosomal components. Consistently, in sucrose gradient analysis, Tsr3 was found in low-molecular weight fractions rather than with pre-ribosome containing high-molecular weight fractions. In contrast to several enzymes that catalyze base specific modifications in rRNAs Tsr3 is not an essential protein. Typically, other small subunit rRNA methyltransferases as Dim1, Bud23 and Nep1/Emg1 carry dual functions, in ribosome biogenesis and rRNA modification, and it is their involvement in pre-RNA processing that is essential rather than their RNA-methylating activity (, discussed in 7). In contrast, for several Tsr3 mutants (SAM-binding and cysteine mutations) we found a systematic correlation between the loss of acp modification and the efficiency of 18S rRNA maturation. This demonstrates that, unlike the other small subunit rRNA base modifications, the acp modification is required for efficient pre-rRNA processing. Recently, structural, functional, and CRAC (cross-linking and cDNA analysis) experiments of late assembly factors involved in cytoplasmic processing of 40S subunits, along with cryo-EM studies of the late pre-40S subunits have provided important insights into late pre-40S processing. Apart from most of the ribosomal proteins, cytoplasmic pre-40S particles contain 20S rRNA and at least seven non-ribosomal proteins including the D-site endonuclease Nob1 as well as Tsr1, a putative GTPase and Rio2 which block the mRNA channel and the initiator tRNA binding site, respectively, thus preventing translation initiation. After structural changes, possibly driven by GTP hydrolysis, which go together with the formation of the decoding site, the 20S pre-rRNA becomes accessible for Nob1 cleavage at site D. This also involves joining of pre-40S and 60S subunits to 80S-like particles in a translation-like cycle promoted by eIF5B. The cleavage step most likely acts as a quality control check that ensures the proper 40S subunit assembly with only completely processed precursors. Finally, termination factor Rli1, an ATPase, promotes the dissociation of assembly factors and the 80S-like complex dissociates and releases the mature 40S subunit. Interestingly, differences in the level of acp modification were demonstrated for different steps of the cytoplasmic pre-40S subunit maturation after analyzing purified 20S pre-rRNAs using different purification bait proteins. Early cytoplasmic pre-40S subunits still containing the ribosome assembly factors Tsr1, Ltv1, Enp1 and Rio2 were not or only partially acp modified. In contrast, late pre-40S subunits containing Nob1 and Rio1 or already associated with 60S subunits in 80S-like particles showed acp modification levels comparable to mature 40S subunits. Thus, the acp transfer to m1Ψ1191 occurs during the step at which Rio2 leaves the pre-40S particle. These data and the finding that a missing acp modification hinders pre-20S rRNA processing, suggest that the acp modification together with the release of Rio2 promotes the formation of the decoding site and thus D-site cleavage by Nob1. The interrelation between acp modification and Rio2 release is also supported by CRAC analysis showing that Rio2 binds to helix 31 next to the Ψ1191 residue that receives the acp modification. Therefore, Rio2 either blocks the access of Tsr3 to helix 31, and acp modification can only occur after Rio2 is released, or the acp modification of m1Ψ1191 and putative subsequent conformational changes of 20S rRNA weaken the binding of Rio2 to helix 31 and support its release from the pre-rRNA. In summary, by identifying Tsr3 as the enzyme responsible for introducing the acp group to the hypermodified m1acp3Ψ nucleotide at position 1191 (yeast)/ 1248 (humans) of 18S rRNA we added one of the last remaining pieces to the puzzle of eukaryotic small ribosomal subunit rRNA modifications. The current data together with the finding that acp modification takes place at the very last step in pre-40S subunit maturation indicate that the acp modification probably supports the formation of the decoding site and efficient 20S pre-rRNA D-site cleavage. Furthermore, our structural data unravelled how the regioselectivity of SAM-dependent group transfer reactions can be tuned by distinct small evolutionary adaptions of the ligand binding pocket of SAM-binding enzymes. ACCESSION NUMBERS Coordinates and structure factors have been deposited in the Protein Data Bank under accession codes PDB 5APG (VdTsr3/SAM-complex) and PDB 5AP8 (SsTsr3). Supplementary Material SUPPLEMENTARY DATA Supplementary Data are available at NAR Online. FUNDING DFG grant [En134/9-1]; SFB 902 (Molecular Principles of RNA-based Regulation); DFG SPP1784 (Chemical Biology of Native Nucleic Acid Modifications, DFG grants) [En134/13-1, Wo 901/5-1]; European Community's Seventh Framework Programme [FP7/2007-2013] under BioStruct-X [283570]; Goethe University and the State of Hesse; EMBO long-term fellowship [ALTF 644-2014 to S.S.]; Research in the Lab of DLJL is supported by the Université Libre de Bruxelles (ULB); Fonds National de la Recherche (F.R.S./FNRS); Walloon Region [DGO6]; Fédération Wallonie-Bruxelles; European Research Development Fund (ERDF). Funding for open access charge: DFG SPP1784 (Chemical Biology of Native Nucleic Acid Modifications, DFG grants) [En134/13-1, Wo 901/5-1]. Conflict of interest statement. None declared. REFERENCES Ribosome biogenesis in the yeast Saccharomyces cerevisiae Diverse diseases from a ubiquitous process: the ribosomopathy paradox Genetics. 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ribosomal subunit biogenesis Proofreading of pre-40S ribosome maturation by a translation initiation factor and 60S subunits A translation-like cycle is a quality control checkpoint for maturing 40S ribosome 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+[{"sourceid":"4880283","sourcedb":"","project":"","target":"","text":"Crystal Structures of Putative Sugar Kinases from Synechococcus Elongatus PCC 7942 and Arabidopsis Thaliana The genome of the Synechococcus elongatus strain PCC 7942 encodes a putative sugar kinase (SePSK), which shares 44.9% sequence identity with the xylulose kinase-1 (AtXK-1) from Arabidopsis thaliana. Sequence alignment suggests that both kinases belong to the ribulokinase-like carbohydrate kinases, a sub-family of FGGY family carbohydrate kinases. However, their exact physiological function and real substrates remain unknown. Here we solved the structures of SePSK and AtXK-1 in both their apo forms and in complex with nucleotide substrates. The two kinases exhibit nearly identical overall architecture, with both kinases possessing ATP hydrolysis activity in the absence of substrates. In addition, our enzymatic assays suggested that SePSK has the capability to phosphorylate D-ribulose. In order to understand the catalytic mechanism of SePSK, we solved the structure of SePSK in complex with D-ribulose and found two potential substrate binding pockets in SePSK. Using mutation and activity analysis, we further verified the key residues important for its catalytic activity. Moreover, our structural comparison with other family members suggests that there are major conformational changes in SePSK upon substrate binding, facilitating the catalytic process. Together, these results provide important information for a more detailed understanding of the cofactor and substrate binding mode as well as the catalytic mechanism of SePSK, and possible similarities with its plant homologue AtXK-1. Introduction Carbohydrates are essential cellular compounds involved in the metabolic processes present in all organisms. Phosphorylation is one of the various pivotal modifications of carbohydrates, and is catalyzed by specific sugar kinases. These kinases exhibit considerable differences in their folding pattern and substrate specificity. Based on sequence analysis, they can be divided into four families, namely HSP 70_NBD family, FGGY family, Mer_B like family and Parm_like family. The FGGY family carbohydrate kinases contain different types of sugar kinases, all of which possess different catalytic substrates with preferences for short-chained sugar substrates, ranging from triose to heptose. These sugar substrates include L-ribulose, erythritol, L-fuculose, D-glycerol, D-gluconate, L-xylulose, D-ribulose, L-rhamnulose and D-xylulose. Structures reported in the Protein Data Bank of the FGGY family carbohydrate kinases exhibit a similar overall architecture containing two protein domains, one of which is responsible for the binding of substrate, while the second is used for binding cofactor ATP. While the binding pockets for substrates are at the same position, each FGGY family carbohydrate kinases uses different substrate-binding residues, resulting in high substrate specificity. Synpcc7942_2462 from the cyanobacteria Synechococcus elongatus PCC 7942 encodes a putative sugar kinase (SePSK), and this kinase contains 426 amino acids. The At2g21370 gene product from Arabidopsis thaliana, xylulose kinase-1 (AtXK-1), whose mature form contains 436 amino acids, is located in the chloroplast (ChloroP 1.1 Server). SePSK and AtXK-1 display a sequence identity of 44.9%, and belong to the ribulokinase-like carbohydrate kinases, a sub-family of FGGY family carbohydrate kinases. Members of this sub-family are responsible for the phosphorylation of sugars similar to L-ribulose and D-ribulose. The sequence and the substrate specificity of ribulokinase-like carbohydrate kinases are different, but they share the common folding feature with two domains. Domain I exhibits a ribonuclease H-like folding pattern, and is responsible for the substrate binding, while domain II possesses an actin-like ATPase domain that binds cofactor ATP. Two possible xylulose kinases (xylulose kinase-1: XK-1 and xylulose kinase-2: XK-2) from Arabidopsis thaliana were previously proposed. It was shown that XK-2 (At5g49650) located in the cytosol is indeed xylulose kinase. However, the function of XK-1 (At2g21370) inside the chloroplast stroma has remained unknown. SePSK from Synechococcus elongatus strain PCC 7942 is the homolog of AtXK-1, though its physiological function and substrates remain unclear. In order to obtain functional and structural information about these two proteins, here we reported the crystal structures of SePSK and AtXK-1. Our findings provide new details of the catalytic mechanism of SePSK and lay the foundation for future studies into its homologs in eukaryotes. Results and Discussion Overall structures of apo-SePSK and apo-AtXK-1 The attempt to solve the SePSK structure by molecular replacement method failed with ribulokinase from Bacillus halodurans (PDB code: 3QDK, 15.7% sequence identity) as an initial model. We therefore used single isomorphous replacement anomalous scattering method (SIRAS) for successful solution of the apo-SePSK structure at a resolution of 2.3 Å. Subsequently, the apo-SePSK structure was used as molecular replacement model to solve all other structures identified in this study. Our structural analysis showed that apo-SePSK consists of one SePSK protein molecule in an asymmetric unit. The amino-acid residues were traced from Val2 to His419, except for the Met1 residue and the seven residues at the C-termini. Apo-SePSK contains two domains referred to further on as domain I and domain II (Fig 1A). Domain I consists of non-contiguous portions of the polypeptide chains (aa. 2–228 and aa. 402–419), exhibiting 11 α-helices and 11 β-sheets. Among all these structural elements, α4/α5/α11/α18, β3/β2/β1/β6/β19/β20/β17 and α21/α32 form three patches, referred to as A1, B1 and A2, exhibiting the core region. In addition, four β-sheets (β7, β10, β12 and β16) and five α-helices (α8, α9, α13, α14 and α15) flank the left side of the core region. Domain II is comprised of aa. 229–401 and classified into B2 (β31/β29/β22/β23/β25/β24) and A3 (α26/α27/α28/α30) (Fig 1A and S1 Fig). In the SePSK structure, B1 and B2 are sandwiched by A1, A2 and A3, and the whole structure shows the A1/B1/A2/B2/A3 (α/β/α/β/α) folding pattern, which is in common with other members of FGGY family carbohydrate kinases (S2 Fig). The overall folding of SePSK resembles a clip, with A2 of domain I acting as a hinge region. As a consequence, a deep cleft is formed between the two domains. Overall structures of SePSK and AtXK-1. (A) Three-dimensional structure of apo-SePSK. The secondary structural elements are indicated (α-helix: cyan, β-sheet: yellow). (B) Three-dimensional structure of apo-AtXK-1. The secondary structural elements are indicated (α-helix: green, β-sheet: wheat). Apo-AtXK-1 exhibits a folding pattern similar to that of SePSK in line with their high sequence identity (Fig 1B and S1 Fig). However, superposition of structures of AtXK-1 and SePSK shows some differences, especially at the loop regions. A considerable difference is found in the loop3 linking β3 and α4, which is stretched out in the AtXK-1 structure, while in the SePSK structure, it is bent back towards the inner part. The corresponding residues between these two structures (SePSK-Lys35 and AtXK-1-Lys48) have a distance of 15.4 Å (S3 Fig). Activity assays of SePSK and AtXK-1 In order to understand the function of these two kinases, we performed structural comparison using Dali server. The structures most closely related to SePSK are xylulose kinase, glycerol kinase and ribulose kinase, implying that SePSK and AtXK-1 might function similarly to these kinases. We first tested whether both enzymes possessed ATP hydrolysis activity in the absence of substrates. As shown in Fig 2A, both SePSK and AtXK-1 exhibited ATP hydrolysis activity. This finding is in agreement with a previous result showing that xylulose kinase (PDB code: 2ITM) possessed ATP hydrolysis activity without adding substrate. To further identify the actual substrate of SePSK and AtXK-1, five different sugar molecules, including D-ribulose, L-ribulose, D-xylulose, L-xylulose and Glycerol, were used in enzymatic activity assays. As shown in Fig 2B, the ATP hydrolysis activity of SePSK greatly increased upon adding D-ribulose than adding other potential substrates, suggesting that it has D-ribulose kinase activity. In contrary, limited increasing of ATP hydrolysis activity was detected for AtXK-1 upon addition of D-ribulose (Fig 2C), despite its structural similarity with SePSK. The enzymatic activity assays of SePSK and AtXK-1. (A) The ATP hydrolysis activity of SePSK and AtXK-1. Both SePSK and AtXK-1 showed ATP hydrolysis activity in the absence of substrate. While the ATP hydrolysis activity of SePSK greatly increases upon addition of D-ribulose (DR). (B) The ATP hydrolysis activity of SePSK with addition of five different substrates. The substrates are DR (D-ribulose), LR (L-ribulose), DX (D-xylulose), LX (L-xylulose) and GLY (Glycerol). (C) The ATP hydrolysis activity of SePSK and AtXK-1 with or without D-ribulose. (D) The ATP hydrolysis activity of wild-type (WT) and single-site mutants of SePSK. Three single-site mutants of SePSK are D8A-SePSK, T11A-SePSK and D221A-SePSK. The ATP hydrolysis activity measured via luminescent ADP-Glo assay (Promega). To understand the catalytic mechanism of SePSK, we performed structural comparisons among xylulose kinase, glycerol kinase, ribulose kinase and SePSK. Our results suggested that three conserved residues (D8, T11 and D221 of SePSK) play an important role in SePSK function. Mutations of the corresponding residue in xylulose kinase and glycerol kinase from Escherichia coli greatly reduced their activity. To identify the function of these three residues of SePSK, we constructed D8A, T11A and D221A mutants. Using enzymatic activity assays, we found that all of these mutants exhibit much lower activity of ATP hydrolysis after adding D-ribulose than that of wild type, indicating the possibility that these three residues are involved in the catalytic process of phosphorylation D-ribulose and are vital for the function of SePSK (Fig 2D). SePSK and AtXK-1 possess a similar ATP binding site To obtain more detailed information of SePSK and AtXK-1 in complex with ATP, we soaked the apo-crystals in the reservoir adding cofactor ATP, and obtained the structures of SePSK and AtXK-1 bound with ATP at the resolution of 2.3 Å and 1.8 Å, respectively. In both structures, a strong electron density was found in the conserved ATP binding pocket, but can only be fitted with an ADP molecule (S4 Fig). Thus the two structures were named ADP-SePSK and ADP-AtXK-1, respectively. The extremely weak electron densities of ATP γ-phosphate in both structures suggest that the γ-phosphate group of ATP is either flexible or hydrolyzed by SePSK and AtXK-1. This result was consistent with our enzymatic activity assays where SePSK and AtXK-1 showed ATP hydrolysis activity without adding any substrates (Fig 2A and 2C). To avoid hydrolysis of ATP, we soaked the crystals of apo-SePSK and apo-AtXK-1 into the reservoir adding AMP-PNP. However, we found that the electron densities of γ-phosphate group of AMP-PNP (AMP-PNP γ-phosphate) are still weak in the AMP-PNP-SePSK and AMP-PNP-AtXK-1 structures, suggesting high flexibility of ATP-γ-phosphate. The γ-phosphate group of ATP is transferred to the sugar substrate during the reaction process, so this flexibility might be important for the ability of these kinases. The overall structures as well as the coordination modes of ADP and AMP-PNP in the AMP-PNP-AtXK-1, ADP-AtXK-1, ADP-SePSK and AMP-PNP-SePSK structures are nearly identical (S5 Fig), therefore the structure of AMP-PNP-SePSK is used here to describe the structural details and to compare with those of other family members. As shown in Fig 3A, one SePSK protein molecule is in an asymmetric unit with one AMP-PNP molecule. The AMP-PNP is bound at the domain II, where it fits well inside a positively charged groove. The AMP-PNP binding pocket consists of four α-helices (α26, α28, α27 and α30) and forms a shape resembling a half-fist (Fig 3A and 3B). The head group of the AMP-PNP is embedded in a pocket surrounded by Trp383, Asn380, Gly376 and Gly377. The purine ring of AMP-PNP is positioned in parallel to the indole ring of Trp383. In addition, it is hydrogen-bonded with the side chain amide of Asn380 (Fig 3B). The tail of AMP-PNP points to the hinge region of SePSK, and its α-phosphate and β-phosphate groups are stabilized by Gly376 and Ser243, respectively. Together, this structure clearly shows that the AMP-PNP-β-phosphate is sticking out of the ATP binding pocket, thus the γ-phosphate group is at the empty space between domain I and domain II and is unconstrained in its movement by the protein. Structure of SePSK in complex with AMP-PNP. (A) The electron density of AMP-PNP. The SePSK structure is shown in the electrostatic potential surface mode. The AMP-PNP is depicted as sticks with its ǀFoǀ-ǀFcǀ map contoured at 3 σ shown as cyan mesh. (B) The AMP-PNP binding pocket. The head of AMP-PNP is sandwiched by four residues (Leu293, Gly376, Gly377 and Trp383). The protein skeleton is shown as cartoon (cyan). The four α-helices (α26, α28, α27 and α30) are labeled in red. The AMP-PNP and coordinated residues are shown as sticks. The interactions between them are represented as black dashed lines. The numerical note near the black dashed line indicates the distance (Å). The potential substrate binding site in SePSK The results from our activity assays suggested that SePSK has D-ribulose kinase activity. To better understand the interaction pattern between SePSK and D-ribulose, the apo-SePSK crystals were soaked into the reservoir with 10 mM D-ribulose (RBL) and the RBL-SePSK structure was solved. As shown in S6 Fig, two residual electron densities are visible in domain I, which can be interpreted as two D-ribulose molecules with reasonable fit. As shown in Fig 4A, the nearest distance between the carbon skeleton of two D-ribulose molecules are approx. 7.1 Å (RBL1-C4 and RBL2-C1). RBL1 is located in the pocket consisting of α21 and the loop between β6 and β7. The O4 and O5 of RBL1 are coordinated with the side chain carboxyl group of Asp221. Furthermore, the O2 of RBL1 interacts with the main chain amide nitrogen of Ser72 (Fig 4B). This pocket is at a similar position of substrate binding site of other sugar kinase, such as L-ribulokinase (PDB code: 3QDK) (S7 Fig). However, structural comparison shows that the substrate ligating residues between the two structures are not strictly conserved. Based on the structures, the ligating residues of RBL1 in RBL-SePSK structure are Ser72, Asp221 and Ser222, and the interacting residues of L-ribulose with L-ribulokinase are Ala96, Lys208, Asp274 and Glu329 (S7 Fig). Glu329 in 3QDK has no counterpart in RBL-SePSK structure. In addition, although Lys208 of L-ribulokinase has the corresponding residue (Lys163) in RBL-SePSK structure, the hydrogen bond of Lys163 is broken because of the conformational change of two α-helices (α9 and α13) of SePSK. These differences might account for their different substrate specificity. The binding of D-ribulose (RBL) with SePSK. (A) The electrostatic potential surface map of RBL-SePSK and a zoom-in view of RBL binding site. The RBL1 and RBL2 are depicted as sticks. (B) Interaction of two D-ribulose molecules (RBL1 and RBL2) with SePSK. The RBL molecules (carbon atoms colored yellow) and amino acid residues of SePSK (carbon atoms colored green) involved in RBL interaction are shown as sticks. The hydrogen bonds are indicated by the black dashed lines and the numbers near the dashed lines are the distances (Å). (C) The binding affinity assays of SePSK with D-ribulose. Single-cycle kinetic data are reflecting the interaction of SePSK and D8A-SePSK with D-ribulose. It shows two experimental sensorgrams after minus the empty sensorgrams. The original data is shown as black curve, and the fitted data is shown as different color (wild type SePSK: red curve, D8A-SePSK: green curve). Dissociation rate constant of wild type and D8A-SePSK are 3 ms-1 and 9 ms-1, respectively. The binding pocket of RBL2 with relatively weak electron density is near the N-terminal region of SePSK and is negatively charged. The side chain of Asp8 interacts strongly with O3 and O4 of RBL2. The hydroxyl group of Ser12 coordinates with O2 of RBL2. The backbone amide nitrogens of Gly13 and Arg15 also keep hydrogen bonds with RBL2 (Fig 4B). Structural comparison of SePSK and AtXK-1 showed that while the RBL1 binding pocket is conserved, the RBL2 pocket is disrupted in AtXK-1 structure, despite the fact that the residues interacting with RBL2 are highly conserved between the two proteins. In the RBL-SePSK structure, a 2.6 Å hydrogen bond is present between RBL2 and Ser12 (Fig 4B), while in the AtXK-1 structure this hydrogen bond with the corresponding residue (Ser22) is broken. This break is probably induced by the conformational change of the two β-sheets (β1 and β2), with the result that the linking loop (loop 1) is located further away from the RBL2 binding site. This change might be the reason that AtXK-1 only shows limited increasing in its ATP hydrolysis ability upon adding D-ribulose as a substrate after comparing with SePSK (Fig 2C). Our SePSK structure shows that the Asp8 residue forms strong hydrogen bond with RBL2 (Fig 4B). In addition, our enzymatic assays indicated that Asp8 is important for the activity of SePSK (Fig 2D). To further verified this result, we measured the binding affinity for D-ribulose of both wild type (WT) and D8A mutant of SePSK using a surface plasmon resonance method. The results showed that the affinity of D8A-SePSK with D-ribulose is weaker than that of WT with a reduction of approx. two third (Fig 4C). Dissociation rate constant (Kd) of wild type and D8A-SePSK are 3 ms-1 and 9 ms-1, respectively. The results implied that the second RBL binding site plays a role in the D-ribulose kinase function of SePSK. However, considering the high concentration of D-ribulose used for crystal soaking, as well as the relatively weak electron density of RBL2, it is also possible that the second binding site of D-ribulose in SePSK is an artifact. Simulated conformational change of SePSK during the catalytic process It was reported earlier that the crossing angle between the domain I and domain II in FGGY family carbohydrate kinases is different. In addition, this difference may be caused by the binding of substrates and/or ATP. As reported previously, members of the sugar kinase family undergo a conformational change to narrow the crossing angle between two domains and reduce the distance between substrate and ATP in order to facilitate the catalytic reaction of phosphorylation of sugar substrates. After comparing structures of apo-SePSK, RBL-SePSK and AMP-PNP-SePSK, we noticed that these structures presented here are similar. Superposing the structures of RBL-SePSK and AMP-PNP-SePSK, the results show that the nearest distance between AMP-PNP γ-phosphate and RBL1/RBL2 is 7.5 Å (RBL1-O5)/6.7 Å (RBL2-O1) (S8 Fig). This distance is too long to transfer the γ-phosphate group from ATP to the substrate. Since the two domains of SePSK are widely separated in this structure, we hypothesize that our structures of SePSK represent its open form, and that a conformational rearrangement must occur to switch to the closed state in order to facilitate the catalytic process of phosphorylation of sugar substrates. For studying such potential conformational change, a simulation on the Hingeprot Server was performed to predict the movement of different SePSK domains. The results showed that domain I and domain II are closer to each other with Ala228 and Thr401 in A2 as Hinge-residues. Based on the above results, SePSK is divided into two rigid parts. The domain I of RBL-SePSK (aa. 1–228, aa. 402–421) and the domain II of AMP-PNP-SePSK (aa. 229–401) were superposed with structures, including apo-AtXK-1, apo-SePSK, xylulose kinase from Lactobacillus acidophilus (PDB code: 3LL3) and the S58W mutant form of glycerol kinase from Escherichia coli (PDB code: 1GLJ). The results of superposition displayed different crossing angle between these two domains. After superposition, the distances of AMP-PNP γ-phosphate and the fifth hydroxyl group of RBL1 are 7.9 Å (superposed with AtXK-1), 7.4 Å (superposed with SePSK), 6.6 Å (superposed with 3LL3) and 6.1 Å (superposed with 1GLJ). Meanwhile, the distances of AMP-PNP γ-phosphate and the first hydroxyl group of RBL2 are 7.2 Å (superposed with AtXK-1), 6.7 Å (superposed with SePSK), 3.7 Å (superposed with 3LL3), until AMP-PNP γ-phosphate fully contacts RBL2 after superposition with 1GLJ (Fig 5). This distance between RBL2 and AMP-PNP-γ-phosphate is close enough to facilitate phosphate transferring. Together, our superposition results provided snapshots of the conformational changes at different catalytic stages of SePSK and potentially revealed the closed form of SePSK. Simulated conformational change of SePSK during the catalytic process. The structures are shown as cartoon and the ligands are shown as sticks. Domain I from D-ribulose-SePSK (green) and Domain II from AMP-PNP-SePSK (cyan) are superposed with apo-AtXK-1 (1st), apo-SePSK (2nd), 3LL3 (3rd) and 1GLJ (4th), respectively. The numbers near the black dashed lines show the distances (Å) between two nearest atoms of RBL and AMP-PNP. In summary, our structural and enzymatic analyses provide evidence that SePSK shows D-ribulose kinase activity, and exhibits the conserved features of FGGY family carbohydrate kinases. Three conserved residues in SePSK were identified to be essential for this function. Our results provide the detailed information about the interaction of SePSK with ATP and substrates. Moreover, structural superposition results enable us to visualize the conformational change of SePSK during the catalytic process. In conclusion, our results provide important information for a more detailed understanding of the mechanisms of SePSK and other members of FGGY family carbohydrate kinases. Materials and Methods Cloning, expression and purification of SePSK The gene encoding SePSK was amplified by polymerase chain reaction (PCR) with forward primer 5' CATGCCATGGGCATGGTCGTTGCACTTGGCCTCG 3' containing an internal Nco I restriction site (underlined) and reverse primer 5' CCGCTCGAGGGTTCTCTTTAACCCCGCCG 3' including an internal Xho I restriction site (underlined) from Synechococcus elongatus PCC 7942 genomic DNA. The amplified PCR product was digested with Nco I and Xho I (Takara) and ligated into linearized pET28-a vector (Novagen) between Nco I and Xho I restriction sites with a C-terminal his6 tag. The recombinant plasmids were transformed into competent Escherichia coli Trans10 cells for DNA production and purification, and the final constructs were verified by sequencing. The recombinant vectors were transformed into Escherichia coli BL21 (DE3) to express the protein. After induction with the 1 mM IPTG at 289 K in Luria-Bertani medium until the cell density reached an OD 600 nm of 0.6–0.8, the cells were harvested by centrifugation at 6,000 g at 4°C for 15 min, re-suspended in buffer A (20 mM Tris-HCl, pH 8.0, 500 mM NaCl, 5 mM imidazole) and disrupted by sonication. After centrifuge 40,000 g for 30 min, the protein was purified by passage through a Ni2+ affinity column in buffer A, and then washed the unbound protein with buffer B (20 mM Tris-HCl, pH 8.0, 500 mM NaCl, 60 mM imidazole), and eluted the fraction with the buffer C (20 mM Tris-HCl, pH 8.0, 500 mM NaCl, 500 mM imidazole). After that, the protein was further purified by size exclusion chromatography with Superdex 200 10/300 GL (GE Healthcare) equilibrated with the buffer D (20 mM Tris-HCl, pH 8.0, 300 mM NaCl). The eluted major peak fraction was concentrated to 20 mg/mL protein using 10,000 MCWO centrifugal filter units (Millipore) and stored at -80°C for crystallization trials. The purified product was analyzed by SDS-PAGE with a single band visible only. Cloning, expression and purification of AtXK-1 The gene encoding AtXK-1 was amplified by PCR using a forward primer 5' TACTTCCAATCCAATGCTGTTATGAGTGGCAATAAAGGAACGA 3' and reverse primer 5' TTATCCACTTCCAATGTTACAAACCACTGTTCTGTTTTGCGCCC 3' from cDNA library of Arabidopsis thaliana. The underlined nucleotides were used for the ligation-independent cloning. The PCR product was treated by T4 DNA polymerase (LIC-qualified, Novagen) and then cloned into linearized pMCSG7 vector treated by T4 DNA polymerase (LIC-qualified, Novagen) with an N-terminal his6 tag though ligation-independent cloning method. The final construct was confirmed by DNA sequencing after amplified in competent Escherichia coli Trans10 cells. The recombinant vectors were transformed into Escherichia coli BL21 (DE3) for protein expression. After induction with 1 mM IPTG at 289 K in Luria-Bertani medium, cells were grown until the cell density reached an OD 600 nm of 0.6–0.8. Subsequent purification was identical to that used for SePSK, except that there was one additional step, during which tobacco etch virus protease was used to digest the crude AtXK-1 protein for removal of the N-terminal his6 tag following Ni2+ affinity purification. Ni2+ affinity column buffer contained extra 20% glycerol. The protein was further purified by size exclusion chromatography with Superdex 200 10/300 GL (GE Healthcare) in elution buffer consisting of 20 mM HEPES, pH 7.5, 100 mM NaCl. Finally, AtXK-1 protein was concentrated to 40 mg/mL protein using 10,000 MCWO centrifugal filter units (Millipore) and stored at -80°C prior to crystallization trials. Purity was verified by SDS-PAGE with a single band visible only. Site-directed mutagenesis of SePSK The gene of D8A and T11A mutations were amplified by PCR with the forward primer 5' CATGCCATGGGCATGGTCGTTGCACTTGGCCTCGCCTTCGGCAC 3' and forward primer 5' CATGCCATGGGCATGGTCGTTGCACTTGGCCTCGACTTCGGCGCCTCTGGAGCCC 3' (mismatched base pairs are underlined). The reverse primers of D8A and T11A mutants, the further constructions and purification procedures were identical with those used for wild type SePSK. The N-terminal sequence of D221A was amplified with forward primer (T7 promoter primer) 5' TAATACGACTCACTATA 3' and reverse primer 5' AGCAGCAATGCTAGCCGTTGTACCG 3’, and the C-terminal sequence of D221A was amplified with forward primer 5' TGCCGGTACAACGGCTAGCATTGCT 3' and reverse primer (T7 terminator primer) CGATCAATAACGAGTCGCC (mismatched base pairs are underlined). The second cycle PCR used the above PCR products as templates, and the construction and purification procedures were identical to those used for wild type SePSK. Crystallization and data collection Crystallization trials of SePSK and AtXK-1 were carried out at 281 K by mixing equal volume of 20 mg/ml protein and reservoir solution with the sitting-drop vapor diffusion method. The reservoir solution was PEG Rx I-35 (0.1 M BIS-TRIS pH 6.5, 20% w/v Polyethylene glycol monomethyl ether 5,000) (Hampton research). After 2 or 3 days, the rod-like crystals could be observed. For phasing, the high-quality apo-SePSK crystals were soaked in mother liquor containing 1 mM ethylmercuricthiosalicylic acid, sodium salt (Hampton research, heavy atom kit) overnight at 281 K. In order to get the complexes with ATP and AMP-PNP, the crystals of apo-SePSK and apo-AtXK-1 were incubated with the reservoir including 10 mM ATP and 20 mM MgCl2 as well as 10 mM AMP-PNP and 20 mM MgCl2, respectively. The apo-SePSK crystals were incubated with the reservoir including 10 mM D-ribulose in order to obtain the complex D-ribulose-bound SePSK (RBL-SePSK). The crystals of three mutants (D8A, T11A and D221A) grew in the same condition as that of the wild type SePSK. The crystals were dipped into reservoir solution supplemented with 15% glycerol and then flash frozen in a nitrogen gas stream at 100 K. All data sets were collected at Shanghai Synchrotron Radiation Facility, Photo Factory in Japan and Institute of Biophysics, Chinese Academy of Sciences. Diffraction data were processed using the HKL2000 package. Structure determination and refinement The initial phases of SePSK were obtained from the Hg-derivative crystals by single isomorphous replacement anomalous scattering (SIRAS) using AutoSol from the PHENIX suite. AutoBuild from the PHENIX suite was used to build 75% of the main chain of apo-SePSK, and the remaining residues were built manually by Coot. All other structures were solved by molecular replacement method using apo-SePSK as an initial model. The model was refined using phenix.refine and REFMAC5. The final model was checked with PROCHECK. All structural figures were prepared by PyMOL. The summary of the data-collection and structure-refinement statistics is shown in Table 1 and S1 Table. Atomic coordinates and structure factors in this article have been deposited in the Protein Data Bank. The deposited codes of all structures listed in the Table 1 and S1 Table. Data collection and refinement statistics. Data set\tHg-SePSK\tapo-SePSK\tAMP-PNP-SePSK\tRBL-SePSK\tapo-AtXK-1\t \tData collection\t\t\t\t\t\t \tSpace group\tC 1 2 1\tC 1 2 1\tC 1 2 1\tC 1 2 1\tP21\t \tWavelength (Å)\t1.54178\t1.54178\t1.54178\t1.54178\t1.54178\t \tCell parameters\t\t\t\t\t\t \ta/b/c(Å)\t103.1, 46.6, 88.3\t110.2, 49.0, 86.9\t103.5, 46.6, 88.0\t102.6, 47.0, 88.7\t49.7, 87.9, 53.6\t \tα/β/γ(°)\t90.0, 91.9, 90.0\t90.0, 110.3, 90.0\t90.0, 91.0, 90.0\t90.0, 91.4, 90.0\t90.0, 97.0, 90.0\t \tResolution (Å)a\t50.00–2.20(2.28–2.20)\t50.00–2.30(2.38–2.30)\t50.00–2.30(2.38–2.30)\t50.00–2.35(2.43–2.35)\t50.00–2.00(2.07–2.00)\t \tR mergeb\t0.105(0.514)\t0.149(0.501)\t0.082(0.503)\t0.095(0.507)\t0.106(0.454)\t \t〈 I/σ(I)〉\t28.89(4.07)\t13.85(4.10)\t10.18(1.79)\t19.4(4.6)\t12.91(4.08)\t \tCompleteness (%)\t92.3(99.2)\t96.1(94.2)\t98.9(99.8)\t99.8(100.0)\t97.1(94.5)\t \tRedundancy\t6.7(5.1)\t7.4(7.5)\t2.4(2.4)\t6.9(6.7)\t7.2(6.9)\t \tRefinement statistics\t\t\t\t\t\t \tResolution (Å)\t\t32.501–2.301\t24.707–2.300\t24.475–2.344\t23.771–1.998\t \tRwork/ Rfreec\t\t0.1834/0.2276\t0.1975/0.2327\t0.2336/0.2687\t0.1893/0.2161\t \tNo. atoms\t\t\t\t\t\t \tProtein\t\t3503\t3196\t3209\t3256\t \tligand/ion\t\t-\t31\t20\t-\t \tWater\t\t313\t146\t143\t486\t \tRMSD Bond lengths (Å)d\t\t0.003\t0.005\t0.003\t0.003\t \tRMSD Bond angles (°)d\t\t0.674\t0.886\t0.649\t0.838\t \tRamachandran plot (%)\t\t\t\t\t\t \tfavoured\t\t98.1\t97.8\t96.7\t99.1\t \tallowed\t\t1.9\t2.2\t3.3\t0.9\t \tdisallowed\t\t0.0\t0.0\t0.0\t0.0\t \tPDB code\t\t5HTN\t5HTP\t5HV7\t5HTR\t \t a The values in parentheses correspond to the highest resolution shell. b Rmerge = ∑j∑h|Ij,h-\u003cIh\u003e|/∑j∑h\u003cIh\u003e where h are unique reflection indices and Ij,h are intensities of symmetry-related reflections and \u003cIh\u003e is the mean intensity. c R-work and R-free were calculated as follows: R = Σ (|Fobs-Fcalc|)/Σ |Fobs| ×100, where Fobs and Fcalc are the observed and calculated structure factor amplitudes, respectively. d Root mean square deviations (r.m.s.d.) from standard values. ADP-Glo kinase assay ADP-Glo kinase assay was used according to the manufacturer’s instructions (Promega). Each reaction mixture system consisted of 8 uM enzyme, 100 uM ATP, 1 mM MgCl2, 20 mM HEPES (pH 7.4), 5 mM substrate. The reaction was initiated by adding the purified enzyme into the reaction system. After incubation at 298 K for different time, equal volume ADP-Glo™ reagent was added to terminate the kinase reaction and to deplete any remaining ATP. Subsequently, kinase detection reagent with double volume of reaction system was added to convert ADP to ATP and allowed the newly synthesized ATP to be measured using a luciferase/luciferin reaction which produced luminescence signal and could be recorded. After incubation at room temperature for another 60 min, luminescence was detected by Varioskan Flash Multimode Reader (Thermo). The reference experiment was carried out in the same reaction system without the enzyme. For each assay, at least three repeats were performed for the calculation of mean values and standard deviations (SDs). The purity of five substrates in the activity assays was ≥98% (D-ribulose, Santa cruz), 99.7% (L-ribulose, Carbosynth), 99.3% (D-xylulose, Carbosynth), 99.5% (L-xylulose, Carbosynth) and 99.0% (Glycerol, AMRESCO). Surface plasmon resonance Surface plasmon resonance (SPR) was used to analyze the interaction of SePSK and D-ribulose. The SPR experiments were performed on a Biacore T100 system using series S CM5 sensor chips (GE Healthcare). All sensorgrams were recorded at 298 K. The proteins in buffer containing 20 mM HEPES, pH 7.5, 100 mM NaCl, was diluted to 40 ug/ml by 10 mM sodium acetate buffer at pH 4.5. All flow cells on a CM5 sensor chip were activated with a freshly prepared solution of 0.2 M 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDC) and 0.05 M N-hydroxysuccinimide (NHS) in a ratio of 1:1 at a constant flow rate of 10 ul/min for 420 s. Deactivation of the surface was performed with an injection of a 1 M solution of ethanolamine-HCl (pH 8.5) using the same flow rate and duration. Kinetic parameters were derived from data sets acquired in single-cycle mode. Each run consisted of five consecutive analytic injections at 125, 250, 500, 1000 and 2000 uM. Analytic injections lasted for 60 s, separated by 30 s dissociation periods. Each cycle was completed with an extended dissociation period of 300 s. The specific binding to a blank flow cell was subtracted to obtain corrected sensorgrams. Biacore data were analyzed using BiaEvaluation software (GE Healthcare) by fitting to a 1:1 Langmuir binding fitting model. Accession Codes Coordinates and structure factors for all the structures in this article have been deposited in the Protein Data Bank. These accession codes are 5HTN, 5HTP, 5HUX, 5HV7, 5HTJ, 5HU2, 5HTY, 5HTR, 5HTV and 5HTX. The corresponding-structures are apo-SePSK, AMP-PNP-SePSK, ADP-SePSK, RBL-SePSK, D8A-SePSK, T11A-SePSK, D221A-SePSK, apo-AtXK1, AMP-PNP-AtXK1 and ADP-AtXK1, respectively. Supporting Information References The FGGY carbohydrate kinase family: insights into the evolution of functional specificities Structure and function of human xylulokinase, an enzyme with important roles in carbohydrate metabolism Structural insight into mechanism and diverse substrate selection strategy of L-ribulokinase Structural and Kinetic Studies of Induced Fit in Xylulose Kinase from Escherichia coli ChloroP, a neural network-based method for predicting chloroplast transit peptides and their cleavage sites A cytosolic Arabidopsis D-xylulose kinase catalyzes the phosphorylation of 1-deoxy-D-xylulose into a precursor of the plastidial isoprenoid pathway Crystal structures of Escherichia coli glycerol kinase variant S58→ W in complex with nonhydrolyzable ATP analogues reveal a putative active conformation of the enzyme as a result of domain motion Glycerol kinase from Escherichia coli and an Ala65→ Thr mutant: the crystal structures reveal conformational changes with implications for allosteric regulation Structure and Reaction Mechanism of l-Rhamnulose Kinase from Escherichia coli Structural and Functional Analysis of Fucose-Processing Enzymes from Streptococcus pneumoniae Conserved active site aspartates and domain–domain interactions in regulatory properties of the sugar kinase superfamily Analyzing a kinetic titration series using affinity biosensors Structures of enterococcal glycerol kinase in the absence and presence of glycerol: correlation of conformation to substrate binding and a mechanism of activation by phosphorylation The sugar kinase/heat shock protein 70/actin superfamily: implications of conserved structure for mechanism HingeProt: automated prediction of hinges in protein structures A new vector for high-throughput, ligation-independent cloning encoding a tobacco etch virus protease cleavage site Processing of X-ray diffraction data collected in oscillation mode PHENIX: a comprehensive Python-based system for macromolecular structure solution Coot: model-building tools for molecular graphics On macromolecular refinement at subatomic resolution with interatomic scatterers PROCHECK: a program to check the stereochemical quality of protein structures DeLano. W. The PyMOL Molecular Graphics System. Available: http://www.pymol.org. Comparison of the luminescent ADP-Glo assay to a standard radiometric assay for measurement of protein kinase 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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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+[{"sourceid":"4887163","sourcedb":"","project":"","target":"","text":"Hotspot autoimmune T cell receptor binding underlies pathogen and insulin peptide cross-reactivity The cross-reactivity of T cells with pathogen- and self-derived peptides has been implicated as a pathway involved in the development of autoimmunity. However, the mechanisms that allow the clonal T cell antigen receptor (TCR) to functionally engage multiple peptide–major histocompatibility complexes (pMHC) are unclear. Here, we studied multiligand discrimination by a human, preproinsulin reactive, MHC class-I–restricted CD8+ T cell clone (1E6) that can recognize over 1 million different peptides. We generated high-resolution structures of the 1E6 TCR bound to 7 altered peptide ligands, including a pathogen-derived peptide that was an order of magnitude more potent than the natural self-peptide. Evaluation of these structures demonstrated that binding was stabilized through a conserved lock-and-key–like minimal binding footprint that enables 1E6 TCR to tolerate vast numbers of substitutions outside of this so-called hotspot. Highly potent antigens of the 1E6 TCR engaged with a strong antipathogen-like binding affinity; this engagement was governed though an energetic switch from an enthalpically to entropically driven interaction compared with the natural autoimmune ligand. Together, these data highlight how T cell cross-reactivity with pathogen-derived antigens might break self-tolerance to induce autoimmune disease. Introduction T cells perform an essential role in adaptive immunity by interrogating the host proteome for anomalies, classically by recognizing peptides bound in major histocompatibility (MHC) molecules at the cell surface. Recent data supports the notion that, to perform this role, the highly variable αβ T cell antigen receptor (TCR) must be able to recognize thousands, if not millions, of different peptide ligands. This ability is required to enable the estimated 25 million distinct TCRs expressed in humans to provide effective immune coverage against all possible foreign peptide antigens. Although essential to avoid blind spots during pathogen recognition, T cell cross-reactivity has also been implicated as a pathway to autoimmunity, possibly mediated by highly reactive pathogen-specific T cells weakly recognizing self-ligands. Several mechanisms, by which TCRs could bind to a large number of different peptide-MHC (pMHC), have been proposed. Structures of unligated and ligated TCRs have shown that the TCR complementarity determining region (CDR) loops can be flexible, perhaps enabling peptide binding using different loop conformations. Both MHC and peptide have also been shown to undergo structural changes upon TCR binding, mediating an induced fit between the TCR and pMHC. Other studies, mainly in the murine system, have demonstrated that the same TCR can interact with different pMHCs using a common or divergent modality. Recent studies in model murine systems demonstrate that TCR cross-reactivity can be governed by recognition of a conserved region in the peptide that allows tolerance of peptide sequence variation outside of this hotspot. We recently reported that the 1E6 human CD8+ T cell clone — which mediates the destruction of β cells through the recognition of a major, HLA-A*0201–restricted, preproinsulin signal peptide (ALWGPDPAAA15–24) — can recognize upwards of 1 million different peptides. CD8+ T cells that recognize HLA-A*0201–ALWGPDPAAA have been shown to populate insulitic lesions in patients with type 1 diabetes (T1D). We demonstrated that the TCR from the 1E6 T cell clone bound to HLA-A*0201–ALWGPDPAAA using a limited footprint and very weak binding affinity. This first experimental evidence of a high level of CD8+ T cell cross-reactivity in a human autoimmune disease system hinted toward molecular mimicry by a more potent pathogenic peptide as a potential mechanism leading to β cell destruction. Here, we solved the structure of the 1E6 TCR with 7 altered peptide ligands (APLs) determined by our previously published combinatorial peptide library (CPL) screening, 2 of which mapped within human pathogens. These APLs differed from the natural preproinsulin peptide by up to 7 of 10 residues. We also solved the structure of each unligated APL to investigate whether structural changes occurred before or after binding — which, combined with an in-depth cellular and biophysical analysis of the 1E6 interaction with each APL, demonstrated the molecular mechanism mediating the high level of cross-reactivity exhibited by this preproinsulin-reactive human CD8+ T cell clone. Results The 1E6 T cell clone recognizes APLs across a large dynamic range. We have previously demonstrated that the 1E6 T cell clone can recognize over 1 million different peptides with a potency comparable with, or better than, the cognate preproinsulin peptide ALWGPDPAAA. From this large functional scan, we selected 7 different APLs that activated the 1E6 T cell clone across a wide (4-log) functional range (Table 1). Two of these peptides, MVWGPDPLYV and RQFGPDWIVA (bold text signifies amino acids that are different from the index preproinsulin–derived sequence), are contained within the proteomes of the human pathogens Bacteroides fragilis/thetaiotaomicron and Clostridium asparagiforme, respectively. Competitive functional testing revealed that the preproinsulin-derived sequence ALWGPDPAAA was one of the least potent targets for 1E6, with only the MVWGPDPLYV and YLGGPDFPTI demonstrating a similar low-activity profile in MIP-1β secretion and target killing assays (Figure 1, A and B). The RQFGPDWIVA sequence (present in C. asparagiforme) activated the 1E6 T cell with around 1 log–greater potency compared with ALWGPDPAAA. At the other end of the spectrum, the RQFGPDFPTI peptide stimulated MIP-1β release and killing by 1E6 at an exogenous peptide concentration 2–3 logs lower compared with ALWGPDPAAA. The pattern of peptide potency was closely mirrored by pMHC tetramer staining experiments (Figure 1C and plots shown in Supplemental Figure 1; supplemental material available online with this article; doi:10.1172/JCI85679DS1). Here, the A2-RQFGPDFPTI tetramer stained 1E6 with the greatest MFI, gradually decreasing to the weakest tetramers: A2-MVWGPDPLYV and -YLGGPDFPTI. To parallel the functional analysis, we also performed thermal melt (Tm) experiments using synchrotron radiation circular dichroism (SRCD) to investigate the stability of each APL (Figure 1D). The range of Tm was between 49.4°C (RQFGPDWIVA) and 60.3°C (YQFGPDFPIA), with an average approximately 55°C, similar to our previous findings. This pattern of stability did not correlate with the T cell activation or tetramer staining experiments, indicating that peptide binding to the MHC do not explain ligand potency. The 1E6 TCR can bind peptides with strong antipathogen-like affinities. We, and others, have previously demonstrated that antipathogenic TCRs tend to bind with stronger affinity compared with self-reactive TCRs, likely a consequence of the deletion of T cells with high-affinity self-reactive TCR during thymic selection. In accordance with this trend, the 1E6 TCR bound the natural preproinsulin peptide, ALWGPDPAAA, with the weakest affinity currently published for a human CD8+ T cell–derived TCR with a biologically relevant ligand (KD \u003e 200 μM; KD, equilibrium binding constant). Surface plasmon resonance (SPR) analysis of the 1E6 TCR–pMHC interaction for all 7 APLs (Figure 2, A–H) demonstrated that stronger binding affinity (represented as ΔG°, kcal/mol) correlated well with the EC50 values (peptide concentration required to reach half-maximal 1E6 T cell killing) for each ligand, demonstrated by a Pearson’s correlation analysis value of 0.8 (P = 0.01) (Figure 2I). It should be noted that this correlation, although consistent with the T cell killing experiments, uses only approximate affinities calculated for the 2 weakest ligands. These experiments revealed 4 important findings. First, the 1E6 T cell could still functionally respond to peptide when the TCR binding affinity was extremely weak, e.g., the 1E6 TCR binding affinity for the A2-MVWGPDPLYV peptide was KD = ~600 μM. Second, the 1E6 TCR bound to A2-RQFGPDFPTI with KD = 0.5 μM, equivalent to the binding affinity of the very strongest antipathogen TCRs. Third, the 1E6 TCR bound to A2-RQFGPDWIVA peptide, within the C. asparagiforme proteome, with approximately 4-fold stronger affinity than A2-ALWGPDPAAA, demonstrating the potential for a pathogen-derived antigen to initiate a response to the self-derived sequence. Finally, these data demonstrate the largest range of binding affinities reported for a natural, endogenous human TCR of more than 3 logs of magnitude (A2-MVWGPDPLYV vs. A2-RQFGPDFPTI). To confirm the affinity spread detected by SPR, and to evaluate whether experiments performed using soluble molecules were biologically relevant to events at the T cell surface, we determined the effective 2D affinity of each APL using an adhesion frequency assay in which a human rbc coated in pMHC acted as an adhesion sensor. In agreement with SPR experiments, the range of 2D affinities we detected differed by around 3 logs, with the A2-MVWGPDPLYV generating the weakest 2D affinity (2.6 × 10–5 AcKa μm4) and A2-RQFGPDFPTI the strongest (4.5 × 10–2 AcKa μm4) (Figure 2J). As with the 3D affinity measurements, the 2D affinity measurements correlated well with the EC50 values for each ligand (Figure 2K) demonstrating a strong correlation (Pearson’s correlation = 0.8, P = 0.01) between T cell antigen sensitivity and TCR binding affinity. Of note, these data demonstrate a close agreement between the 3D affinity values generated using SPR and 2D affinity values generated using adhesion frequency assays. The 1E6 TCR uses a consensus binding mode to engage multiple APLs. Our previous structure of the 1E6-A2-ALWGPDPAAA complex demonstrated a limited binding footprint between the TCR and pMHC. The low number of contacts between the 2 molecules most likely contributed to the weak binding affinity of the interaction. In order to examine the mechanism by which the 1E6 TCR engaged a wide range of peptides with divergent binding affinities, we solved the structure of the 1E6 TCR in complex with all 7 APLs used in Figure 2. All structures were solved in space group P1 to 2–3 Å resolution with crystallographic Rwork/Rfree ratios within accepted limits as shown in the theoretically expected distribution (ref. and Supplemental Table 1). The 1E6 TCR used a very similar overall binding modality to engage all of the APLs, with root mean square deviation ranging between 0.81 and 1.12 Å2 (compared with 1E6-A2-ALWGPDPAAA). The relatively broad range of buried surface areas (1,670–1,920 Å2) did not correlate well with TCR binding affinity (Pearson’s correlation = 0.45, P = 0.2). The surface complementarity values (0.52–0.7) correlated slightly with affinity (Pearson’s correlation = 0.7, P = 0.05) but could not explain all differences in binding (Figure 3A and Table 2). The TCR CDR loops were in a very similar position in all complexes, apart from some slight deviations in the TCR β-chain (Figure 3B); the peptides were all presented in a similar conformation (Figure 3C); and there was minimal variation in crossing angles of the TCR (42.3°–45.6°) (Figure 3D). Overall, the 1E6 TCR used a canonical binding mode to engage each APL with the TCR α-chain positioned over the MHC class I (MHCI) α2-helix and the TCR β-chain over the MHCI α-1 helix, straddling the peptide cargo. However, subtle differences in the respective interfaces were apparent (discussed below) and resulted in altered binding affinities of the respective complexes. Interactions between the 1E6 TCR and different APLs are focused around a conserved GPD peptide motif. We next performed an in-depth atomic analysis of the contacts between the 1E6 TCR and each APL to determine the structural basis for the altered T cell peptide sensitivities and TCR binding affinities (Table 2). Concomitant with our global analysis of 1E6 TCR binding to the APLs, we observed a common interaction element, consistent with our previous findings, that utilized TCR residues Tyr97α and Trp97β, forming an aromatic cap over a central GPD motif that was present in all of the APLs (Figure 4). Interactions between these 2 TCR and 3 peptide residues accounted for 41%–50% of the total contacts across all complexes (Table 2), demonstrating the conserved peptide centric binding mode utilized by the 1E6 TCR. This fixed anchoring between the 2 molecules was important for stabilization of the TCR-pMHC complex, as — although other peptides without the ‘GDP’ motif were tested and shown to activate the 1E6 T cell clone — we were unable to measure robust affinities using SPR (data not shown). These data support the requirement for a conserved interaction between the 1E6 TCR and the GPD motif, as we observed in our previously published 1E6-A2-ALWGPDPAAA structure. Focused hotspot binding around a conserved GPD motif enables the 1E6 TCR to tolerate peptide degeneracy. Although the 1E6 TCR formed a similar overall interaction with each APL, the stabilization between the TCR and the GPD motif enabled fine differences in the contact network with both the peptide and MHC surface that allowed discrimination between each ligand (Figure 5). For example, the 1E6 TCR made only 47 peptide contacts with A2-MVWGPDPLYV (KD = ~600 μM) compared with 63 and 57 contacts with A2-YQFGPDFPIA (KD = 7.4 μM) and A2-RQFGPDFPTI (KD = 0.5 μM), respectively. Although the number of peptide contacts was a good predictor of TCR binding affinity for some of the APLs, for others, the correlation was poor (Pearson’s correlation = 0.045, P = 0.92), possibly because of different resolutions for each complex structure. For example, the 1E6 TCR made 64 peptide contacts with A2-YLGGPDFPTI (KD = ~400 μM) compared with 43 contacts with A2-RQWGPDPAAV (KD = 7.8 μM). The most important peptide modification in terms of generating new contacts was peptide position 1. The stronger ligands all encoded larger side chains (Arg or Tyr) at peptide position 1 (Figure 5, E–H), enabling interactions with 1E6 that were not present in the weaker APLs that lacked large side chains in this position (Figure 5, A, C, and D). We have previously shown that the 1E6 TCR uses a rigid lock-and-key mechanism during binding to A2-ALWGPDPAAA. These data demonstrated that the unligated structure of the 1E6 TCR was virtually identical to its ligated counterparts. In order to determine whether any of the APLs required an induced fit mechanism during binding that could explain the difference in free binding energy (ΔG) between each complex (Table 2), we solved the unligated structures of all 7 APLs (the A2-ALWGPDPAAA structure has been previously published and was used in this comparison, ref.) (Figure 6 and Supplemental Table 2). The unligated A2-MVWGPDPLYV (KD = ~600 μM) structure revealed that the side chain Tyr9 swung around 8 Å in the complex structure, subsequently making contacts with TCR residues Asp30β and Asn51β (Figure 6A and Figure 5A, respectively). This movement could result in an entropic penalty contributing to the weak TCR binding affinity we observed for this ligand. Additional small movements in the Cα backbone of the peptide around peptide residue Asp6 were apparent in the A2-YLGGPDFPTI (KD = ~400 μM), A2-ALWGPDPAAA (KD = ~208 μM), and A2-RQFGPDWIVA (KD = 44.4 μM) structures (Figure 6, B, C, and E). The unligated structures of A2-AQWGPDAAA, A2-RQWGPDPAAV, A2-YQFGPDFPIA, and A2-RQFGPDFPTI were virtually identical when in complex with 1E6 (Figure 6, D and F–H). Apart from the case of A2-AQWGPDAAA (KD = 61.9 μM), these observations support the conclusion that the higher-affinity ligands required less conformational melding during binding, which could be energetically beneficial (lower entopic cost) during ligation with the 1E6 TCR. Peptide modifications alter the interaction between the 1E6 TCR and the MHC surface. In addition to changes between the TCR and peptide component, we also observed that different APLs had different knock-on effects between the TCR and MHC. MHC residue Arg65 that forms part of the MHC restriction triad (Arg65, Ala69, and Gln155) played a central role in TCR-MHC contacts, with Gln155 playing a less important role and Ala69 playing no role in binding at the interface (Figure 7). Generally, the weaker-affinity APLs made fewer contacts with the MHC surface (27–29 interactions) compared with the stronger-affinity APLs (29–35 contacts), consistent with a better Pearson’s correlation value (0.55) compared with TCR-peptide interactions versus affinity (0.045). For instance, contacts were made between TCR residue Val53β and MHC residue Gln72 in all APLs except for in the weakest affinity ligand pair, 1E6-A2-MVWGPDPLYV, in which a subtle change in TCR conformation — probably mediated by different peptide contacts — abrogated this interaction (Figure 7A). An energetic switch from unfavorable to favorable entropy (order-to-disorder) correlates with antigen potency. Our analysis of the contact network provided some clues that could explain the different antigen potencies and binding affinities between the 1E6 TCR and the different APLs. However, there were clear outliers in which the number of contacts did not match with the strength/potency of the interaction. For example, the 1E6 TCR bound to A2-RQWGPDPAAV with the third strongest affinity (KD = 7.8 μM) but made fewer contacts than with A2-ALWGPDPAAA (KD = ~208 μM) (Table 2). However, it is not necessarily the quantity of contacts that determines the strength of an interaction, but the quality of the contacts. Thus, we performed an in-depth thermodynamic analysis of 6 of the ligands under investigation (Figure 8 and Supplemental Table 3). The weak binding affinity between 1E6 and A2-MVWGPDPLYV and A2-YLGGPDFPTI generated thermodynamic data that were not robust enough to gain insight into the enthalpic (ΔH°) and entropic (TΔS°) changes that contributed to the different binding affinities/potencies for each APL. The overall free binding energies (ΔG°) were between –4.4 and –8.6 kcal/mol, reflecting the wide range of TCR binding affinities we observed for the different APLs. The enthalpic contribution in each complex did not follow a clear trend with affinity, with all but the 1E6-A2-RQFGPDFPTI interaction (ΔH° = 6.3 kcal/mol) generating an energetically favorable enthalpy value (ΔH° = –3.7 to –11.4 kcal/mol); this indicated a net gain in electrostatic interactions during complex formation. However, there was a clear switch in entropy between the weaker-affinity and stronger-affinity ligands, indicated by a strong Pearson’s correlation value between entropy and affinity (Pearson’s correlation value 0.93, P =0.007). For instance, the A2-ALWGPDPAAA, A2-AQWGPDAAA, and A2-RQFGPDWIVA (KD = ~208 μM, KD = 61.9 μM, and KD = 44.4 μM, respectively) were all entropically unfavorable (TΔS° = –2.9 to –5.6 kcal/mol), indicating a net change from disorder to order. Conversely, the stronger-affinity ligands A2-RQWGPDPAAV (KD = 7.8 μM), A2-YQFGPDFPIA (KD = 7.4 μM), and A2-RQFGPDFPTI (KD = 0.5 μM) exhibited favorable entropy (TΔS° = 2.2 to 14.9 kcal/mol), indicating an order-to-disorder change during binding, possibly through the expulsion of ordered water molecules. Furthermore, the structures of the unligated pMHCs demonstrated that, for these stronger-affinity ligands, there was less conformational difference between the TCR ligated pMHCs compared with the weaker-affinity ligands (Figure 6). The potential requirement for a larger degree of induced fit during binding to these weaker-affinity ligands is consistent with the larger entropic penalties observed for these interactions. Potential epitopes for 1E6 TCR occur commonly in the viral proteome. We searched a database of over 1,924,572 unique decamer peptides from the proteome of viral pathogens that are known, or strongly suspected, to infect humans. Three hundred forty-two of these decamers conformed to the motif xxxGPDxxxx. Of these, 53 peptides contained the motif xOxGPDxxxO, where O is one of the hydrophobic amino acid residues A,V, I, L, M, Y, F, and W that might allow binding to HLA-A*0201 (Supplemental Table 4). Thus, there are many pathogen-encoded peptides that could act as agonists for the 1E6 T cell beyond the MVWGPDPLYV and RQFGPDWIVA sequences studied here. Extension of these analyses to include the larger genomes of bacterial pathogens would be expected to considerably increase these numbers. The binding affinity of the 1E6 TCR interaction with A2-RQFGPDWIVA is considerably higher than with the disease-implicated A2-ALWGPDPAAA sequence (KD = 44.4 μM and KD \u003e 200 μM, respectively), highlighting how a pathogen-derived sequence might be capable of priming a 1E6-like T cell. Discussion T cell antigen discrimination is governed by an interaction between the clonally expressed TCR and pMHC, mediated by the chemical characteristics of the interacting molecules. It has recently become clear that TCR cross-reactivity with large numbers of different pMHC ligands is essential to plug holes in T cell immune coverage that pathogens could exploit. Flexibility at the interface between the TCR and pMHC, demonstrated in various studies, has been suggested as a mechanism mediating T cell cross-reactivity with multiple distinct epitopes. This notion is attractive because the CDR loops, which form the TCR antigen-binding site, are usually the most flexible part of the TCR and have the ability to mold around differently shaped ligands. Focused binding around a minimal peptide motif has also been implicated as an alternative mechanism enabling TCR cross-reactivity. Notably among these studies, Garcia and colleagues recently used the alloreactive murine TCR-MHC pair of the 42F3 TCR and H2-Ld to demonstrate recognition of a large number of different peptides via conserved hotspot contacts with prominent up-facing peptide residues. Sethi and colleagues recently demonstrated that the MHCII-restricted Hy.1B11 TCR, which was isolated from a patient with multiple sclerosis, could anchor into a deep pocket formed from peptide residues 2, 3, and 5 (from MBP85–99 bound to HLA-DQ1). This motif was conserved in at least 2 potential foreign peptides, originating from Herpes simplex virus and Pseudomonas aeruginosa, enabling TCR recognition of foreign epitopes. Although these data provided some clues into the molecular mechanism of T cell recognition, there still remain several gaps in our understanding. First, we currently know nothing about how human MHCI–restricted TCRs mediate cross-reactivity in the context of a clinically relevant model of autoimmunity, thought to be a major pathway of disease initiation in several autoimmune diseases. Second, molecular studies have not yet revealed a broad set of rules that determine TCR cross-reactivity because, with the exception of the allo–TCR-MHC pair of the 42F3 TCR and H2-Ld that did not encounter each other during T cell development, studies have been limited to structures of a TCR with only 2 or 3 different ligands. Finally, no studies have included characterization of pathogen-derived ligands recognized by self-reactive T cells with greater potency than the autoantigen, a potentially important facet to break self-tolerance. Here, we investigated a highly cross-reactive MHCI-restricted TCR isolated from a patient with T1D that recognizes an HLA-A*0201–restricted preproinsulin signal peptide (ALWGPDPAAA15–24). Human CD8+ T cell clones expressing TCRs with this specificity mediate the destruction of β cells, have been found in islets early in infection, and are proposed to be a major driver of disease. We solved the structure of the 1E6 TCR with 7 APLs to enable a comprehensive analysis of the molecular basis of TCR degeneracy. The epitopes we selected exhibited a broad range of potencies and could activate the 1E6 T cell clone at exogenously supplied concentrations more than 4 logs apart. Overall, the difference in antigen potency correlated well with the binding energy (ΔG° kcal/mol) of the 1E6 TCR for the different epitopes, which ranged from values of ΔG° = ~–4.4 to –8.6 kcal/mol (calculated from 3D affinity data) or 2D affinity values of AcKa = 2.5 × 10–5 to 4.4 × 10–2 μm4. The weaker end of this spectrum extends our understanding of the limits in which T cells can functionally operate in terms of TCR 3D binding affinity and is in line with the types of very low affinity, yet fully functional self-reactive CD8+ T cells we have observed in tumor-infiltrating lymphocytes. Previous studies of autoreactive TCRs have shown that their binding mode is generally atypical, either due to an unusual binding manner, weak TCR binding affinity, an unstable pMHC, or a combination of these factors. Our data demonstrate the potential for an autoreactive TCR to bind with a conventional binding mode to a stable pMHC with antipathogen-like affinity (KD = 0.5 μM) depending on the peptide sequence. Our structural analysis revealed that the 1E6 TCR bound with a conserved conformation across all APLs investigated. This binding orientation was mediated through a focused interaction with TCR residues Tyr97α and Trp97β that formed an aromatic cap over a central ‘GDP’ motif that was common to all APLs. We have previously demonstrated the importance of the GPD motif using a peptide library scan, as well as a CPL scan approach. Although the 1E6 T cell was able to activate weakly with peptides that lacked this motif, we were unable to robustly measure binding affinities or generate complex structures with these ligands, highlighting the central role of this interaction during 1E6 T cell antigen recognition. This hotspot binding, defined as a localized cluster of interactions that dominate binding energy during protein-protein interactions, has been previously shown to contribute to TCR recognition of MHC as a mechanism that tunes T cell cross-reactivity by providing fixed anchor points that enable TCRs to tolerate a variable peptide cargo. Alternatively, interactions between the TCR and peptide have been shown to dominate the energetic landscape during ligand engagement, ensuring that T cells retain peptide specificity. The binding mechanism utilized by the 1E6 TCR during pMHC recognition is consistent with both of these models. Ligand engagement is dominated by peptide interactions, but hotspot-like interactions with the central GPD motif enable the 1E6 TCR to tolerate peptide residues that vary outside of this region, explaining how T cells expressing this TCR may cross-react with a large number of different peptides. These findings are also analogous to the observed binding mode of the Hy.1B11 TCR, in which one aromatic residue of the TCR CDR3α loop anchored into a pocket created by a conserved peptide motif. In both of these examples, self-recognition is mediated by TCR residues with aromatic side chains. These large, generally hydrophobic amino acids can form strong interactions with other residues through π-π stacking. Combined with evidence demonstrating that aromatic side chains are conserved in the CDR2 loops of TCRs from many species, we speculate that these aromatic residues could impart a level of “stickiness” to TCRs, which might be enriched in an autoimmune setting when the TCR often binds in a nonoptimal fashion. Despite some weak statistical correlation between the surface complementarity (SC) and affinity, closer inspection of the interface revealed no obvious structural signature that could definitively explain the differences in antigen potency and TCR binding strength between the different ligands. However, similar to our findings in other systems, modifications to residues outside of the canonical central peptide bulge were important for generating new interactions. For example, all of the stronger ligands encoded larger side chains (Arg or Tyr) at peptide position 1 that enabled new interactions with 1E6 not present with the Ala at this position in the natural preproinsulin peptide. These data also explain our previous findings that alteration of the anchor residue at peptide position 2 (Leu-Gln) has a direct effect on 1E6 TCR binding affinity because our structural analysis demonstrated that 1E6 made 3 additional bonds with A2-AQWGPDPAAA compared with A2-ALWGPDPAAA, consistent with the \u003e3-fold stronger binding affinity. We have recently demonstrated how a suboptimal position 2 anchor in a melanoma-derived antigen can improve TCR binding through a similar mechanism. These results challenge the notion that the most potent peptide antigens exhibit the greatest pMHC stability and have implications for the design of anchor residue–modified heteroclitic peptides for vaccination. Early thermodynamic analysis of TCR-pMHC interactions suggested a common energetic signature, driven by favorable enthalpy (generally mediated through an increase in electrostatic interactions) and unfavorable entropy (changes from disorder to order). These parameters aligned well with structural data, demonstrating that TCRs engaged pMHC using an induced fit binding mode. However, more recent data have shown that TCRs can utilize a range of energetic strategies during pMHC binding, currently with no obvious pattern in terms of TCR affinity, binding mechanism, or specificity (pathogen, cancer, or self-ligands). Although no energetic signature appears to exist for different TCRs, we used thermodynamic analysis here to explore whether changes in energetics could help explain ligand discrimination by a single TCR. This analysis demonstrated a strong relationship (according to the Pearson’s correlation analysis) between the energetic signature used by the 1E6 TCR and the sensitivity of the 1E6 T cell clone to different APLs. The weaker APL ligands were characterized by favorable enthalpy and unfavorable entropy, whereas the stronger ligands progressively shifted to favorable entropy. These differences were consistent with a greater degree of movement between the unligated and ligated pMHCs for the weaker ligands, suggesting a greater requirement for disorder-to-order changes during TCR binding. Thus, the enhanced antigen potency was probably mediated through a shift from an induced fit to a lock-and-key interaction between the stronger ligands (less requirement for energetically unfavorable disorder-to-order changes), resulting in a more energetically favorable ΔG value. Importantly, the preproinsulin-derived epitope was one of the least potent peptides, demonstrating that the 1E6 T cell clone had the ability to respond to different peptide sequences with far greater potency. The RQFGPDWIVA peptide, which was substantially more potent than the preproinsulin peptide, is within the proteome of a common human pathogen (C. asparagiforme), demonstrating the potential for an encounter between a naive 1E6-like T cell and a foreign peptide with a more potent ligand that might then break self-tolerance. Indeed, we found over 50 decamer peptides from the proteome of likely, or known, human viral pathogens alone that contained both the conserved central GPD motif and anchor residues at positions 2 and 10 that would enable binding to HLA-A*02:01. Further experiments will be required to determine whether any naturally presented, human pathogen–derived peptides act as active ligands for 1E6, but our work presented here demonstrates that it is at least feasible for an autoimmune TCR to bind to a different peptide sequence that could be present in a pathogen proteome with substantially higher affinity and potency than the interaction it might use to attack self-tissue. In summary, this investigation into the molecular basis of T cell cross-reactivity using a clinically relevant cytotoxic CD8+ T cell clone that kills human pancreatic β cells provides answers to a number of previously outstanding questions. First, our data shows that a single TCR has the potential to functionally (assessed through T cell activation) bind to different ligands with affinities ranging across 3 orders of magnitude. Second, this is the first example in which ligands have been identified and characterized for a human autoreactive TCR that are substantially more potent than the natural self-ligand, demonstrating the potential for a pathogenic ligand to break self-tolerance and prime self-reactive T cells. Third, this first structural analysis of a cross-reactive human MHCI–restricted autoimmune TCR showed that degeneracy was mediated through TCR-pMHC anchoring by a conserved minimal binding peptide motif. Finally, TCR ligand discrimination was characterized by an energetic shift from an enthalpically to entropically driven interaction. Our demonstration of the molecular mechanism governing cross-reactivity by this preproinsulin reactive human CD8+ T cell clone supports the notion first put forward by Wucherpfennig and Strominger that molecular mimicry could mediate autoimmunity and has far-reaching implications for the complex nature of T cell antigen discrimination. Methods T cell maintenance and culture. The 1E6 T cell clone was generated as previously described and stored in vapor phase liquid nitrogen in freezing buffer (90% FCS and 10% DMSO). Cells were defrosted rapidly in a 37°C water bath until a small amount of frozen cells were left and then immediately washed in 15–20 ml of R10 media (RPMI 1640 with 10% FCS, 100 IU/ml penicillin, 100 μg/ml streptomycin, 2 mM l-glutamine) by centrifuging at 300 g for 5 minutes. Defrosted cells were cultured in T cell media: R10 with 1× nonessential amino acids, 1 mM sodium pyruvate, 10 mM HEPES buffer (all from Invitrogen), 20 IU/ml of IL-2 (aldesleukin, brand name Proleukin, Prometheus) and 25 ng/ml IL-15 (PeproTech), for 24 hours; then, 0.75 × 106 to 1.5 × 106 cells expanded by coculture with 15 × 106 irradiated (3,100 cGy) PBMCs from 3 donors in a 25 cm2 tissue culture flask with 1 μg/ml of phytoheamagglutinin (Alere Inc.) and T cell media as above. The clone was transferred to 24-well tissue culture plates (3 × 106 to 4 × 106 per well in 2 ml) at day 7, and the IL-2 increased to 200 IU/ml. For the purpose of this study, the clone was passaged 3 times and used between weeks 2 and 4 after expansion. T cell activation assays and tetramer staining. The [51Cr] release cytotoxicity assay was performed as previously described. Target A2 CIR cells were labeled for 1 hour at 37°C with 30 μCi chromium (sodium chromate in normal saline, PerkinElmer) per 1 × 106 cells, washed with R10, and allowed to leach for a further hour at 37°C in R10 to remove any excess chromium from the cells. After chromium labeling, target cells were washed and plated at 1,000 cells/well in 96-well tissue culture plates and pulsed with peptide at the indicated concentrations for 2 hours at 37°C. T cells were added to give the desired T cell/target cell (5:1) ratio and a final volume of 150 μl R10. Target cells were also incubated alone or with 1% Triton X-100 detergent (Sigma-Aldrich) to give the spontaneous and total chromium released from the target cells, respectively. After overnight incubation, at 37°C and 5% CO2, the supernatants were both (i) assayed for MIP1β by ELISA (R\u0026D Systems) and (ii) harvested (10% of total volume), mixed with 150 μl Optipahse supermix scintillation mixture (PerkinElmer) in 96-well polyethylene terephthalate plates (PerkinElmer), and sealed; the amount of released chromium was measured indirectly on a 1450 Microbeta counter (PerkinElmer). The percentage of specific target cell lysis by T cells was calculated according to the following formula: (experimental release [with T cells and target cells] − spontaneous release from target cells)/(total release from target cells − spontaneous release from target cells) × 100. Experiments were independently completed in triplicate. Tetrameric pMHCI reagents (tetramers) were constructed by the addition of PE-conjugated streptavidin (Invitrogen) at a pMHCI/streptavidin molar ratio of 4:1. A total of 50,000 T cells were stained with PE-conjugated tetramer (25 μg/ml) folded around the indicated peptides for 30 minutes on ice and washed with PBS before staining with 2 μl (1:40 dilution of the DMSO stock in PBS) of the violet LIVE/DEAD fixable dead cell stain Vivid (Invitrogen) for 5 minutes at room temperature before direct addition of 2 μl of anti–CD8-APC antibody (clone BW135/80, Miltenyi Biotec) and incubated for a further 20 minutes on ice before being washed in FACS buffer (2% FCS in PBS). Data were acquired using a FACSCanto II flow cytometer (BD Biosciences) and analyzed with FlowJo software (Tree Star Inc.). Protein expression, refolding, and purification. The 1E6 TCR, HLA-A*0201, and human β2m chain were generated as described previously. The 1E6 TCR and HLA-A*0201 peptide variants were refolded and purified as described previously. Biotinylated pMHCI and pMHC tetramer were prepared as previously described. pMHC stability assays. Thermal stability of the HLA-A*0201–peptide complexes was assessed by circular dichroism (CD) spectroscopy monitoring the change in ellipticities at 218 nm. Data were collected, in duplicate, using a nitrogen-flushed Module B end-station spectrophotometer at the B23 Synchrotron Radiation CD Beamline at the Diamond Light Source (DLS). Samples were prepared in phosphate buffered saline, pH 7.4, and concentrated to ~10 mM. Spectra were measured every 5°C over a temperature range between 5°C and 90°C with 5 minutes of equilibration time for each temperature. Four scans were acquired using an integration time of 1 second, a path length of 0.02 cm, and a slit width of 0.5 mm equivalent to a 1.2-nm bandwidth. Reversibility was monitored by measuring the spectrum at 20°C after cooling from 90°C with 30 minutes of incubation. Melting curves were analyzed assuming a 2-state trimer-to-monomer transition from the native (N) to unfolded (U) conformation N3 ↔ 3U with an equilibrium constant K = (U)3/(N3) = F/(3c2[1-F]3), where F and c are the degree of folding and protein concentration, respectively. Data were fitted as described. Fitted parameters were the melting temperature Tm, van’t Hoff’s enthalpy ΔHvH, and the slope and intercept of the native baseline. As all protein complexes aggregated to various degrees upon unfolding, the ellipticity of the unfolded state was set as a constant of –4,500 deg cm2/dmol. SPR analysis. Binding analysis was performed using a BIAcore T200 equipped with a CM5 sensor chip as previously described. Binding analysis was performed 3× in independent experiments using pMHC monomers generated in-house. Approximately 200–500 RU of each HLA-A*0201–peptide complex was attached to the CM5 sensor chip at a slow flow rate of 10 μl/min to ensure uniform distribution on the chip surface. The 1E6 TCR was purified and concentrated to approximately 40–350 μM on the same day of SPR analysis to reduce the likelihood of TCR aggregation. For equilibrium analysis, 10 serial dilutions were prepared in triplicate for each sample and injected over the relevant sensor chips at 25°C. TCR was injected over the chip surface using kinetic injections at a flow rate of 45 μl/min using HLA-A*0201–ELAGIGILTV as a negative control surface on flow cell 1. For the thermodynamics experiments, this method was repeated at the following temperatures: 5°C, 13°C, 18°C, 25°C, 30°C, and 37°C. Results were analyzed using BIAevaluation 3.1, Excel, and Origin 6.0 software. The KD values were calculated assuming a 1:1 interaction by plotting specific equilibrium-binding responses against protein concentrations, followed by nonlinear least squares fitting of the Langmuir binding equation. The thermodynamic parameters were calculated using the nonlinear van’t Hoff equation (RT ln KD = ΔH° –TΔS° + ΔCp°[T-T0] – TΔCp° ln [T/T0]) with T0=298 K. Adhesion frequency assay. We used an adhesion frequency assay to measure the 2D affinity of TCR-pMHC interactions at the cell membrane as previously described. Briefly, human 1E6 T cells were mounted onto 1 micropipette, and, on the other pipette, human rbcs coated with pMHC by biotin-streptavidin coupling served as both a surrogate APC and an adhesion sensor for detecting the TCR-pMHC interaction. Site densities of TCR and pMHC were measured by flow cytometry as previously described. All assays were performed using at least 5 cell pairs and calculated as an average of 100 cell-cell contacts. Crystal structure determination. All protein crystals were grown at 18°C by vapor diffusion via the sitting drop technique. Each pMHCI (200 nl, 10 mg/ml) in crystallization buffer (10 mM TRIS [pH 8.1] and 10 mM NaCl) was added to 200 nl of reservoir solution. HLA-A*0201–MVWGPDPLYV (A2-MVW) crystals were grown in 0.2 M ammonium chloride, 0.1 M TRIS (pH 8), 20% PEG 6000; HLA-A*0201–YLGGPDFPTI (A2-YLG) crystals were grown in 0.2 M sodium nitrate, 0.1 M BIS TRIS propane (pH 6.5), 20% PEG 3350; HLA-A*0201–AQWGPDPAAA (A2-AQW) crystals were grown in 0.2 M sodium malonate, 0.1 M BIS TRIS propane (pH 6.5), 20% PEG3350; HLA-A*0201–RQFGPDWIVA (A2-RQF[A]) crystals were grown in 0.2 M sodium sulphate, 0.1 M BIS TRIS propane (pH 6.5), 20% PEG 3350; HLA-A*0201–RQWGPDPAAV (A2-RQW) crystals were grown in 0.1 M TRIS (pH 8), 20% PEG 8000, 15% glycerol; HLA-A*0201–YQFGPDFPTA (A2-YQF) crystals were grown in 0.1 M TRIS (pH 8), 25% PEG 4000, 15% glycerol; HLA-A*0201–RQFGPDFPTI (A2-RQF[I]) crystals were grown in 0.2 M potassium/sodium tartrate, 0.1 M BIS TRIS propane (pH 8.5), 20% PEG 3350; 1E6-A2-MVW crystals were grown in 0.1 M HEPES (pH 7.5), 15% PEG 4000, 0.2 M sodium acetate; 1E6-A2-YLG crystals were grown in 0.1 M sodium cacodylate (pH 6.5), 15% PEG 4000, 0.2 M sodium acetate; 1E6-A2-AQW crystals were grown in 0.2 M sodium citrate, 0.1 M BIS TRIS propane (pH 6.5), 20% PEG 3350; 1E6-A2-RQF(A) crystals were grown in 0.1 M HEPES (pH 7), 15% PEG 4000, 0.2 M sodium acetate; 1E6-A2-RQW crystals were grown in 0.2 M sodium cholride, 0.1 M MES (pH 6), 20% PEG 6000; 1E6-A2-YQF crystals were grown in 0.2 M sodium cholride, 0.1 M HEPES (pH 7), 20% PEG 3350; and 1E6-A2-RQF(I) crystals were grown in 0.1 M HEPES (pH 7.5), 15% PEG 4000, 0.2 M sodium acetate. Crystallization screens were conducted using an Art-Robbins Phoenix dispensing robot (Alpha Biotech Ltd.), and data were collected at 100 K at the DLS at a wavelength of 0.98 Å using an ADSC Q315 CCD detector. Reflection intensities were estimated using XIA2, and the data were analyzed with Scala and the CCP4 package. Structures were solved with molecular replacement using Phaser. Sequences were adjusted with Coot, and the models were refined with REFMAC5. Graphical representations were prepared with PyMOL. The reflection data and final model coordinates were deposited with the PDB database (A2-MVW PDB: 5C0H; A2-YLG PDB: 5C0G; A2-AQW PDB: 5C0D; A2-RQF[A] PDB: 5C0J; A2-RQW PDB: 5C0F; A2-YQF PDB: 5C0E; A2-RQF[I] PDB: 5C0I; 1E6-A2-MVW PDB: 5C0A; 1E6-A2-YLG PDB: 5C09; 1E6-A2-AQW PDB: 5HYJ; 1E6-A2-RQF[A] PDB: 5C0C; 1E6-A2-RQW PDB: 5C08; 1E6-A2-YQF PDB: 5C07; and 1E6-A2-RQF[I] PDB: 5C0B). Peptide motif predictions. Peptide motif predictions were performed by searching a viral database compiled using publicly available protein sequences of over 1,924,572 unique decamer peptides from the proteome of viral pathogens. The motif xOxGPDxxxO — where O is anyone of the hydrophobic amino acid residues A,V, I, L, M, Y, F, and W that might allow binding to HLA-A*0201 — was used as the search parameter. Statistics. Pearson’s correlation analysis was performed to determine the relationship between TCR binding affinity and antigen potency, structural correlates, or thermodynamics using Origin Lab 9.0 pro. Author contributions AMB, GD, AJS, BS, WR, AT, PJ, AF, AS, JJM, LW, PJR, and DKC performed experiments and analyzed the data. AKS, JR, CZ, JJM, MP, and DKC wrote the manuscript. AKS and DKC conceived and directed the study. AKS and DKC funded the study. All authors contributed to discussions. Supplementary Material Conflict of interest: The authors have declared that no conflict of interest exists. License: This work is licensed under the Creative Commons Attribution 4.0 International License. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/. Reference information:J Clin Invest. 2016;126(6):2191–2204. doi:10.1172/JCI85679. See the related Commentary beginning on page . Deconstructing the peptide-MHC specificity of T cell recognition Specificity and degeneracy of T cells A single autoimmune T cell receptor recognizes more than a million different peptides A very high level of crossreactivity is an essential feature of the T-cell receptor Why must T cells be cross-reactive? A direct estimate of the human alphabeta T cell receptor diversity Infection as a cause of type 1 diabetes? Motifs for a deadly encounter Molecular mimicry in T cell-mediated autoimmunity: viral peptides activate human T cell clones specific for myelin basic protein T cell-mediated autoimmune disease due to low-affinity crossreactivity to common microbial peptides CDR3 loop flexibility contributes to the degeneracy of TCR recognition Conformational changes and flexibility in T-cell receptor recognition of peptide-MHC complexes T cell receptor cross-reactivity directed by antigen-dependent tuning of peptide-MHC molecular flexibility T cell allorecognition via molecular mimicry T cell receptor recognition of a ‘super-bulged’ major histocompatibility complex class I-bound peptide A T cell receptor flattens a bulged antigenic peptide presented by a major histocompatibility complex class I molecule T cell receptor signaling is limited by docking geometry to peptide-major histocompatibility complex Crystal structure of a T cell receptor bound to an allogeneic MHC molecule Crossreactivity of a human autoimmune TCR is dominated by a single TCR loop Minimal conformational plasticity enables TCR cross-reactivity to different MHC class II heterodimers How a single T cell receptor recognizes both self and foreign MHC Structural interplay between germline interactions and adaptive recognition determines the bandwidth of TCR-peptide-MHC cross-reactivity Structural basis for the killing of human β cells by CD8(+) T cells in type 1 diabetes Human beta-cell killing by autoreactive preproinsulin-specific CD8 T cells is predominantly granule-mediated with the potency dependent upon T-cell receptor avidity CTLs are targeted to kill β cells in patients with type 1 diabetes through recognition of a glucose-regulated preproinsulin epitope Demonstration of islet-autoreactive CD8 T cells in insulitic lesions from recent onset and long-term type 1 diabetes patients A molecular switch in immunodominant HIV-1-specific CD8 T-cell epitopes shapes differential HLA-restricted escape Structural and biophysical determinants of αβ T-cell antigen recognition T cell antigen receptor recognition of antigen-presenting molecules Measuring two-dimensional receptor-ligand binding kinetics by micropipette The kinetics of two-dimensional TCR and pMHC interactions determine T-cell responsiveness Rfree and the rfree ratio Hard wiring of T cell receptor specificity for the major histocompatibility complex is underpinned by TCR adaptability Alloreactivity between disparate cognate and allogeneic pMHC-I complexes is the result of highly focused, peptide-dependent structural mimicry Microorganisms and autoimmunity: making the barren field fertile? TCRs used in cancer gene therapy cross-react with MART-1/Melan-A tumor antigens via distinct mechanisms Four A6-TCR/peptide/HLA-A2 structures that generate very different T cell signals are nearly identical Crossreactive T Cells spotlight the germline rules for alphabeta T cell-receptor interactions with MHC molecules Structural insights into the editing of germ-line-encoded interactions between T-cell receptor and MHC class II by Valpha CDR3 A structural basis for varied αβ TCR usage against an immunodominant EBV antigen restricted to a HLA-B8 molecule Comparison of peptide-major histocompatibility complex tetramers and dextramers for the identification of antigen-specific T cells More tricks with tetramers: a practical guide to staining T cells with peptide-MHC multimers Antibody stabilization of peptide-MHC multimers reveals functional T cells bearing extremely low-affinity TCRs Unconventional topology of self peptide-major histocompatibility complex binding by a human autoimmune T cell receptor Structure of a human autoimmune TCR bound to a myelin basic protein self-peptide and a multiple sclerosis-associated MHC class II molecule A highly tilted binding mode by a self-reactive T cell receptor results in altered engagement of peptide and MHC T cell receptor reversed polarity recognition of a self-antigen major histocompatibility complex Structure of a TCR with high affinity for self-antigen reveals basis for escape from negative selection Distortion of the MHC class I binding groove to accommodate an insulin-derived 10-mer peptide A hot spot of binding energy in a hormone-receptor interface Identification of a crucial energetic footprint on the alpha1 helix of human histocompatibility leukocyte antigen (HLA)-A2 that provides functional interactions for recognition by tax peptide/HLA-A2-specific T cell receptors Interface-disrupting amino acids establish specificity between T cell receptors and complexes of major histocompatibility complex and peptide Two-step binding mechanism for T-cell receptor recognition of peptide MHC The CDR3 regions of an immunodominant T cell receptor dictate the ‘energetic landscape’ of peptide-MHC recognition TCR-peptide specificity overrides affinity enhancing TCR-MHC interactions Evolutionarily conserved features contribute to alphabeta T cell receptor specificity Germ line-governed recognition of a cancer epitope by an immunodominant human T-cell receptor Structural basis for ineffective T-cell responses to MHC anchor residue-improved “heteroclitic” peptides T-cell receptor-optimized peptide skewing of the T-cell repertoire can enhance antigen targeting Modification of MHC anchor residues generates heteroclitic peptides that alter TCR binding and T cell recognition Thermodynamics of T cell receptor binding to peptide-MHC: evidence for a general mechanism of molecular scanning TCR binding to peptide-MHC stabilizes a flexible recognition interface Structural basis of T cell recognition Two different T cell receptors use different thermodynamic strategies to recognize the same peptide/MHC ligand Thermodynamics of T-cell receptor-peptide/MHC interactions: progress and opportunities The structural dynamics and energetics of an immunodominant T cell receptor are programmed by its Vbeta domain The energetic basis underpinning T-cell receptor recognition of a super-bulged peptide bound to a major histocompatibility complex class I molecule Interaction between the CD8 coreceptor and major histocompatibility complex class I stabilizes T cell receptor-antigen complexes at the cell surface Measuring circular dichroism in a capillary cell using the b23 synchrotron radiation CD beamline at diamond light source Analysis of circular dichroism data How to measure and predict the molar absorption coefficient of a protein Circular dichroic analysis of denatured proteins: inclusion of denatured proteins in the reference set T cell receptor engagement of peptide-major histocompatibility complex class I does not modify CD8 binding Kinetics of MHC-CD8 interaction at the T cell membrane Decision making in xia2 Collaborative Computational Project, number 4: providing programs for protein crystallography Phaser crystallographic software Coot: model-building tools for molecular graphics  Identification of human viral protein-derived ligands recognized by individual major histocompatibility complex class I (MHCI)-restricted T-cell receptors The 1E6 T cell clone reacts with a broad sensitivity range to APLs. (A and B) The 1E6 T cell clone was tested in a peptide dilution assay, in triplicate, with MVWGPDPLYV (gray), YLGGPDFPTI (red), ALWGPDPAAA (blue), AQWGPDPAAA (green), RQFGPDWIVA (dark blue), RQWGPDPAAV (purple), YQFGPDFPTA (yellow), and RQFGPDFPTI (cyan) peptides presented by HLA-A*0201–expressing C1R cells for release of MIP-1β (A) and killing (B). (C) The 1E6 T cell clone was stained, in duplicate, with tetramers composed of each APL (colored as above) presented by HLA-A*0201. (D) The stability of each APL (colored as above) was tested, in duplicate, using CD by recording the peak at 218 nm absorbance from 5°C–90°C. Tm values were calculated using a Boltzmann fit to each set of data. 3D and 2D binding analysis of the 1E6 TCR with A2-ALW and the APLs. (A–H) Binding affinity of the 1E6 TCR interaction at 25°C using SPR. Eight serial dilutions of the 1E6 TCR were measured (shown in the inset); representative data from 3 independent experiments are plotted. The equilibrium binding constant (KD) values were calculated using a nonlinear curve fit (y= [P1x]/[P2 + X]). In order to calculate each response, the 1E6 TCR was also injected over a control sample (HLA-A*0201–ILAKFLHWL) that was deducted from the experimental data. (A) 1E6-A2-MVWGPDPLYV (approximate value); (B) 1E6-A2-YLGGPDFPTI (approximate value); (C) 1E6-A2-ALWGPDPAAA; (D) 1E6-A2-AQWGPDPAAA; (E) 1E6-A2-RQFGPDWIVA; (F) 1E6-A2-RQWGPDPAAV; (G) 1E6-A2-YQFGPDFPTA; and (H) 1E6-A2-RQFGPDFPTI. (I) ΔG values, calculated from SPR experiments, plotted against 1/EC50 (the reciprocal peptide concentration required to reach half-maximal 1E6 T cell killing) showing Pearson’s coefficient analysis (r) and P value (including approximate values from A and B). (J) Effective 2D affinity (AcKa) calculated using adhesion frequency assays, using at least 5 cell pairs, and calculated as an average of 100 cell cell contacts. (K) Effective 2D affinity plotted against 1/EC50 showing Pearson’s coefficient analysis (r) and P value. The 1E6 TCR uses a conserved binding mode to engage A2-ALWGPDPAAA and the APLs. (A) Superposition of the 1E6 TCR (multicolored illustration) in complex with all 7 APLs (multicolored sticks) and the A2-ALWGPDPAAA ligand using the HLA-A*0201 (gray illustration) molecule to align all of the structures. The 1E6 TCR and each peptide are colored according to the APL used in the complex as in Figure 1. (B) Position of the 1E6 TCR CDR loops (multicolored lines) in each complex. The ALWGPDPAAA peptide (green sticks) is shown in the HLA-A*0201 binding groove (gray surface). (C) The Cα backbone conformation of each APL (multicolored illustration) in the context of the HLA-A*0201 α1 helices (gray illustration). (D) Crossing angle of the 1E6 TCR (multicolored lines) calculated using previously published parameters in the context of the ALWGPDPAAA peptide (green sticks) bound in the HLA-A*0201 binding groove (gray surface). A conserved interaction with a GPD motif underpins the 1E6 TCR interaction with the APLs. Interaction between 1E6 TCR (gray illustration) residues Tyr97α and Tyr97β (the position of these side chains in the TCR in complex with all 7 APLs, and the previously reported A2-ALWGPDPAAA epitope, is shown in multicolored sticks; ref.) and the GPD peptide motif (the position of these side chains in all 7 APLs and A2-ALWGPDPAAA in complex with the 1E6 TCR is shown in multicolored sticks). The rest of the peptide, and the MHCα1 helix, are shown as a gray illustration. The 1E6 TCR makes distinct peptide contacts with peripheral APL residues. Interactions between the 1E6 TCR and peptide residues outside of the conserved GPD motif. The MHCα1 helix is shown in gray illustrations. Hydrogen bonds are shown as red dotted lines; van der Waals (vdW) contacts are shown as black dotted lines. Boxes show total contacts between the 1E6 TCR and each peptide ligand. (A) Interaction between the 1E6 TCR (black illustration and sticks) and A2-MVWGPDPLYV (black illustration and sticks). (B) Interaction between the 1E6 TCR (red illustration and sticks) and A2-YLGGPDFPTI (red illustration and sticks). (C) Interaction between the 1E6 TCR (blue illustration and sticks) and A2-ALWGPDPAAA (blue illustration and sticks) reproduced from previous published data. (D) Interaction between the 1E6 TCR (green illustration and sticks) and A2-AQWGPDPAAA (green illustration and sticks). (E) Interaction between the 1E6 TCR (dark blue illustration and sticks) and A2-RQFGPDWIVA (dark blue illustration and sticks). (F) Interaction between the 1E6 TCR (purple illustration and sticks) and A2-RQWGPDPAAV (purple illustration and sticks). (G) Interaction between the 1E6 TCR (yellow illustration and sticks) and A2-YQFGPDFPTA (yellow illustration and sticks). (H) Interaction between the 1E6 TCR (cyan illustration and sticks) and A2-RQFGPDFPTI (cyan illustration and sticks). Comparison of ligated and unligated APLs. Superposition of each APL in unligated form and ligated to the 1E6 TCR. All unligated pMHCs are shown as light green illustrations. Peptide sequences are shown underneath each structure aligned with the peptide structure. Black arrows denote the direction of the side chain. Upward arrows indicates solvent exposed, downward arrows indicates anchor position, and no arrow indicates an intermediate position. (A) A2-MVWGPDPLYV (black sticks). A large conformational shift was observed for Tyr8 in the ligated versus unligated states (black circle). (B) A2-YLGGPDFPTI (red sticks). (C) A2-ALWGPDPAAA (blue sticks) reproduced from previous published data. (D) A2-AQWGPDPAAA (green sticks). (E) A2-RQFGPDWIVA (dark blue sticks). (F) A2-RQWGPDPAAV (purple sticks). (G) A2-YQFGPDFPTA (yellow sticks). (H) A2-RQFGPDFPTI (cyan sticks). The 1E6 TCR makes distinct peptide contacts with the MHC surface depending on the peptide cargo. Interactions between the 1E6 TCR and the MHC α1 helix residues Arg65, Lys66, and Gln72. Hydrogen bonds are shown as red dotted lines; vdW contacts are shown as black dotted lines. MHCα1 helix are shown in gray illustrations. Boxes show total contacts between the 1E6 TCR and these key residues (green boxes are MHC residues; white boxes are TCR residues). (A) Interaction between the 1E6 TCR (black illustration and sticks) and A2-MVWGPDPLYV (black illustration and sticks). (B) Interaction between the 1E6 TCR (red illustration and sticks) and A2-YLGGPDFPTI (red illustration and sticks). (C) Interaction between the 1E6 TCR (blue illustration and sticks) and A2-ALWGPDPAAA (blue illustration and sticks) reproduced from previous published data. (D) Interaction between the 1E6 TCR (green illustration and sticks) and A2-AQWGPDPAAA (green illustration and sticks). (E) Interaction between the 1E6 TCR (dark blue illustration and sticks) and A2-RQFGPDWIVA (dark blue illustration and sticks). (F) Interaction between the 1E6 TCR (purple illustration and sticks) and A2-RQWGPDPAAV (purple illustration and sticks). (G) Interaction between the 1E6 TCR (yellow illustration and sticks) and A2-YQFGPDFPTA (yellow illustration and sticks). (H) Interaction between the 1E6 TCR (cyan illustration and sticks) and A2-RQFGPDFPTI (cyan illustration and sticks). Thermodynamic analysis of the 1E6 TCR with A2-ALWGPDPAAA and the APLs. Eight serial dilutions of the 1E6 TCR were injected, in duplicate, over each immobilized APL and A2-ALW at 5°C, 13°C, 18°C, 25°C, 30°C, and 37°C. The equilibrium binding constant (KD) values were calculated using a nonlinear curve fit (y = [P1x]/[P2 + X]), and thermodynamic parameters were calculated according to the Gibbs-Helmholtz equation (ΔG° = ΔH − TΔS°). The binding free energies, ΔG° (ΔG° = RTlnKD), were plotted against temperature (K) using nonlinear regression to fit the 3-parameters van’t Hoff equation (RT ln KD = ΔH° – TΔS° + ΔCp°[T-T0] – TΔCp° ln [T/T0] with T0 = 298 K). (A) 1E6-A2-ALWGPDPAAA; (B) 1E6-A2-AQWGPDPAAA; (C) 1E6-A2-RQFGPDWIVA; (D) 1E6-A2-RQWGPDPAAV, (E) 1E6-A2-YQFGPDFPTA; and (F) 1E6-A2-RQFGPDFPTI. 1E6 TCR-pMHC contacts, affinity measurements and thermodynamics Peptides used in this 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diff --git a/annotated_BioC_JSON/PMC4887326_ann.json b/annotated_BioC_JSON/PMC4887326_ann.json
new file mode 100644
index 0000000000000000000000000000000000000000..d914a87dbf96e06223b07ab1b685901788d15009
--- /dev/null
+++ b/annotated_BioC_JSON/PMC4887326_ann.json
@@ -0,0 +1 @@
+[{"sourceid":"4887326","sourcedb":"","project":"","target":"","text":"Structural insights into the regulatory mechanism of the Pseudomonas aeruginosa YfiBNR system YfiBNR is a recently identified bis-(3’-5’)-cyclic dimeric GMP (c-di-GMP) signaling system in opportunistic pathogens. It is a key regulator of biofilm formation, which is correlated with prolonged persistence of infection and antibiotic drug resistance. In response to cell stress, YfiB in the outer membrane can sequester the periplasmic protein YfiR, releasing its inhibition of YfiN on the inner membrane and thus provoking the diguanylate cyclase activity of YfiN to induce c-di-GMP production. However, the detailed regulatory mechanism remains elusive. Here, we report the crystal structures of YfiB alone and of an active mutant YfiBL43P complexed with YfiR with 2:2 stoichiometry. Structural analyses revealed that in contrast to the compact conformation of the dimeric YfiB alone, YfiBL43P adopts a stretched conformation allowing activated YfiB to penetrate the peptidoglycan (PG) layer and access YfiR. YfiBL43P shows a more compact PG-binding pocket and much higher PG binding affinity than wild-type YfiB, suggesting a tight correlation between PG binding and YfiB activation. In addition, our crystallographic analyses revealed that YfiR binds Vitamin B6 (VB6) or L-Trp at a YfiB-binding site and that both VB6 and L-Trp are able to reduce YfiBL43P-induced biofilm formation. Based on the structural and biochemical data, we propose an updated regulatory model of the YfiBNR system. INTRODUCTION Bis-(3’-5’)-cyclic dimeric GMP (c-di-GMP) is a ubiquitous second messenger that bacteria use to facilitate behavioral adaptations to their ever-changing environment. An increase in c-di-GMP promotes biofilm formation, and a decrease results in biofilm degradation (Boehm et al.,; Duerig et al.,; Hickman et al.,; Jenal,; Romling et al.,). The c-di-GMP level is regulated by two reciprocal enzyme systems, namely, diguanylate cyclases (DGCs) that synthesize c-di-GMP and phosphodiesterases (PDEs) that hydrolyze c-di-GMP (Kulasakara et al.,; Ross et al.,; Ross et al.,). Many of these enzymes are multiple-domain proteins containing a variable N-terminal domain that commonly acts as a signal sensor or transduction module, followed by the relatively conserved GGDEF motif in DGCs or EAL/HD-GYP domains in PDEs (Hengge,; Navarro et al.,; Schirmer and Jenal,). Intriguingly, studies in diverse species have revealed that a single bacterium can have dozens of DGCs and PDEs (Hickman et al.,; Kirillina et al.,; Kulasakara et al.,; Tamayo et al.,). In Pseudomonas aeruginosa in particular, 42 genes containing putative DGCs and/or PDEs were identified (Kulasakara et al.,). The functional role of a number of downstream effectors of c-di-GMP has been characterized as affecting exopolysaccharide (EPS) production, transcription, motility, and surface attachment (Caly et al.,; Camilli and Bassler,; Ha and O’Toole,; Pesavento and Hengge,). However, due to the intricacy of c-di-GMP signaling networks and the diversity of experimental cues, the detailed mechanisms by which these signaling pathways specifically sense and integrate different inputs remain largely elusive. Biofilm formation protects pathogenic bacteria from antibiotic treatment, and c-di-GMP-regulated biofilm formation has been extensively studied in P. aeruginosa (Evans,; Kirisits et al.,; Malone,; Reinhardt et al.,). In the lungs of cystic fibrosis (CF) patients, adherent biofilm formation and the appearance of small colony variant (SCV) morphologies of P. aeruginosa correlate with prolonged persistence of infection and poor lung function (Govan and Deretic,; Haussler et al.,; Haussler et al.,; Parsek and Singh,; Smith et al.,). Recently, Malone and coworkers identified the tripartite c-di-GMP signaling module system YfiBNR (also known as AwsXRO (Beaumont et al.,; Giddens et al.,) or Tbp (Ueda and Wood,)) by genetic screening for mutants that displayed SCV phenotypes in P. aeruginosa PAO1 (Malone et al.,; Malone et al.,). The YfiBNR system contains three protein members and modulates intracellular c-di-GMP levels in response to signals received in the periplasm (Malone et al.,). More recently, this system was also reported in other Gram-negative bacteria, such as Escherichia coli (Hufnagel et al.,; Raterman et al.,; Sanchez-Torres et al.,), Klebsiella pneumonia (Huertas et al.,) and Yersinia pestis (Ren et al.,). YfiN is an integral inner-membrane protein with two potential transmembrane helices, a periplasmic Per-Arnt-Sim (PAS) domain, and cytosolic domains containing a HAMP domain (mediate input-output signaling in histidine kinases, adenylyl cyclases, methyl-accepting chemotaxis proteins, and phosphatases) and a C-terminal GGDEF domain indicating a DGC’s function (Giardina et al.,; Malone et al.,). YfiN is repressed by specific interaction between its periplasmic PAS domain and the periplasmic protein YfiR (Malone et al.,). YfiB is an OmpA/Pal-like outer-membrane lipoprotein (Parsons et al.,) that can activate YfiN by sequestering YfiR (Malone et al.,) in an unknown manner. Whether YfiB directly recruits YfiR or recruits YfiR via a third partner is an open question. After the sequestration of YfiR by YfiB, the c-di-GMP produced by activated YfiN increases the biosynthesis of the Pel and Psl EPSs, resulting in the appearance of the SCV phenotype, which indicates enhanced biofilm formation (Malone et al.,). It has been reported that the activation of YfiN may be induced by redox-driven misfolding of YfiR (Giardina et al.,; Malone et al.,; Malone et al.,). It is also proposed that the sequestration of YfiR by YfiB can be induced by certain YfiB-mediated cell wall stress, and mutagenesis studies revealed a number of activation residues of YfiB that were located in close proximity to the predicted first helix of the periplasmic domain (Malone et al.,). In addition, quorum sensing-related dephosphorylation of the PAS domain of YfiN may also be involved in the regulation (Ueda and Wood,; Xu et al.,). Recently, we solved the crystal structure of YfiR in both the non-oxidized and the oxidized states, revealing breakage/formation of one disulfide bond (Cys71-Cys110) and local conformational change around the other one (Cys145-Cys152), indicating that Cys145-Cys152 plays an important role in maintaining the correct folding of YfiR (Yang et al.,). In the present study, we solved the crystal structures of an N-terminal truncated form of YfiB (34–168) and YfiR in complex with an active mutant YfiBL43P. Most recently, Li and coworkers reported the crystal structures of YfiB (27–168) alone and YfiRC71S in complex with YfiB (59–168) (Li et al.,). Compared with the reported complex structure, YfiBL43P in our YfiB-YfiR complex structure has additional visible N-terminal residues 44–58 that are shown to play essential roles in YfiB activation and biofilm formation. Therefore, we are able to visualize the detailed allosteric arrangement of the N-terminal structure of YfiB and its important role in YfiB-YfiR interaction. In addition, we found that the YfiBL43P shows a much higher PG-binding affinity than wild-type YfiB, most likely due to its more compact PG-binding pocket. Moreover, we found that Vitamin B6 (VB6) or L-Trp can bind YfiR with an affinity in the ten millimolar range. Together with functional data, these results provide new mechanistic insights into how activated YfiB sequesters YfiR and releases the suppression of YfiN. These findings may facilitate the development and optimization of anti-biofilm drugs for the treatment of chronic infections. RESULTS Overall structure of YfiB We obtained two crystal forms of YfiB (residues 34–168, lacking the signal peptide from residues 1–26 and periplasmic residues 27–33), crystal forms I and II, belonging to space groups P21 and P41, respectively. Overall structure of YfiB. (A) The overall structure of the YfiB monomer. (B) A topology diagram of the YfiB monomer. (C and D) The analytical ultracentrifugation experiment results for the wild-type YfiB and YfiBL43P\n Two dimeric types of YfiB dimer. (A–C) The “head to head” dimer. (D–F) The “back to back” dimer. (A) and (E) indicate the front views of the two dimers, (B) and (F) indicate the top views of the two dimers, and (C) and (D) indicate the details of the two dimeric interfaces The crystal structure of YfiB monomer consists of a five-stranded β-sheet (β1-2-5-3-4) flanked by five α-helices (α1–5) on one side. In addition, there is a short helix turn connecting the β4 strand and α4 helix (Fig. 1A and 1B). Each crystal form contains three different dimeric types of YfiB, two of which are present in both, suggesting that the rest of the dimeric types may result from crystal packing. Here, we refer to the two dimeric types as “head to head” and “back to back” according to the interacting mode (Fig. 2A and 2E), with the total buried surface areas being 316.8 Å2 and 554.3 Å2, respectively. The “head to head” dimer exhibits a clamp shape. The dimerization occurs mainly via hydrophobic interactions formed by A37 and I40 on the α1 helices, L50 on the β1 strands, and W55 on the β2 strands of both molecules, making a hydrophobic interacting core (Fig. 2A–C). The “back to back” dimer presents a Y shape. The dimeric interaction is mainly hydrophilic, occurring among the main-chain and side-chain atoms of N68, L69, D70 and R71 on the α2-α3 loops and R116 and S120 on the α4 helices of both molecules, resulting in a complex hydrogen bond network (Fig. 2D–F). The YfiB-YfiR interaction Overall structure of the YfiB-YfiR complex and the conserved surface in YfiR. (A) The overall structure of the YfiB-YfiR complex. The YfiBL43P molecules are shown in cyan and yellow. The YfiR molecules are shown in green and magenta. Two interacting regions are highlighted by red rectangles. (B) Structural superposition of apo YfiB and YfiR-bound YfiBL43P. To illustrate the differences between apo YfiB and YfiR-bound YfiBL43P, the apo YfiB is shown in pink, except residues 34–70 are shown in red, whereas the YfiR-bound YfiBL43P is shown in cyan, except residues 44–70 are shown in blue. (C) Close-up view of the differences between apo YfiB and YfiR-bound YfiBL43P. The residues proposed to contribute to YfiB activation are illustrated in sticks. The key residues in apo YfiB are shown in red and those in YfiBL43P are shown in blue. (D) Close-up views showing interactions in regions I and II. YfiBL43P and YfiR are shown in cyan and green, respectively. (E and F) The conserved surface in YfiR contributes to the interaction with YfiB. (G) The residues of YfiR responsible for interacting with YfiB are shown in green sticks, and the proposed YfiN-interacting residues are shown in yellow sticks. The red sticks, which represent the YfiB-interacting residues, are also responsible for the proposed interactions with YfiN To gain structural insights into the YfiB-YfiR interaction, we co-expressed YfiB (residues 34–168) and YfiR (residues 35–190, lacking the signal peptide), but failed to obtain the complex, in accordance with a previous report in which no stable complex of YfiB-YfiR was observed (Malone et al.,). It has been reported that single mutants of Q39, L43, F48 and W55 contribute to YfiB activation leading to the induction of the SCV phenotype in P. aeruginosa PAO1 (Malone et al.,). It is likely that these residues may be involved in the conformational changes of YfiB that are related to YfiR sequestration (Fig. 3C). Therefore, we constructed two such single mutants of YfiB (YfiBL43P and YfiBF48S). As expected, both mutants form a stable complex with YfiR. Finally, we crystalized YfiR in complex with the YfiBL43P mutant and solved the structure at 1.78 Å resolution by molecular replacement using YfiR and YfiB as models. The YfiB-YfiR complex is a 2:2 heterotetramer (Fig. 3A) in which the YfiR dimer is clamped by two separated YfiBL43P molecules with a total buried surface area of 3161.2 Å2. The YfiR dimer in the complex is identical to the non-oxidized YfiR dimer alone (Yang et al.,), with only Cys145-Cys152 of the two disulfide bonds well formed, suggesting Cys71-Cys110 disulfide bond formation is not essential for forming YfiB-YfiR complex. The N-terminal structural conformation of YfiBL43P, from the foremost N-terminus to residue D70, is significantly altered compared with that of the apo YfiB. The majority of the α1 helix (residues 34–43) is invisible on the electron density map, and the α2 helix and β1 and β2 strands are rearranged to form a long loop containing two short α-helix turns (Fig. 3B and 3C), thus embracing the YfiR dimer. The observed changes in conformation of YfiB and the results of mutagenesis suggest a mechanism by which YfiB sequesters YfiR. The YfiB-YfiR interface can be divided into two regions (Fig. 3A and 3D). Region I is formed by numerous main-chain and side-chain hydrophilic interactions between residues E45, G47 and E53 from the N-terminal extended loop of YfiB and residues S57, R60, A89 and H177 from YfiR (Fig. 3D-I(i)). Additionally, three hydrophobic anchoring sites exist in region I. The residues F48 and W55 of YfiB are inserted into the hydrophobic cores mainly formed by the main chain and side chain carbon atoms of residues S57/Q88/A89/N90 and R60/R175/H177 of YfiR, respectively; and F57 of YfiB is inserted into the hydrophobic pocket formed by L166/I169/V176/P178/L181 of YfiR (Fig. 3D-I(ii)). In region II, the side chains of R96, E98 and E157 from YfiB interact with the side chains of E163, S146 and R171 from YfiR, respectively. Additionally, the main chains of I163 and V165 from YfiB form hydrogen bonds with the main chains of L166 and A164 from YfiR, respectively, and the main chain of P166 from YfiB interacts with the side chain of R185 from YfiR (Fig. 3D-II). These two regions contribute a robust hydrogen-bonding network to the YfiB-YfiR interface, resulting in a tightly bound complex. Based on the observations that two separated YfiBL43P molecules form a 2:2 complex structure with YfiR dimer, we performed an analytical ultracentrifugation experiment to check the oligomeric states of wild-type YfiB and YfiBL43P. The results showed that wild-type YfiB exists in both monomeric and dimeric states in solution, while YfiBL43P primarily adopts the monomer state in solution (Fig. 1C–D). This suggests that the N-terminus of YfiB plays an important role in forming the dimeric YfiB in solution and that the conformational change of residue L43 is associated with the stretch of the N-terminus and opening of the dimer. Therefore, it is possible that both dimeric types might exist in solution. For simplicity, we only discuss the “head to head” dimer in the following text. The PG-binding site of YfiB The PG-binding site in YfiB. (A) Structural superposition of the PG-binding sites of the H. influenzae Pal/PG-P complex and YfiR-bound YfiBL43P complexed with sulfate ions. (B) Close-up view showing the key residues of Pal interacting with the m-Dap5 ε-carboxylate group of PG-P. Pal is shown in wheat and PG-P is in magenta. (C) Close-up view showing the key residues of YfiR-bound YfiBL43P interacting with a sulfate ion. YfiR-bound YfiBL43P is shown in cyan; the sulfate ion, in green; and the water molecule, in yellow. (D) Structural superposition of the PG-binding sites of apo YfiB and YfiR-bound YfiBL43P, the key residues are shown in stick. Apo YfiB is shown in yellow and YfiR-bound YfiBL43P in cyan. (E and F) MST data and analysis for binding affinities of (E) YfiB wild-type and (F) YfiBL43P with PG. (G) The sequence alignment of P. aeruginosa and E. coli sources of YfiB, Pal and the periplasmic domain of OmpA PG-associated lipoprotein (Pal) is highly conserved in Gram-negative bacteria and anchors to the outer membrane through an N-terminal lipid attachment and to PG layer through its periplasmic domain, which is implicated in maintaining outer membrane integrity. Previous homology modeling studies suggested that YfiB contains a Pal-like PG-binding site (Parsons et al.,), and the mutation of two residues at this site, D102 and G105, reduces the ability for biofilm formation and surface attachment (Malone et al.,). In the YfiB-YfiR complex, one sulfate ion is found at the bottom of each YfiBL43P molecule (Fig. 3A) and forms a strong hydrogen bond with D102 of YfiBL43P (Fig. 4A and 4C). Structural superposition between YfiBL43P and Haemophilus influenzae Pal complexed with biosynthetic peptidoglycan precursor (PG-P), UDP-N-acetylmuramyl-L-Ala-α-D-Glu-m-Dap-D-Ala-D-Ala (m-Dap is meso-diaminopimelate) (PDB code: 2aiz) (Parsons et al.,), revealed that the sulfate ion is located at the position of the m-Dap5 ϵ-carboxylate group in the Pal/PG-P complex (Fig. 4A). In the Pal/PG-P complex structure, the m-Dap5 ϵ-carboxylate group interacts with the side-chain atoms of D71 and the main-chain amide of D37 (Fig. 4B). Similarly, in the YfiR-bound YfiBL43P structure, the sulfate ion interacts with the side-chain atoms of D102 (corresponding to D71 in Pal) and R117 (corresponding to R86 in Pal) and the main-chain amide of N68 (corresponding to D37 in Pal). Moreover, a water molecule was found to bridge the sulfate ion and the side chains of N67 and D102, strengthening the hydrogen bond network (Fig. 4C). In addition, sequence alignment of YfiB with Pal and the periplasmic domain of OmpA (proteins containing PG-binding site) showed that N68 and D102 are highly conserved (Fig. 4G, blue stars), suggesting that these residues contribute to the PG-binding ability of YfiB. Interestingly, superposition of apo YfiB with YfiR-bound YfiBL43P revealed that the PG-binding region is largely altered mainly due to different conformation of the N68 containing loop. Compared to YfiBL43P, the N68-containing loop of the apo YfiB flips away about 7 Å, and D102 and R117 swing slightly outward; thus, the PG-binding pocket is enlarged with no sulfate ion or water bound (Fig. 4D). Therefore, we proposed that the PG-binding ability of inactive YfiB might be weaker than that of active YfiB. To validate this, we performed a microscale thermophoresis (MST) assay to measure the binding affinities of PG to wild-type YfiB and YfiBL43P, respectively. The results indicated that the PG-binding affinity of YfiBL43P is 65.5 μmol/L, which is about 16-fold stronger than that of wild-type YfiB (Kd = 1.1 mmol/L) (Fig. 4E–F). As the experiment is performed in the absence of YfiR, it suggests that an increase in the PG-binding affinity of YfiB is not a result of YfiB-YfiR interaction and is highly coupled to the activation of YfiB characterized by a stretched N-terminal conformation. The conserved surface in YfiR is functional for binding YfiB and YfiN Calculation using the ConSurf Server (http://consurf.tau.ac.il/), which estimates the evolutionary conservation of amino acid positions and visualizes information on the structure surface, revealed a conserved surface on YfiR that contributes to the interaction with YfiB (Fig. 3E and 3F). Interestingly, the majority of this conserved surface contributes to the interaction with YfiB (Fig. 3E and 3F). Malone JG et al. have reported that F151, E163, I169 and Q187, located near the C-terminus of YfiR, comprise a putative YfiN binding site (Malone et al.,). Interestingly, these residues are part of the conserved surface of YfiR (Fig. 3G). F151, E163 and I169 form a hydrophobic core while, Q187 is located at the end of the α6 helix. E163 and I169 are YfiB-interacting residues of YfiR, in which E163 forms a hydrogen bond with R96 of YfiB (Fig. 3D-II) and I169 is involved in forming the L166/I169/V176/P178/L181 hydrophobic core for anchoring F57 of YfiB (Fig. 3D-I(ii)). Collectively, a part of the YfiB-YfiR interface overlaps with the proposed YfiR-YfiN interface, suggesting alteration in the association-disassociation equilibrium of YfiR-YfiN and hence the ability of YfiB to sequester YfiR. YfiR binds small molecules Previous studies indicated that YfiR constitutes a YfiB-independent sensing device that can activate YfiN in response to the redox status of the periplasm, and we have reported YfiR structures in both the non-oxidized and the oxidized states earlier, revealing that the Cys145-Cys152 disulfide bond plays an essential role in maintaining the correct folding of YfiR (Yang et al.,). However, whether YfiR is involved in other regulatory mechanisms is still an open question. Overall Structures of VB6-bound and Trp-bound YfiR. (A) Superposition of the overall structures of VB6-bound and Trp-bound YfiR. (B) Close-up views showing the key residues of YfiR that bind VB6 and L-Trp. The electron densities of VB6 and Trp are countered at 3.0σ and 2.3σ, respectively, in |Fo|-|Fc| maps. (C) Superposition of the hydrophobic pocket of YfiR with VB6, L-Trp and F57 of YfiB Intriguingly, a Dali search (Holm and Rosenstrom,) indicated that the closest homologs of YfiR shared the characteristic of being able to bind several structurally similar small molecules, such as L-Trp, L-Phe, B-group vitamins and their analogs, encouraging us to test whether YfiR can recognize these molecules. For this purpose, we co-crystallized YfiR or soaked YfiR crystals with different small molecules, including L-Trp and B-group vitamins. Fortunately, we found obvious small-molecule density in the VB6-bound and Trp-bound YfiR crystal structures (Fig. 5A and 5B), and in both structures, the YfiR dimers resemble the oxidized YfiR structure in which both two disulfide bonds are well formed (Yang et al.,). Functional analysis of VB6 and L-Trp. (A and B) The effect of increasing concentrations of VB6 or L-Trp on YfiBL43P-induced attachment (bars). The relative optical density is represented as curves. Wild-type YfiB is used as negative control. (C and D) BIAcore data and analysis for binding affinities of (C) VB6 and (D) L-Trp with YfiR. (E–G) ITC data and analysis for titration of (E) YfiB wild-type, (F) YfiBL43P, and (G) YfiBL43P/F57A into YfiR Structural analyses revealed that the VB6 and L-Trp molecules are bound at the periphery of the YfiR dimer, but not at the dimer interface. Interestingly, VB6 and L-Trp were found to occupy the same hydrophobic pocket, formed by L166/I169/V176/P178/L181 of YfiR, which is also a binding pocket for F57 of YfiB, as observed in the YfiB-YfiR complex (Fig. 5C). To evaluate the importance of F57 in YfiBL43P-YfiR interaction, the binding affinities of YfiBL43P and YfiBL43P/F57A for YfiR were measured by isothermal titration calorimetry (ITC). The results showed Kd values of 1.4 × 10−7 mol/L and 5.3 × 10−7 mol/L for YfiBL43P and YfiBL43P/F57A, respectively, revealing that the YfiBL43P/F57A mutant caused a 3.8-fold reduction in the binding affinity compared with the YfiBL43P mutant (Fig. 6F and 6G). In parallel, to better understand the putative functional role of VB6 and L-Trp, yfiB was deleted in a PAO1 wild-type strain, and a construct expressing the YfiBL43P mutant was transformed into the PAO1 ΔyfiB strain to trigger YfiBL43P-induced biofilm formation. Growth and surface attachment assays were carried out for the yfiB-L43P strain in the presence of increasing concentrations of VB6 or L-Trp. As shown in Fig. 6A and 6B, the over-expression of YfiBL43P induced strong surface attachment and much slower growth of the yfiB-L43P strain, and as expected, a certain amount of VB6 or L-Trp (4–6 mmol/L for VB6 and 6–10 mmol/L for L-Trp) could reduce the surface attachment. Interestingly, at a concentration higher than 8 mmol/L, VB6 lost its ability to inhibit biofilm formation, implying that the VB6-involving regulatory mechanism is highly complicated and remains to be further investigated. Of note, both VB6 and L-Trp have been reported to correlate with biofilm formation in certain Gram-negative bacteria (Grubman et al.,; Shimazaki et al.,). In Helicobacter pylori in particular, VB6 biosynthetic enzymes act as novel virulence factors, and VB6 is required for full motility and virulence (Grubman et al.,). In E. coli, mutants with decreased tryptophan synthesis show greater biofilm formation, and matured biofilm is degraded by L-tryptophan addition (Shimazaki et al.,). However, the detailed mechanism remains elusive. To answer the question whether competition of VB6 or L-Trp for the YfiB F57-binding pocket of YfiR plays an essential role in inhibiting biofilm formation, we measured the binding affinities of VB6 and L-Trp for YfiR via BIAcore experiments. The results showed relatively weak Kd values of 35.2 mmol/L and 76.9 mmol/L for VB6 and L-Trp, respectively (Fig. 6C and 6D). Based on our results, we concluded that VB6 or L-Trp can bind to YfiR, however, VB6 or L-Trp alone may have little effects in interrupting the YfiB-YfiR interaction, the mechanism by which VB6 or L-Trp inhibits biofilm formation remains unclear and requires further investigation. DISCUSSION Previous studies suggested that in response to cell stress, YfiB in the outer membrane sequesters the periplasmic protein YfiR, releasing its inhibition of YfiN on the inner membrane and thus inducing the diguanylate cyclase activity of YfiN to allow c-di-GMP production (Giardina et al.,; Malone et al.,; Malone et al.,). However, the pattern of interaction between these proteins and the detailed regulatory mechanism remain unknown due to a lack of structural information. Here, we report the crystal structures of YfiB alone and an active mutant YfiBL43P in complex with YfiR, indicating that YfiR forms a 2:2 complex with YfiB via a region composed of conserved residues. Our structural data analysis shows that the activated YfiB has an N-terminal portion that is largely altered, adopting a stretched conformation compared with the compact conformation of the apo YfiB. The apo YfiB structure constructed beginning at residue 34 has a compact conformation of approximately 45 Å in length. In addition to the preceding 8 aa loop (from the lipid acceptor Cys26 to Gly34), the full length of the periplasmic portion of apo YfiB can reach approximately 60 Å. It was reported that the distance between the outer membrane and the cell wall is approximately 50 Å and that the thickness of the PG layer is approximately 70 Å (Matias et al.,). Thus, YfiB alone represents an inactive form that may only partially insert into the PG matrix. By contrast, YfiR-bound YfiBL43P (residues 44–168) has a stretched conformation of approximately 55 Å in length. In addition to the 17 preceding intracellular residues (from the lipid acceptor Cys26 to Leu43), the length of the intracellular portion of active YfiB may extend over 100 Å, assuming a fully stretched conformation. Provided that the diameter of the widest part of the YfiB dimer is approximately 64 Å, which is slightly smaller than the smallest diameter of the PG pore (70 Å) (Meroueh et al.,), the YfiB dimer should be able to penetrate the PG layer. Regulatory model of the YfiBNR tripartite system. The periplasmic domain of YfiB and the YfiB-YfiR complex are depicted according to the crystal structures. The lipid acceptor Cys26 is indicated as blue ball. The loop connecting Cys26 and Gly34 of YfiB is modeled. The PAS domain of YfiN is shown as pink oval. Once activated by certain cell stress, the dimeric YfiB transforms from a compact conformation to a stretched conformation, allowing the periplasmic domain of the membrane-anchored YfiB to penetrate the cell wall and sequester the YfiR dimer, thus relieving the repression of YfiN These results, together with our observation that activated YfiB has a much higher cell wall binding affinity, and previous mutagenesis data showing that (1) both PG binding and membrane anchoring are required for YfiB activity and (2) activating mutations possessing an altered N-terminal loop length are dominant over the loss of PG binding (Malone et al.,), suggest an updated regulatory model of the YfiBNR system (Fig. 7). In this model, in response to a particular cell stress that is yet to be identified, the dimeric YfiB is activated from a compact, inactive conformation to a stretched conformation, which possesses increased PG binding affinity. This allows the C-terminal portion of the membrane-anchored YfiB to reach, bind and penetrate the cell wall and sequester the YfiR dimer. The YfiBNR system provides a good example of a delicate homeostatic system that integrates multiple signals to regulate the c-di-GMP level. Homologs of the YfiBNR system are functionally conserved in P. aeruginosa (Malone et al.,; Malone et al.,), E. coli (Hufnagel et al.,; Raterman et al.,; Sanchez-Torres et al.,), K. pneumonia (Huertas et al.,) and Y. pestis (Ren et al.,), where they affect c-di-GMP production and biofilm formation. The mechanism by which activated YfiB relieves the repression of YfiN may be applicable to the YfiBNR system in other bacteria and to analogous outside-in signaling for c-di-GMP production, which in turn may be relevant to the development of drugs that can circumvent complicated antibiotic resistance. MATERIALS AND METHODS Protein expression and purification P. aeruginosa YfiR (residues 35–190, lacking the predicted N-terminal periplasmic localization signaling peptide) and YfiB (residues 34–168, lacking the signal peptide from residues 1–26 and periplasmic residues 27–33) were cloned into ORF1 of the pETDuet-1 (Merck Millipore, Darmstadt, Germany) vector via the BamHI and HindIII restriction sites, with a constructed N-terminal His6 and a TEV cleavage site, respectively. In addition, YfiB (residues 34–168) was ligated into the NdeI and XhoI restriction sites of ORF2 in the previously constructed YfiR expression vector. Site-directed mutagenesis was carried out using a QuikChange kit (Agilent Technologies, Santa Clara, CA), following the manufacturer’s instructions. The proteins were over-expressed in the E. coli BL21-CodonPlus(DE3)-RIPL strain. Protein expression was induced by adding 0.5–1 mmol/L IPTG at an OD600 of approximately 0.8. The cell cultures were then incubated for an additional 4.5 h at 37°C. The cells were subsequently harvested by centrifugation and stored at −80°C. Cell suspensions were thawed and homogenized using a high-pressure homogenizer (JNBIO, Beijing, China). YfiR was first purified by Ni affinity chromatography and then incubated with His6-tagged TEV protease overnight. The His6-TEV cleavage site was subsequently removed by incubation with Ni-NTA resin. Finally, YfiR was purified with a HiTrap STM column (GE Healthcare), followed by a Superdex 200 (GE Healthcare) column. YfiB was purified with Ni affinity chromatography, followed by a Superdex 200 (GE Healthcare) column. The YfiB-YfiR complex was first purified by Ni affinity chromatography, then by a Superdex 200 (GE Healthcare) column, and finally by a HiTrap STM column (GE Healthcare). All of the purified fractions were collected and concentrated to ~40 mg/mL in 20 mmol/L Tris-HCl (pH 8.0) and 200 mmol/L NaCl, frozen in liquid nitrogen and stored at −80°C. Crystallization and data collection Crystal screening was performed with commercial screening kits (Hampton Research, CA, USA) using the sitting-drop vapor diffusion method, and positive hits were optimized using the hanging-drop vapor diffusion method at 293 K. Crystals of the YfiB protein were obtained and optimized in buffer containing 0.2 mol/L lithium sulfate monohydrate, 0.1 mol/L Tris-HCl (pH 8.0) and 30% w/v polyethylene glycol 4000. After being soaked for a few seconds in cryoprotection solution (well solution complemented with 25% xylitol), the crystals were cooled by plunging them into liquid nitrogen. Diffraction-quality crystals of the YfiB-YfiR complex were grown in buffer containing 0.2 mol/L ammonium sulfate, 0.1 mol/L Tris-HCl (pH 8.0) and 12% w/v polyethylene glycol 8000. The crystals were cryoprotected with 8% (w/v) polyethylene glycol 8000 and 0.1 mol/L Tris-HCl (pH 7.5) supplemented with saturated sucrose prior to being flash frozen. Crystals of the native YfiR were obtained and optimized in 0.1 mol/L HEPES (pH 7.5) and 1.8 mol/L ammonium sulfate. VB6-bound YfiR crystals were obtained by soaking the native YfiR crystals in 2 mmol/L VB6 molecules. Trp-bound YfiR crystals were obtained by co-crystalizing the YfiR protein and 4 mmol/L L-Trp molecules in 0.2 mol/L NaCl, 0.1 mol/L BIS-TRIS (pH 5.5), and 25% w/v polyethylene glycol 3350. For cryoprotection, both the VB6-bound and the L-Trp-bound YfiR crystals were soaked in 2.5 mol/L lithium sulfate monohydrate for a few seconds before data collection. Diffraction data for the YfiB crystal belonging to space group P21 was collected in house, the data for the YfiB crystal belonging to space group P41 and for the Trp-bound YfiR crystal were collected on beamline BL17U at the Shanghai Synchrotron Radiation Facility (SSRF), and the data for the VB6-bound YfiR crystal were collected on beamline BL18U at SSRF. Finally, the data for the YfiB-YfiR complex crystal were collected on beamline BL-1A at the Photon Factory in Japan. The diffraction data were processed with the HKL2000 software program (Otwinowski and Minor,). Structure determination and refinement Data collection, phasing and refinement statistics Data collection\tYfiB (crystal form I)\tYfiB (crystal form II)\tVB6-bound YfiR\tTrp-bound YfiR\tYfiBL43P-YfiR\t \tSpace group\tP21\tP41\tP43212\tP43212\tP1\t \tWavelength (Å)\t1.54187\t0.9791\t0.97861\t0.9791\t1.10000\t \tResolution (Å)a\t50.0–2.15 (2.19–2.15)\t50.0–2.80 (2.85–2.8)\t50.0–2.4 (2.44–2.4)\t50.0–2.5 (2.54–2.5)\t50–1.78 (1.86–1.78)\t \tCell dimensions\t \t a, b, c (Å)\t65.85, 90.45, 66.30\t46.95, 46.95, 154.24\t120.24, 120.24, 84.99\t120.88, 120.88, 88.46\t49.50, 58.57, 69.86\t \t α, β, γ (°)\t90, 113.87, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t72.93, 96.98, 90.19\t \t Unique reflections\t37,625 (1866)\t8,105 (412)\t24,776 (1202)\t23170 (1132)\t67,774 (6615)\t \t I/σI\t19.59 (2.62)\t12.36 (4.15)\t20.17 (2.4)\t39.5 (4.68)\t17.75 (1.89)\t \t Completeness (%)\t97.1 (95.4)\t97.8 (100)\t99.1 (98.8)\t99.9 (100)\t96.5 (94.6)\t \t Rmerge (%)\t6.5 (44.5)\t14.6 (49.7)\t8.9 (56.8)\t9.4 (89.2)\t5.6 (46.3)\t \t Rmeas (%)\t7.4 (51.6)\t15.4 (52.0)\t9.6 (61.7)\t9.6 (90.8)\t6.6 (55.1)\t \t CC1/2b\t0.747\t0.952\t0.899\t0.974\t0.849\t \tRefinement\t \t Rwork (%)\t20.14\t19.17\t17.82\t18.66\t17.90\t \t Rfree(%)\t26.29\t26.49\t19.81\t23.05\t20.61\t \tAverage B factors (Å2)\t \t Protein\t25.54\t42.70\t38.68\t35.03\t32.54\t \t VB6\t-\t-\t44.08\t-\t-\t \t Trp\t-\t-\t-\t87.51\t-\t \t SO42−\t37.16\t66.52\t51.55\t41.93\t45.51\t \t H2O\t32.91\t36.09\t40.58\t34.75\t43.52\t \tRoot mean square deviations\t \t Bond lengths (Å)\t0.009\t0.009\t0.007\t0.007\t0.007\t \t Bond angles (°)\t1.085\t1.132\t1.021\t0.977\t1.110\t \tRamachandran plot\t \t Most favored (%)\t92.6\t87.7\t96.5\t98.1\t94.2\t \t Additionally allowed (%)\t7.4\t12.3\t3.5\t1.9\t5.8\t \t Generously allowed (%)\t0\t0\t0\t0\t0\t \t Disallowed\t0\t0\t0\t0\t0\t \t \na Numbers in parentheses are for the highest resolution shell \nb The values of CC1/2 are for the highest resolution shell The two YfiB crystal structures respectively belonging to space groups P21 and P41 were both solved by molecular replacement (Lebedev et al.,) using the putative MotB-like protein DVU_2228 from D. vulgaris as a model (PDB code: 3khn) at 2.15 Å and 2.8 Å resolution, respectively. Both the VB6-bound and the Trp-bound YfiR crystals belonging to space group P43212, with a dimer in the asymmetric unit, were solved by molecular replacement (Lebedev et al.,) using native YfiR as a model (PDB code: 4YN7) at 2.4 Å and 2.5 Å resolution, respectively. The YfiB-YfiR crystal belonging to space group P1, with a 2:2 heterotetramer in the asymmetric unit, was solved by molecular replacement using YfiR and YfiB as models. Electron density maps were calculated using PHENIX (Adams et al.,). Model building was performed using COOT (Emsley et al.,) and refined with PHENIX (Adams et al.,; Afonine et al.,). The final structures were analyzed with PROCHECK (Laskowski et al.,). Data collection and refinement statistics are presented in Table 1. The figures depicting structures were prepared using PyMOL (http://www.pymol.org). Atomic coordinates and structure factors have been deposited in the RCSB Protein Data Bank (http://www.pdb.org) under accession codes 5EAZ, 5EB0, 5EB1, 5EB2 and 5EB3. Analytical ultracentrifugation Sedimentation velocity measurements were performed on a Beckman ProteomeLab XL-I at 25°C. All protein samples were diluted to an OD280 of 0.7 in 20 mmol/L Tris (pH 8.0) and 200 mmol/L NaCl. Data were collected at 60,000 rpm. (262,000 ×g) every 3 min at a wavelength of 280 nm. Interference sedimentation coefficient distributions, or c(M), were calculated from the sedimentation velocity data using SEDFIT (Schuck,). PG preparation PG was extracted from the E. coli DH5α strain by following a method described previously (Desmarais et al.,). Briefly, cells were cultured until they reached an OD600 of 0.7–0.8 and then collected at 5,000 ×g, 4°C. The collected bacteria were dripped into the boiling 6% (w/v) SDS and stirred at 500 rpm in a boiling water bath for 3 h before incubating overnight at room temperature. The large PG polymers were collected by ultracentrifugation at 130,000 ×g for 1 h at room temperature and washed repeatedly to remove SDS. The pellet was treated with Pronase E (200 μg/mL final concentration) for 3 h at 60°C followed by SDS to remove contaminating proteins and washed three times to remove the SDS by ultracentrifugation. Next, the samples were treated with lysozyme (200 μg/mL final concentration) for 16 h at 37°C. Finally, the purified PG is obtained by treating the samples in a boiling water bath for 10 min and centrifuging it at 13,000 ×g to remove the contaminating lysozyme. Microscale thermophoresis (MST) Purified YfiB wild-type and it mutant YfiBL43P were fluorescently labeled using the NanoTemper blue protein-labeling kit according to the manufacturer’s protocol. This resulted in coupling of the fluorescent dye NT-495. PG was titrated in 1:1 dilutions starting at 1 mmol/L. To determine of the Kd values, 10 μL labeled protein was mixed with 10 μL PG at various concentrations in Hepes buffer (20 mmol/L Hepes, 200 mmol/L NaCl, 0.005% Tween-20, pH 7.5). After 10 min of incubation, all binding reaction mixtures were loaded into the MST-grade glass capillaries (NanoTemper Technologies), and thermophoresis was measured with a NanoTemper Monolith-NT115 system (20% light-emitting diode, 20% IR laser power). Deletion of the yfiB genes The yfiB deletion construct was produced by SOE-PCR (Hmelo et al.,) and contained homologous flanking regions to the target gene. This construct was ligated into the pEX18Gm vector between the HindIII and the KpnI sites. The resulting vector was then used to delete yfiB by two-step allelic exchange (Hmelo et al.,). After being introduced into PAO1 via biparental mating with E. coli SM10 (λpir), single crossovers were selected on Vogel-Bonner Minimal Medium (VBMM), which was used for counter-selection against E. coli (P. aeruginosa can utilize citrate as a sole carbon source and energy source, whereas E. coli cannot), containing 50 μg/mL gentamycin. Restreaking was then performed on no-salt Luria-Bertani (NSLB) agar that contained 15% sucrose to force the resolution of double crossovers. Deletion of yfiB in the strains was confirmed by colony PCR. For complementation experiments, yfiB wild-type and L43P mutant genes were cloned into the pJN105 vector via the EcoRI and XbaI restriction sites, respectively. The plasmids were then individually transformed into the PAO1 ΔyfiB strain using the rapid electroporation method described in Choi KH et al. (Choi et al.,). Transformants were selected on LB plates containing 50 μg/mL gentamycin. For induction, arabinose was added to a final concentration of 0.2%. Attachment assays The attachment assays were carried out using the MBECTM (Minimum Biofilm Eradication Concentration, Innovotech, Inc.) biofilm inoculator, which consists of a plastic lid with 96 pegs and 96 individual wells. The MBEC plates containing 150 μL LB medium/well were inoculated with 1% overnight cultures of the yfiB-L43P strain and incubated overnight at 37°C without shaking. VB6, L-Trp and arabinose were added as appropriate. The peg lids were washed with distilled water, and the attached cell material was then stained with 0.1% crystal violet solution (5% methanol, 5% isopropanol) before further washing to remove excess dye. The crystal violet was re-dissolved in 20% acetic acid solution, and the absorbance was measured at 600 nm. Assays were performed with 12 wells/strain and repeated independently for each experiment. BIAcore analysis The interaction kinetics of YfiR with VB6 and L-Trp were examined on a SPR machine Biacore 3000 (GE Healthcare) at 25°C. The running buffer (20 mmol/L HEPES, pH 7.5, 150 mmol/L NaCl, 0.005% (v/v) Tween-20) was vacuum filtered, and degassed immediately prior to use. YfiR at 10 μg/mL in 10 mmol/L sodium acetate (pH 5.5) was immobilized to 3000 response units on the carboxymethylated dextran surface-modified chip (CM5 chip). The binding affinities were evaluated over a range of 2.5–40 mmol/L concentrations. Meanwhile, for both binding assays, the concentration of 10 mmol/L was repeated as an internal control. All of the data collected were analyzed using BIAevaluation software version 4.1. ITC assays ITC experiments were performed in a buffer composed of 20 mmol/L Tris (pH 8.0) and 150 mmol/L NaCl at 25°C using an iTC200 calorimeter (GE Healthcare). YfiB wild-type or its mutants (YfiBL43P, YfiBL43P/F57A) (0.4 mmol/L, in the syringe) was titrated into YfiR (0.04 mmol/L, in the cell), respectively. The titration sequence included a single 0.5 µL injection, followed by 19 injections of 2 µL each, with a 2-min interval between injections and a stirring rate of 1000 rpm. The calorimetric data were then analyzed with OriginLab software (GE Healthcare). Min Xu, Xuan Yang and Xiu-An Yang have contributed equally to this work. References PHENIX: a comprehensive Python-based system for macromolecular structure solution Towards automated crystallographic structure refinement with phenix.refine Experimental evolution of bet hedging Second messenger-mediated adjustment of bacterial swimming velocity Targeting cyclic di-GMP signalling: a strategy to control biofilm formation? Bacterial small-molecule signaling pathways A 10-min method for preparation of highly electrocompetent Pseudomonas aeruginosa cells: application for DNA fragment transfer between chromosomes and plasmid transformation Desmarais SM, Cava F, de Pedro MA, Huang KC (2014) Isolation and preparation of bacterial cell walls for compositional analysis by ultra performance liquid chromatography. J Vis Exp 83:e51183 Second messenger-mediated spatiotemporal control of protein degradation regulates bacterial cell cycle progression Features and development of Coot Small colony variants of Pseudomonas aeruginosa in chronic bacterial infection of the lung in cystic fibrosis Investigating the allosteric regulation of YfiN from Pseudomonas aeruginosa: clues from the structure of the catalytic domain Mutational activation of niche-specific genes provides insight into regulatory networks and bacterial function in a complex environment Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia Grubman A, Phillips A, Thibonnier M, Kaparakis-Liaskos M, Johnson C, Thiberge JM, Radcliff FJ, Ecobichon C, Labigne A, de Reuse H. et al (2010) Vitamin B6 is required for full motility and virulence in Helicobacter pylori. MBio 1 Ha DG, O’Toole GA (2015) c-di-GMP and its effects on biofilm formation and dispersion: a Pseudomonas aeruginosa review. Microbiol Spectr 3, MB-0003-2014 Small-colony variants of Pseudomonas aeruginosa in cystic fibrosis Highly adherent small-colony variants of Pseudomonas aeruginosa in cystic fibrosis lung infection Principles of c-di-GMP signalling in bacteria A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels Precision-engineering the Pseudomonas aeruginosa genome with two-step allelic exchange Dali server: conservation mapping in 3D Klebsiella pneumoniae yfiRNB operon affects biofilm formation, polysaccharide production and drug susceptibility The disulfide bonding system suppresses CsgD-independent cellulose production in Escherichia coli Cyclic di-guanosine-monophosphate comes of age: a novel secondary messenger involved in modulating cell surface structures in bacteria? HmsP, a putative phosphodiesterase, and HmsT, a putative diguanylate cyclase, control Hms-dependent biofilm formation in Yersinia pestis Characterization of colony morphology variants isolated from Pseudomonas aeruginosa biofilms Analysis of Pseudomonas aeruginosa diguanylate cyclases and phosphodiesterases reveals a role for bis-(3′-5′)-cyclic-GMP in virulence PROCHECK: a program to check the stereochemical quality of protein structures Model preparation in MOLREP and examples of model improvement using X-ray data Structural insights into YfiR sequestering by YfiB in Pseudomonas aeruginosa PAO1 Role of small colony variants in persistence of Pseudomonas aeruginosa infections in cystic fibrosis lungs YfiBNR mediates cyclic di-GMP dependent small colony variant formation and persistence in Pseudomonas aeruginosa The YfiBNR signal transduction mechanism reveals novel targets for the evolution of persistent Pseudomonas aeruginosa in cystic fibrosis airways Cryo-transmission electron microscopy of frozen-hydrated sections of Escherichia coli and Pseudomonas aeruginosa Three-dimensional structure of the bacterial cell wall peptidoglycan Structural basis for c-di-GMP-mediated inside-out signaling controlling periplasmic proteolysis Processing of X-ray diffraction data collected in oscillation mode Bacterial biofilms: an emerging link to disease pathogenesis Peptidoglycan recognition by Pal, an outer membrane lipoprotein Bacterial nucleotide-based second messengers Genetic analysis of the role of yfiR in the ability of Escherichia coli CFT073 to control cellular cyclic dimeric GMP levels and to persist in the urinary tract Development and persistence of antimicrobial resistance in Pseudomonas aeruginosa: a longitudinal observation in mechanically ventilated patients HmsC, a periplasmic protein, controls biofilm formation via repression of HmsD, a diguanylate cyclase in Yersinia pestis Cyclic di-GMP: the first 25 years of a universal bacterial second messenger Regulation of cellulose synthesis in Acetobacter xylinum by cyclic diguanylic acid Cellulose biosynthesis and function in bacteria GGDEF proteins YeaI, YedQ, and YfiN reduce early biofilm formation and swimming motility in Escherichia coli Structural and mechanistic determinants of c-di-GMP signalling Size-distribution analysis of macromolecules by sedimentation velocity ultracentrifugation and lamm equation modeling L-Tryptophan prevents Escherichia coli biofilm formation and triggers biofilm degradation Genetic adaptation by Pseudomonas aeruginosa to the airways of cystic fibrosis patients The EAL domain protein VieA is a cyclic diguanylate phosphodiesterase Connecting quorum sensing, c-di-GMP, pel polysaccharide, and biofilm formation in Pseudomonas aeruginosa through tyrosine phosphatase TpbA (PA3885) Structural and biochemical analysis of tyrosine phosphatase related to biofilm formation A (TpbA) from the opportunistic pathogen Pseudomonas aeruginosa PAO1 Crystal structures of 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diff --git a/annotated_BioC_JSON/PMC4888278_ann.json b/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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1,3-dihydro-2,1,3-benzothiadiazole 2,2-dioxide analogs as new RORC modulators Minor Structural Change to Tertiary Sulfonamide RORc Ligands Led to Opposite Mechanisms of Action Discovery of novel pyrazole-containing benzamides as potent RORγ inverse agonists Discovery of biaryl carboxylamides as potent RORγ inverse 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diff --git a/annotated_BioC_JSON/PMC4896748_ann.json b/annotated_BioC_JSON/PMC4896748_ann.json
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--- /dev/null
+++ b/annotated_BioC_JSON/PMC4896748_ann.json
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+[{"sourceid":"4896748","sourcedb":"","project":"","target":"","text":"Ensemble cryo-EM uncovers inchworm-like translocation of a viral IRES through the ribosome Internal ribosome entry sites (IRESs) mediate cap-independent translation of viral mRNAs. Using electron cryo-microscopy of a single specimen, we present five ribosome structures formed with the Taura syndrome virus IRES and translocase eEF2•GTP bound with sordarin. The structures suggest a trajectory of IRES translocation, required for translation initiation, and provide an unprecedented view of eEF2 dynamics. The IRES rearranges from extended to bent to extended conformations. This inchworm-like movement is coupled with ribosomal inter-subunit rotation and 40S head swivel. eEF2, attached to the 60S subunit, slides along the rotating 40S subunit to enter the A site. Its diphthamide-bearing tip at domain IV separates the tRNA-mRNA-like pseudoknot I (PKI) of the IRES from the decoding center. This unlocks 40S domains, facilitating head swivel and biasing IRES translocation via hitherto-elusive intermediates with PKI captured between the A and P sites. The structures suggest missing links in our understanding of tRNA translocation. DOI: http://dx.doi.org/10.7554/eLife.14874.001 Introduction Virus propagation relies on the host translational apparatus. To efficiently compete with host mRNAs and engage in translation under stress, some viral mRNAs undergo cap-independent translation. To this end, internal ribosome entry site (IRES) RNAs are employed (reviewed in. An IRES is located at the 5’ untranslated region of the viral mRNA, preceding an open reading frame (ORF). To initiate translation, a structured IRES RNA interacts with the 40S subunit or the 80S ribosome, resulting in precise positioning of the downstream start codon in the small 40S subunit. The canonical scenario of cap-dependent and IRES-dependent initiation involves positioning of the AUG start codon and the initiator tRNAMet in the ribosomal peptidyl-tRNA (P) site, facilitated by interaction with initiation factors. Subsequent binding of an elongator aminoacyl-tRNA to the ribosomal A site transitions the initiation complex into the elongation cycle of translation. Upon peptide bond formation, the two tRNAs and their respective mRNA codons translocate from the A and P to P and E (exit) sites, freeing the A site for the next elongator tRNA. An unusual strategy of initiation is used by intergenic-region (IGR) IRESs found in Dicistroviridae arthropod-infecting viruses. These include shrimp-infecting Taura syndrome virus (TSV), and insect viruses Plautia stali intestine virus (PSIV) and Cricket paralysis virus (CrPV). The IGR IRES mRNAs do not contain an AUG start codon. The IGR-IRES-driven initiation does not involve initiator tRNAMet and initiation factors. As such, this group of IRESs represents the most streamlined mechanism of eukaryotic translation initiation. A recent demonstration of bacterial translation initiation by an IGR IRES indicates that the IRESs take advantage of conserved structural and dynamic properties of the ribosome. Early electron cryo-microscopy (cryo-EM) studies have found that the CrPV IRES packs in the ribosome intersubunit space. Recent cryo-EM structures of ribosome-bound TSV IRES and CrPV IRES revealed that IGR IRESs position the ORF by mimicking a translating ribosome bound with tRNA and mRNA. The ~200-nt IRES RNAs span from the A site beyond the E site. A conserved tRNA-mRNA–like structural element of pseudoknot I (PKI) interacts with the decoding center in the A site of the 40S subunit. The codon-anticodon-like helix of PKI is stabilized by interactions with the universally conserved decoding-center nucleotides G577, A1755 and A1756 (G530, A1492 and A1493 in E. coli 16S ribosomal RNA, or rRNA). The downstream initiation codon—coding for alanine—is placed in the mRNA tunnel, preceding the decoding center. PKI of IGR IRESs therefore mimics an A-site elongator tRNA interacting with an mRNA sense codon, but not a P-site initiator tRNAMet and the AUG start codon. How this non-canonical initiation complex transitions to the elongation step is not fully understood. For a cognate aminoacyl-tRNA to bind the first viral mRNA codon, PKI has to be translocated from the A site, so that the first codon can be presented in the A site. A cryo-EM structure of the ribosome bound with a CrPV IRES and release factor eRF1 occupying the A site provided insight into the post-translocation state. In this structure, PKI is positioned in the P site and the first mRNA codon is located in the A site. How the large IRES RNA translocates within the ribosome, allowing PKI translocation from the A to P site is not known. The structural similarity of PKI and the tRNA anticodon stem loop (ASL) bound to a codon suggests that their mechanisms of translocation are similar to some extent. Translocation of the IRES or tRNA-mRNA requires eukaryotic elongation factor 2 (eEF2), a structural and functional homolog of the well-studied bacterial EF-G. Pre-translocation tRNA-bound ribosomes contain a peptidyl- and deacyl-tRNA, both base-paired to mRNA codons in the A and P sites (termed 2tRNA•mRNA complex). Translocation of 2tRNA•mRNA involves two major large-scale ribosome rearrangements (Figure 1—figure supplement 1) (reviewed in). First, studies of bacterial ribosomes showed that a ~10° rotation of the small subunit relative to the large subunit, known as intersubunit rotation, or ratcheting, is required for translocation. Intersubunit rotation occurs spontaneously upon peptidyl transfer, and is coupled with formation of hybrid tRNA states. In the rotated pre-translocation ribosome, the peptidyl-tRNA binds the A site of the small subunit with its ASL and the P site of the large subunit with the CCA 3’ end (A/P hybrid state). Concurrently, the deacyl-tRNA interacts with the P site of the small subunit and the E site of the large subunit (P/E hybrid state). The ribosome can undergo spontaneous, thermally-driven forward-reverse rotation that shifts the two tRNAs between the hybrid and 'classical' states while the anticodon stem loops remain non-translocated. Binding of EF-G next to the A site and reverse rotation of the small subunit results in translocation of both ASLs on the small subunit. EF-G is thought to 'unlock' the pre-translocation ribosome, allowing movement of the 2tRNA•mRNA complex, however the structural details of this unlocking are not known. The second large-scale rearrangement involves rotation, or swiveling, of the head of the small subunit relative to the body. The head can rotate by up to ~20° around the axis nearly orthogonal to that of intersubunit rotation, in the absence of tRNA or in the presence of a single P/E tRNA and eEF2 or EF-G. Förster resonance energy transfer (FRET) data suggest that head swivel of the rotated small subunit facilitates EF-G-mediated movement of 2tRNA•mRNA. Structures of the 70S•EF-G complex bound with two nearly translocated tRNAs, exhibit a large 18° to 21° head swivel in a mid-rotated subunit, whereas no head swivel is observed in the fully rotated pre-translocation or in the non-rotated post-translocation 70S•2tRNA•EF-G structures. The structural role of head swivel is not fully understood. The head swivel was proposed to facilitate transition of the tRNA from the P to E site by widening a constriction between these sites on the 30S subunit. This widening allows the ASL to sample positions between the P and E sites. Whether and how the head swivel mediates tRNA transition from the A to P site remains unknown. Comparison of 70S•2tRNA•mRNA and 80S•IRES translocation complexes. (a) Structures of bacterial 70S•2tRNA•mRNA translocation complexes, ordered according to the position of the translocating A-\u003eP tRNA (orange). The large ribosomal subunit is shown in cyan; the small subunit in light yellow (head) and wheat-yellow (body), elongation factor G (EF-G) is shown in green. Nucleotides C1054, G966 and G693 of 16S rRNA are shown in black to denote the A, P and E sites, respectively. The extents of the 30S subunit rotation and head swivel relative to their positions in the post-translocation structure are shown with arrows. References and PDB codes of the structures are shown. (b) Structures of the 80S•IRES complexes in the absence and presence of eEF2 (this work). The large ribosomal subunit is shown in cyan; the small subunit in light yellow (head) and wheat-yellow (body); the TSV IRES in red, eEF2 in green. Nucleotides C1274, U1191 of the 40S head and G904 of the platform (corresponding to C1054, G966 and G693 in E. coli 16S rRNA) are shown in black to denote the A, P and E sites, respectively. Unresolved regions of the IRES in densities for Structures III and V are shown in gray. The extents of the 40S subunit rotation and head swivel relative to their positions in the post-translocation structure are shown with arrows. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.004 Schematic of cryo-EM refinement and classification procedures. All particles were initially aligned to a single model. 3D classification using a 3D mask around the 40S head, TSV IRES and eEF2, of the 4x binned stack was used to identify particles containing both the IRES and eEF2. Subsequent 3D classification using a 2D mask comprising PKI and domain IV of eEF2 yielded 5 'purified' classes representing Structures I through V. Sub-classification of each class did not yield additional classes, but helped improve density in the PKI region of class III (estimated resolution and percentage of particles in the sub-classified reconstruction are shown in parentheses). DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.005 Cryo-EM density of Structures I-V. In panels (a-e), the maps are segmented and colored as in Figure 1. The maps in all panels were B-softened by applying a B-factor of 30 Å2. (a-e) Cryo-EM map of Structures I, II, III, IV and V. (f-j) Local resolution of unfiltered and unmasked cryo-EM reconstructions, assessed using Blocres from the BSoft package, for Structures I, II, III, IV and V. (k-o) Cryo-EM density for the TSV IRES (red model) and eEF2 (green model) in Structures I, II, III, IV and V. (p) Fourier shell correlation (FSC) curves for Structures I-V. The horizontal axis is labeled with spatial frequency Å-1 and with Å. The resolutions stated in the text correspond to an FSC threshold value of 0.143, shown as a dotted line, for the FREALIGN-derived FSC ('Part_FSC'). DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.006 Cryo-EM structures of the 80S•TSV IRES bound with eEF2•GDP•sordarin. (a) Structures I through V. In all panels, the large ribosomal subunit is shown in cyan; the small subunit in light yellow (head) and wheat-yellow (body); the TSV IRES in red, eEF2 in green. Nucleotides C1274, U1191 of the 40S head and G904 of the platform (C1054, G966 and G693 in E. coli 16S rRNA) are shown in black to denote the A, P and E sites, respectively. Unresolved regions of the IRES in densities for Structures III and V are shown in gray. (b) Schematic representation of the structures shown in panel a, denoting the conformations of the small subunit relative to the large subunit. A, P and E sites are shown as rectangles. All measurements are relative to the non-rotated 80S•2tRNA•mRNA structure. The colors are as in panel a. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.002 We sought to address the following questions by structural visualization of 80S•IRES•eEF2 translocation complexes: (1) How does a large IRES RNA move through the restricted intersubunit space, bringing PKI from the A to P site of the small subunit? (2) How does eEF2 mediate IRES translocation? (3) Does IRES translocation involve large rearrangements in the ribosome, similar to tRNA translocation? (4) What, if any, is the mechanistic role of 40S head rotation in IRES translocation? We used cryo-EM to visualize 80S•TSV IRES complexes formed in the presence of eEF2•GTP and the translation inhibitor sordarin, which stabilizes eEF2 on the ribosome. Although the mechanism of sordarin action is not fully understood, the inhibitor does not affect the conformation of eEF2•GDPNP on the ribosome, rendering it an excellent tool in translocation studies. Maximum-likelihood classification using FREALIGN identified five IRES-eEF2-bound ribosome structures within a single sample (Figures 1 and 2). The structures differ in the positions and conformations of ribosomal subunits (Figures 1b and 2), IRES RNA (Figures 3 and 4) and eEF2 (Figures 5 and 6). This ensemble of structures allowed us to reconstruct a sequence of steps in IRES translocation induced by eEF2. Results We used single-particle cryo-EM and maximum-likelihood image classification in FREALIGN to obtain three-dimensional density maps from a single specimen. The translocation complex was formed using S. cerevisiae 80S ribosomes, Taura syndrome virus IRES, and S. cerevisiae eEF2 in the presence of GTP and the eEF2-binding translation inhibitor sordarin. Unsupervised cryo-EM data classification was combined with the use of three-dimensional and two-dimensional masking around the ribosomal A site (Figure 1—figure supplement 2). This approach revealed five 80S•IRES•eEF2•GDP structures at average resolutions of 3.5 to 4.2 Å, sufficient to locate IRES domains and to resolve individual residues in the core regions of the ribosome and eEF2 (Figures 3c,d, and 5f,h; see also Figure 1—figure supplement 2 and Figure 5—figure supplement 2), including the post-translational modification diphthamide 699 (Figure 3c). Large-scale rearrangements in Structures I through V, coupled with the movement of PKI from the A to P site and eEF2 entry into the A site. (a) Rotational states of the 40S subunit in the 80S•IRES structure (INIT; PDB 3J6Y) and in 80S•IRES•eEF2 Structures I, II, III, IV and V (this work). For each structure, the triangle outlines the contours of the 40S body; the lower angle illustrates the extent of intersubunit (body) rotation. The sizes of the arrows correspond to the extent of the head swivel (yellow) and subunit rotation (black). The views were obtained by structural alignment of the 25S rRNAs; the sarcin-ricin loop (SRL) of 25S rRNA is shown in gray for reference. (b) Solvent view (opposite from that shown in (a)) of the 40S subunit in the 80S•IRES structure (INIT; PDB 3J6Y) and in 80S•IRES•eEF2 Structures I, II, III, IV and V (this work). The structures are colored as in Figure 1. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.009 Large-scale rearrangements in Structures I through V, coupled with the movement of PKI from the A to P site and eEF2 entry into the A site. (a) Comparison of the 40S-subunit rotational states in Structures I through V, sampling a ~10° range between Structure I (fully rotated) and Structure V (non-rotated). 18S ribosomal RNA is shown and ribosomal proteins are omitted for clarity. The superpositions of Structures I-V were performed by structural alignments of the 25S ribosomal RNAs. (b) Bar graph of the angles characterizing the 40S rotational and 40S head swiveling states in Structures I through V. Measurements for the two 80S•IRES (INIT) structures are included for comparison. All measurements are relative to the non-rotated 80S•2tRNA•mRNA structure. (c) Comparison of the 40S conformations in Structures I through V shows distinct positions of the head relative to the body of the 40S subunit (head swivel). Conformation of the non-swiveled 40S subunit in the S. cerevisiae 80S ribosome bound with two tRNAs is shown for reference (blue). (d) Comparison of conformations of the L1 and P stalks of the large subunit in Structures I through V with those in the 80S•IRES and tRNA-bound 80S structures. Superpositions were performed by structural alignments of 25S ribosomal RNAs. The central protuberance (CP) is labeled. (e) Bar graph of the positions of PKI and domain IV of eEF2 relative to the P site residues of the head (U1191) and body (C1637) in Structures I through V. (f and g) Close-up view of rearrangements in the A and P sites from the initiation state (INIT: PDB ID 3J6Y) to the post-translocation Structure V. The fragment shown within a rectangle in panel f is magnified in panel g. Nucleotides of the 40S body are shown in orange, 40S head in yellow. The superpositions of structures were performed by structural alignments of the 18S ribosomal RNAs excluding the head region (nt 1150–1620). DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.007 Our structures represent hitherto uncharacterized translocation complexes of the TSV IRES captured within globally distinct 80S conformations (Figures 1b and 2). We numbered the structures from I to V, according to the position of the tRNA-mRNA-like PKI on the 40S subunit (Figure 2—source data 1). Specifically, PKI is partially withdrawn from the A site in Structure I, and fully translocated to the P site in Structure V (Figure 4; see also Figure 3—figure supplement 1). Thus Structures I to IV represent different positions of PKI between the A and P sites (Figure 2—source data 1), suggesting that these structures describe intermediate states of translocation. Structure V corresponds to the post-translocation state. Changes in ribosome conformation and eEF2 positions are coupled with IRES movement through the ribosome Intersubunit rotation Using the post-translocation S. cerevisiae 80S ribosome bound with the P and E site tRNAs as a reference (80S•2tRNA•mRNA), in which both the subunit rotation and the head-body swivel are 0°, we found that the ribosome adopts four globally distinct conformations in Structures I through V (Figure 1b; see also Figure 1—figure supplement 1 and Figure 2—source data 1). Structure I comprises the most rotated ribosome conformation (~10°), characteristic of pre-translocation hybrid-tRNA states. From Structure I to V, the body of the small subunit undergoes backward (reverse) rotation (Figure 2b; see also Figure 1—figure supplement 2 and Figure 2—figure supplement 1). Structures II and III are in mid-rotation conformations (~5°). Structure IV adopts a slightly rotated conformation (~1°). Structure V is in a nearly non-rotated conformation (0.5°), very similar to that of post-translocation ribosome-tRNA complexes. Thus, intersubunit rotation of ~9° from Structure I to V covers a nearly complete range of relative subunit positions, similar to what was reported for tRNA-bound yeast, bacterial and mammalian ribosomes. 40S head swivel The pattern of 40S head swivel between the structures is different from that of intersubunit rotation (Figures 2c and d; see also Figure 2—source data 1). As with the intersubunit rotation, the small head swivel (~1°) in the non-rotated Structure V is closest to that in the 80S•2tRNA•mRNA post-translocation ribosome. However in the pre-translocation intermediates (from Structure I to IV), the beak of the head domain first turns toward the large subunit and then backs off (Figure 2—figure supplement 1). This movement reflects the forward and reverse swivel. The head samples a mid-swiveled position in Structure I (12°), then a highly-swiveled position in Structures II and III (17°) and a less swiveled position in Structure IV (14°). The maximum head swivel is observed in the mid-rotated complexes II and III, in which PKI transitions from the A to P site, while eEF2 occupies the A site partially. By comparison, the similarly mid-rotated (4°) 80S•TSV IRES initiation complex, in the absence of eEF2, adopts a mid-swiveled position (~10°) (Figure 2c). These observations suggest that eEF2 is necessary for inducing or stabilizing the large head swivel of the 40S subunit characteristic for IRES translocation intermediates. IRES rearrangements Comparison of the TSV IRES and eEF2 positions in Structures I through V. (a) Positions of the IRES and eEF2 in the initiation, pre-translocation (I) and post-translocation (V) states, relative to the body of the 40S subunit (not shown) (b) Positions of the IRES and eEF2 in the initiation state (INIT) and intermediate steps of translocation (II, III and IV), relative to the body of the 40S subunit (not shown). Superpositions were obtained by structural alignments of the 18S rRNAs excluding the head domains (nt 1150–1620). DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.011 Positions of the IRES relative to proteins uS7, uS11 and eS25. (a) Intra-IRES rearrangements from the 80S*IRES initiation structure (INIT; PDB 3J6Y,) to Structures I through V. For each structure (shown in red), the conformation from a preceding structure is shown in light red for comparison. Superpositions were obtained by structural alignments of 18S rRNA. (b) Positions of the IRES and eEF2 relative to those of classical P- and E-site tRNAs in the 80S•tRNA complex. (c) Positions of the IRES relative to proteins uS11 (40S platform) and uS7 and eS25 (40S head), which interact with the 5′ domain of the IRES in the initiation state (left panel). In all panels, superpositions were obtained by structural alignments of the 18S rRNAs. Ribosomal proteins of the initiation state are shown in gray for comparison. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.012 Positions of the L1stalk, tRNA and TSV IRES relative to proteins uS7 and eS25, in 80S•tRNA structures and 80S•IRES structures I and V (this work). The view shows the vicinity of the ribosomal E site. Loop 1.1 and stem loops 4 and 5 of the IRES are labeled. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.013 Interactions of the stem loops 4 and 5 of the TSV with proteins uS7 and eS25. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.014 Position and interactions of loop 3 (variable loop region) of the PKI domain in Structure V (this work) resembles those of the anticodon stem loop of the E-site tRNA (blue) in the 80S•2tRNA•mRNA complex. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.015 Positions of tRNAs and the TSV IRES relative to the A-site finger (blue, nt 1008–1043 of 25S rRNA) and the P site of the large subunit, comprising helix 84 of 25S rRNA (nt. 2668–2687) and protein uL5 (collectively labeled as central protuberance, CP, in the upper-row first figure, and individually labeled in the lower-row first figure). Structures of translocation complexes of the bacterial 70S ribosome bound with two tRNAs and yeast 80S complexes with tRNAs are shown in the upper row and labeled. Structures of 80S•IRES complexes in the absence of eEF2 (INIT; PDB 3J6Y,) and in the presence of eEF2 (this work) are shown in the lower row and labeled. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.016 Interactions of the TSV IRES with uL5 and eL42. Structures of 80S•IRES complexes in the absence of eEF2 (INIT; PDB 3J6Y,) and in the presence of eEF2 (this work) are shown in the upper row and labeled. Structures of the 80S complexes with tRNAs are shown in the lower row in a view similar to that for the 80S•IRES complex. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.017 Positions of the IRES relative to eEF2 and elements of the ribosome in Structures I through V. (a) Secondary structure of the TSV IRES. The TSV IRES comprises two domains: the 5' domain (blue) and the PKI domain (red). The open reading frame (gray) is immediately following pseudoknot I (PKI). (b) Three-dimensional structure of the TSV IRES (Structure II). Pseudoknots and stem loops are labeled and colored as in (a). (c) Positions of the IRES and eEF2 on the small subunit in Structures I to V. The initiation-state IRES is shown in gray. The insert shows density for interaction of diphthamide 699 (eEF2; green) with the codon-anticodon-like helix (PKI; red) in Structure V. (d and e) Density of the P site in Structure V shows that interactions of PKI with the 18S rRNA nucleotides (c) are nearly identical to those in the P site of the 2tRNA•mRNA-bound 70S ribosome (d). DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.010 In each structure, the TSV IRES adopts a distinct conformation in the intersubunit space of the ribosome (Figures 3 and 4). The IRES (nt 6758–6952) consists of two globular parts (Figure 3a): the 5’-region (domains I and II, nt 6758–6888) and the PKI domain (domain III, nt 6889–6952). We collectively term domains I and II the 5’ domain. The PKI domain comprises PKI and stem loop 3 (SL3), which stacks on top of the stem of PKI. The 6953GCU triplet immediately following the PKI domain is the first codon of the open reading frame. In the eEF2-free 80S•IRES initiation complex (INIT), the bulk of the 5’-domain (nt. 6758–6888) binds near the E site, contacting the ribosome mostly by means of three protruding structural elements: the L1.1 region and stem loops 4 and 5 (SL4 and SL5). In Structures I to IV, these contacts remain as in the initiation complex (Figure 1a). Specifically, the L1.1 region interacts with the L1 stalk of the large subunit, while SL4 and SL5 bind at the side of the 40S head and interact with proteins uS7, uS11 and eS25 (Figure 3—figure supplement 2 and Figure 3—figure supplement 3; ribosomal proteins are termed according to). In Structures I-IV, the minor groove of SL4 (at nt 6840–6846) binds next to an α-helix of uS7, which is rich in positively charged residues (K212, K213, R219 and K222). The tip of SL4 binds in the vicinity of R157 in the β-hairpin of uS7 and of Y58 in uS11. The minor groove of SL5 (at nt 6862–6868) contacts the positively charged region of eS25 (R49, R58 and R68) (Figure 3—figure supplement 4). In Structure V, however, the density for SL5 is missing suggesting that SL5 is mobile, while weak SL4 density suggests that SL4 is shifted along the surface of uS7, ~20 Å away from its initial position (Figure 3—figure supplement 2c). The L1.1 region remains in contact with the L1 stalk (Figure 3—figure supplement 3). Inchworm-like translocation of the TSV IRES. Conformations and positions of the IRES in the initiation state and in Structures I-V are shown relative to those of the A-, P- and E-site tRNAs. The view was obtained by structural alignment of the body domains of 18S rRNAs of the corresponding 80S structures. Distances between nucleotides 6848 and 6913 in SL4 and PKI, respectively, are shown (see also Figure 2—source data 1). DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.018 The shape of the IRES changes considerably from the initiation state to Structures I through V, from an extended to compact to extended conformation (Figure 4; see also Figure 3—figure supplement 2a). Because in Structures I to IV the PKI domain shifts toward the P site, while the 5’ remains unchanged near the E site, the distance between the domains shortens (Figure 4). In the 80S•IRES initiation state, the A-site-bound PKI is separated from SL4 by almost 50 Å (Figure 4). In Structures I and II, the PKI is partially retracted from the A site and the distance from SL4 shortens to ~35 Å. As PKI moves toward the P site in Structures III and IV, the PKI domain approaches to within ~25 Å of SL4. Because the 5’-domain in the following structure (V) moves by ~20 Å along the 40S head, the IRES returns to an extended conformation (~45 Å) that is similar to that in the 80S•IRES initiation complex. Rearrangements of the IRES involve restructuring of several interactions with the ribosome. In Structure I, SL3 of the PKI domain is positioned between the A-site finger (nt 1008–1043 of 25S rRNA) and the P site of the 60S subunit, comprising helix 84 of 25S rRNA (nt. 2668–2687) and protein uL5 (Figure 3—figure supplement 6). This position of SL3 is ~25 Å away from that in the 80S•IRES initiation state, in which PKI and SL3 closely mimic the ASL and elbow of the A-site tRNA, respectively. As such, the transition from the initiation state to Structure I involves repositioning of SL3 around the A-site finger, resembling the transition between the pre-translocation A/P and A/P* tRNA. The second set of major structural changes involves interaction of the P site region of the large subunit with the hinge point of the IRES bending between the 5´ domain and the PKI domain (nt. 6886–6890). In the highly bent Structures III and IV, the hinge region interacts with the universally conserved uL5 and the C-terminal tail of eL42 (Figure 3—figure supplement 7). However, in the extended conformations, these parts of the IRES and the 60S subunit are separated by more than 10 Å, suggesting that an interaction between them stabilizes the bent conformations but not the extended ones. Another local rearrangement concerns loop 3, also known as the variable loop region , which connects the ASL- and mRNA-like parts of PKI. This loop is poorly resolved in Structures I through IV, suggesting conformational flexibility in agreement with structural studies of the isolated PKI and biochemical studies of unbound IRESs. In Structure V, loop 3 is bound in the 40S E site and the backbone of loop 3 near the codon-like part of PKI (at nt. 6945–6946) interacts with R148 and R157 in β-hairpin of uS7. The interaction of loop 3 backbone with uS7 resembles that of the anticodon-stem loop of E-site tRNA in the post-translocation 80S•2tRNA•mRNA structure (Figure 3—figure supplement 5). Ordering of loop 3 suggests that this flexible region contributes to the stabilization of the PKI domain in the post-translocation state. This interpretation is consistent with the recent observation that alterations in loop 3 of the CrPV IRES result in decreased efficiency of translocation. eEF2 structures Elements of the 80S ribosome that contact eEF2 in Structures I through V. The view and colors are as in Figure 5b: eEF2 is shown in green, IRES RNA in red, 40S subunit elements in orange, 60S in cyan/teal. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.020 Cryo-EM density of the GTPase region in Structures I and II. The switch loop I in Structure I is shown in blue. The putative position of the switch loop I, unresolved in the density of Structure II, is shown with a dashed line. Colors for the ribosome and eEF2 are as in Figure 1. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.021 Conformations and interactions of eEF2. (a) Conformations of eEF2 in Structures I-V and domain organization of eEF2 are shown. Roman numerals denote eEF2 domains. Superposition was obtained by structural alignment of domains I and II. (b) Elements of the 80S ribosome in Structures I and V that contact eEF2. eEF2 is shown in green, IRES RNA in red, 40S subunit elements in orange, 60S in cyan/teal. (c) Comparison of conformations of eEF2•sordarin in Structure I (light green) with those of free apo-eEF2 (magenta) and eEF2•sordarin (teal). (d) Interactions of the GTPase domains with the 40S and 60S subunits in Structure I (colored in green/blue, eEF2; orange, 40S; cyan/teal, 60S) and in Structure II (gray). Switch loop I (SWI) in Structure I is in blue; dashed line shows the putative location of unresolved switch loop I in Structure II. Superposition was obtained by structural alignment of the 25S rRNAs. (e) Comparison of the GTP-like conformation of eEF2•GDP in Structure I (light green) with those of 70S-bound elongation factors EF-Tu•GDPCP (teal) and EF-G•GDP•fusidic acid (magenta; fusidic acid not shown). (f) Cryo-EM density showing guanosine diphosphate bound in the GTPase center (green) next to the sarcin-ricin loop of 25S rRNA (cyan) of Structure II. (g) Comparison of the sordarin-binding sites in the ribosome-bound (light green; Structure II) and isolated eEF2 (teal). (h) Cryo-EM density showing the sordarin-binding pocket of eEF2 (Structure II). Sordarin is shown in pink with oxygen atoms in red. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.019 Elongation factor eEF2 in all five structures is bound with GDP and sordarin (Figure 5). The elongation factor consists of three dynamic superdomains: an N-terminal globular superdomain formed by the G (GTPase) domain (domain I) and domain II; a linker domain III; and a C-terminal superdomain comprising domains IV and V (Figure 5a). Domain IV extends from the main body and is critical for translocation catalyzed by eEF2 or EF-G. ADP-ribosylation of eEF2 at the tip of domain IV or deletion of domain IV from EF-G abrogate translocation. In post-translocation-like 80S•tRNA•eEF2 complexes, domain IV binds in the 40S A site, suggesting direct involvement of domain IV in translocation of tRNA from the A to P site. GDP in our structures is bound in the GTPase center (Figures 5d, e and f) and sordarin is sandwiched between the β-platforms of domains III and V (Figures 5g and h), as in the structure of free eEF2•sordarin complex. The global conformations of eEF2 (Figure 5a) are similar in these structures (all-atom RMSD ≤ 2 Å), but the positions of eEF2 relative to the 40S subunit differ substantially as a result of 40S subunit rotation (Figure 2—source data 1). From Structure I to V, eEF2 is rigidly attached to the GTPase-associated center of the 60S subunit. The GTPase-associated center comprises the P stalk (L11 and L7/L12 stalk in bacteria) and the sarcin-ricin loop (SRL, nt 3012–3042). The tips of 25S rRNA helices 43 and 44 of the P stalk (nucleotides G1242 and A1270, respectively) stack on V754 and Y744 of domain V. An αββ motif of the eukaryote-specific protein P0 (aa 126–154) packs in the crevice between the long α-helix D (aa 172–188) of the GTPase domain and the β-sheet region (aa 246–263) of the GTPase domain insert (or G’ insert) of eEF2 (secondary-structure nomenclatures for eEF2 and EF-G are the same). Although the P/L11 stalk is known to be dynamic, its position remains unchanged from Structure I to V: all-atom root-mean-square differences for the 25S rRNA of the P stalk (nt 1223–1286) are within 2.5 Å. However, with respect to its position in the 80S•IRES complex in the absence of eEF2 and in the 80S•2tRNA•mRNA complex, the P stalk is shifted by ~13 Å toward the A site (Figure 2d). The sarcin-ricin loop interacts with the GTP-binding site of eEF2 (Figures 5d and f). While the overall mode of this interaction is similar to that seen in 70S•EF-G crystal structures, there is an important local difference between Structure I and Structures II-V in switch loop I, as discussed below. Repositioning (sliding) of the positively-charged cluster of domain IV of eEF2 over the phosphate backbone (red) of the 18S helices 33 and 34. Structures I through V are shown. Electrostatic surface of eEF2 is shown; negatively and positively charged regions are shown in red and blue, respectively. The view was obtained by structural alignment of the 18S rRNAs. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.023 Interactions of eEF2 with the 40S subunit. (a) eEF2 (green) interacts only with the body in Structure I (eEF2 domains are labeled with roman numerals in white), and with both the head and body in Structures II through V. Colors are as in Figure 1, except for the 40S structural elements that contact eEF2, which are labeled and shown in purple. (b) Entry of eEF2 into the 40S A site, from Structure I through V. Distances to the A-site accommodated eEF2 (Structure V) are shown. The view was obtained by superpositions of the body domains of 18S rRNAs. (c) Rearrangements, from Structure I through V, of a positively charged cluster of eEF2 (K613, R617 and R631) positioned over the phosphate backbone of 18S helices 33 and 34, suggesting a role of electrostatic interactions in eEF2 diffusion over the 40S surface. (d) Shift of the tip of domain III of eEF2, interacting with uS12 upon reverse subunit rotation from Structure I to Structure V. Structure I colored as in Figure 1, except uS12, which is in purple; Structure V is in gray. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.022 There are two modest but noticeable domain rearrangements between Structures I and V. Unlike in free eEF2, which can sample large movements of at least 50 Å of the C-terminal superdomain relative to the N-terminal superdomain (Figure 5c), eEF2 undergoes moderate repositioning of domain IV (~3 Å; Figure 5a) and domain III (~5 Å; Figure 6d). This limited flexibility of the ribosome-bound eEF2 is likely the result of simultaneous fixation of eEF2 superdomains, via domains I and V, by the GTPase-associated center of the large subunit. Domain IV of eEF2 binds at the 40S A site in Structures I to V but the mode of interaction differs in each complex (Figure 6). Because eEF2 is rigidly attached to the 60S subunit and does not undergo large inter-subunit rearrangements, gradual entry of domain IV into the A site between Structures I and V is due to 40S subunit rotation and head swivel. eEF2 settles into the A site from Structure I to V, as the tip of domain IV shifts by ~10 Å relative to the body and by ~20 Å relative to the swiveling head. Modest intra-eEF2 shifts of domain IV between Structures I to V outline a stochastic trajectory (Figure 5a), consistent with local adjustments of the domain in the A site. At the central region of eEF2, domains II and III contact the 40S body (mainly at nucleotides 48–52 and 429–432 of 18S rRNA helix 5 and uS12). From Structure I to V, these central domains migrate by ~10 Å along the 40S surface (Figure 6c). Comparison of eEF2 conformations reveals that in Structure V, domain III is displaced as a result of interaction with uS12, as discussed below. In summary, between Structures I and V, a step-wise translocation of PKI by ~15 Å from the A to P site - within the 40S subunit – occurs simultaneously with the ~11 Å side-way entry of domain IV into the A site coupled with ~3 to 5 Å inter-domain rearrangements in eEF2. These shifts occur during the reverse rotation of the 40S body coupled with the forward-then-reverse head swivel. To elucidate the detailed structural mechanism of IRES translocation and the roles of eEF2 and ribosome rearrangements, we describe in the following sections the interactions of PKI and eEF2 with the ribosomal A and P sites in Structures I through V (Figure 2g; see also Figure 1—figure supplement 1). Structure I represents a pre-translocation IRES and initial entry of eEF2 in a GTP-like state In the fully rotated Structure I, PKI is shifted toward the P site by ~3 Å relative to its position in the initiation complex but maintains interactions with the partially swiveled head. At the head, C1274 of the 18S rRNA (C1054 in E. coli) base pairs with the first nucleotide of the ORF immediately downstream of PKI. The C1274:G6953 base pair provides a stacking platform for the codon-anticodon–like helix of PKI. We therefore define C1274 as the foundation of the 'head A site'. Accordingly, we use U1191 (G966 in E. coli) and C1637 (C1400 in E. coli) as the reference points of the 'head P site' and 'body P site' (Figure 2g), respectively, because these nucleotides form a stacking foundation for the fully translocated mRNA-tRNA helix in tRNA-bound structures and in our post-translocation Structure V discussed below. Interactions of the residues at the eEF2 tip with the decoding center of the IRES-bound ribosome. Key elements of the decoding center of the 'locked' initiation structure, 'unlocked' Structure I, and post-translocation Structure V (this work) are shown. The histidine-diphthamide tip of eEF2 is shown in green. The codon-anticodon-like helix of PKI is shown in red, the downstream first codon of the ORF in magenta. Nucleotides of the 18S rRNA body are in orange and head in yellow; 25S rRNA nucleotide A2256 is blue. A and P sites are schematically demarcated by dotted lines. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.024 The interaction of PKI with the 40S body is substantially rearranged relative to that in the initiation state. In the latter, PKI is stabilized by interactions with the universally conserved decoding-center nucleotides G577, A1755 and A1756 ('body A site'), as in the A-site tRNA bound complexes. In Structure I, PKI does not contact these nucleotides (Figures 2g and 7). The position of eEF2 on the 40S subunit of Structure I is markedly distinct from those in Structures II to V. The translocase interacts with the 40S body but does not contact the head (Figures 5b and 6a; Figure 5—figure supplement 1). Domain IV is partially engaged with the body A site. The tip of domain IV is wedged between PKI and decoding-center nucleotides A1755 and A1756, which are bulged out of h44. This tip contains the histidine-diphthamide triad (H583, H694 and Diph699), which interacts with the codon-anticodon-like helix of PKI and A1756 (Figure 7). Histidines 583 and 694 interact with the phosphate backbone of the anticodon-like strand (at G6907 and C6908). Diphthamide is a unique posttranslational modification conserved in archaeal and eukaryotic EF2 (at residue 699 in S. cerevisiae) and involves addition of a ~7-Å long 3-carboxyamido-3-(trimethylamino)-propyl moiety to the histidine imidazole ring at CE1. The trimethylamino end of Diph699 packs over A1756 (Figure 7). The opposite surface of the tail is oriented toward the minor-groove side of the second base pair of the codon-anticodon helix (G6906:C6951). Thus, in comparison with the initiation state, the histidine-diphthamide tip of eEF2 replaces the codon-anticodon–like helix of PKI. The splitting of the interaction of A1755-A1756 and PKI is achieved by providing the histidine-diphthamine tip as a binding partner for both A1756 and the minor groove of the codon-anticodon helix (Figure 7). Unlike in Structures II to V, the conformation of the eEF2 GTPase center in Structure I resembles that of a GTP-bound translocase (Figure 5e). In translational GTPases, switch loops I and II are involved in the GTPase activity (reviewed in). Switch loop II (aa 105–110), which carries the catalytic H108 (H92 in E. coli EF-G; is well resolved in all five structures. The histidine resides next to the backbone of G3028 of the sarcin-ricin loop and near the diphosphate of GDP (Figure 5e). By contrast, switch loop I (aa 50–70 in S. cerevisiae eEF2) is resolved only in Structure I (Figure 5—figure supplement 2). The N-terminal part of the loop (aa 50–60) is sandwiched between the tip of helix 14 (415CAAA418) of the 18S rRNA of the 40S subunit and helix A (aa 32–42) of eEF2 (Figure 5d). Bulged A416 interacts with the switch loop in the vicinity of D53. Next to GDP, the C-terminal part of the switch loop (aa 61–67) adopts a helical fold. As such, the conformations of SWI and the GTPase center in general are similar to those observed in ribosome-bound EF-Tu and EF-G in the presence of GTP analogs. Structure II reveals PKI between the body A and P sites and eEF2 partially advanced into the A site In Structure II, relative to Structure I, PKI is further shifted along the 40S body, traversing ~4 Å toward the P site (Figures 2e, f, and g), while stacking on C1274 at the head A site. Thus, the intermediate position of PKI is possible due to a large swivel of the head relative to the body, which brings the head A site close to the body P site. Domain IV of eEF2 is further entrenched in the A site by ~3 Å relative to the body and ~8 Å relative to the head, preserving its interactions with PKI. The decoding center residues A1755 and A1756 are rearranged to pack inside helix 44, making room for eEF2. This conformation of decoding center residues is also observed in the absence of A-site ligands. The head interface of domain IV interacts with the 40S head (Figure 6a). Here, a positively charged surface of eEF2, formed by K613, R617 and R631 contacts the phosphate backbone of helix 33 (Figures 6c; see also Figure 6—figure supplement 1). Structure III represents a highly bent IRES with PKI captured between the head A and P sites Consistent with the similar head swivels in Structure III and Structure II, relative positions of the 40S head A site and body P site remain as in Structure II. Among the five structures, the PKI domain is least ordered in Structure III and lacks density for SL3. The map allows placement of PKI at the body P site (Figure 1—figure supplement 3). Thus, in Structure III, PKI has translocated along the 40S body, but the head remains fully swiveled so that PKI is between the head A and P sites. Lower resolution of the map in this region suggests that PKI is somewhat destabilized in the vicinity of the body P site in the absence of stacking with the foundations of the head A site (C1274) or P site (U1191). The position of eEF2 is similar to that in Structure II. Structure IV represents a highly bent IRES with PKI partially accommodated in the P site In Structure IV, the 40S subunit is almost non-rotated relative to the 60S subunit, and the 40S head is mid-swiveled. Unwinding of the head moves the head P-site residue U1191 and body P-site residue C1637 closer together, resulting in a partially restored 40S P site. Whereas C1637 forms a stacking platform for the last base pair of PKI, U1191 does not yet stack on PKI because the head remains partially swiveled. This renders PKI partially accommodated in the P site (Figure 2g). Unwinding of the 40S head also positions the head A site closer to the body A site. This results in rearrangements of eEF2 interactions with the head, allowing eEF2 to advance further into the A site. To this end, the head-interacting interface of domain IV slides along the surface of the head by 5 Å. Helix A of domain IV is positioned next to the backbone of h34, with positively charged residues K613, R617 and R631 rearranged from the backbone of h33 (Figure 6c; see also Figure 6—figure supplement 1). Structure V represents an extended IRES with PKI fully accommodated in the P site and domain IV of eEF2 in the A site In the nearly non-rotated and non-swiveled ribosome conformation in Structure V closely resembling that of the post-translocation 80S•2tRNA•mRNA complex, PKI is fully accommodated in the P site. The codon-anticodon–like helix is stacked on P-site residues U1191 and C1637 (Figure 3d), analogous to stacking of the tRNA-mRNA helix (Figure 3e). A notable conformational change in eEF2 from that in the preceding Structures is visible in the position of domain III, which contacts uS12 (Figure 6d). In Structure V, protein uS12 is shifted along with the 40S body as a result of intersubunit rotation. In this position, uS12 forms extensive interactions with eEF2 domains II and III. Specifically, the C-terminal tail of uS12 packs against the β-barrel of domain II, while the β-barrel of uS12 packs against helix A of domain III. This shifts the tip of helix A of domain III (at aa 500) by ~5 Å (relative to that in Structure I) toward domain I. Although domain III remains in contact with domain V, the shift occurs in the direction that could eventually disconnect the β-platforms of these domains. Domain IV of eEF2 is fully accommodated in the A site. The first codon of the open reading frame is also positioned in the A site, with bases exposed toward eEF2 (Figure 7), resembling the conformations of the A-site codons in EF-G-bound 70S complexes. As in the preceding Structures, the histidine-diphthamide tip is bound in the minor groove of the P-site codon-anticodon helix. Diph699 slightly rearranges, relative to that in Structure I (Figure 7), and interacts with four out of six codon-anticodon nucleotides. The imidazole moiety stacks on G6907 (corresponding to nt 36 in the tRNA anticodon) and hydrogen bonds with O2’ of G6906 (nt 35 of tRNA). The amide at the diphthamide end interacts with N2 of G6906 and O2 and O2’ of C6951 (corresponding to nt 2 of the codon). The trimethylamino-group is positioned over the ribose of C6952 (codon nt 3). Discussion IRES translocation mechanism Animation showing the transition from the initiation 80S•TSV IRES structures (Koh et al., 2014) to eEF2-bound Structures I through V (this work). Four views (scenes) are shown: (1) A view down the intersubunit space, with the head of the 40S subunit oriented toward a viewer, as in Figure 1a; (2) A view at the solvent side of the 40S subunit, with the 40S head shown at the top, as in Figure 2—figure supplement 1; (3) A view down at the subunit interface of the 40S subunit; (4) A close-up view of the decoding center (A site) and the P site, as in Figure 2g. Each scene is shown twice. Colors are as in Figure 1. In scenes 1, 2 and 3, nucleotides C1274, U1191 of the 40S head and G904 of the 40S platform are shown in black to denote the A, P and E sites, respectively. In scene 4, C1274 and U1191 are labeled and shown in yellow; G577, A1755 and A1756 of the 40S body A site and C1637 of the body P site are labeled and shown in orange. DOI:\nhttp://dx.doi.org/10.7554/eLife.14874.025 In this work we have captured the structures of the TSV IRES, whose PKI samples positions between the A and P sites (Structures I–IV), as well as in the P site (Structure V). We propose that together with the previously reported initiation state, these structures represent the trajectory of eEF2-induced IRES translocation (shown as an animation in http://labs.umassmed.edu/korostelevlab/msc/iresmovie.gif and Video 1). Our structures reveal previously unseen intermediate states of eEF2 or EF-G engagement with the A site, providing the structural basis for the mechanism of translocase action. Furthermore, they provide insight into the mechanism of eEF2•GTP association with the pre-translocation ribosome and eEF2•GDP dissociation from the post-translocation ribosome, also delineating the mechanism of translation inhibition by the antifungal drug sordarin. In summary, the reported ensemble of structures substantially enhances our understanding of the translocation mechanism, including that of tRNAs as discussed below. Translocation of the TSV IRES on the 40S subunit globally resembles a step of an inchworm (Figure 4; see also Figure 3—figure supplement 2). At the start (initiation state), the IRES adopts an extended conformation (extended inchworm). The front 'legs' (SL4 and SL5) of the 5’-domain (front end) are attached to the 40S head proteins uS7, uS11 and eS25 (Figure 3—figure supplement 2). PKI, representing the hind end, is bound in the A site. In the first sub-step (Structures I to IV), the hind end advances from the A to the P site and approaches the front end, which remains attached to the 40S surface. This shortens the distance between PKI and SL4 by up to 20 Å relative to the initiating IRES structure, resulting in a bent IRES conformation (bent inchworm). Finally (Structures IV to V), as the hind end is accommodated in the P site, the front 'legs' advance by departing from their initial binding sites. This converts the IRES into an extended conformation, rendering the inchworm prepared for the next translocation step. Notably, at all steps, the head of the IRES inchworm (L1.1 region) is supported by the mobile L1 stalk. In the post-translocation CrPV IRES structure, the 5’-domain similarly protrudes between the subunits and interacts with the L1 stalk, as in the initiation state for this IRES. This underlines structural similarity for the TSV and CrPV IRES translocation mechanisms. Upon translocation, the GCU start codon is positioned in the A site (Structure V), ready for interaction with Ala-tRNAAla upon eEF2 departure. Recent studies have shown that in some cases a fraction of IGR IRES-driven translation results from an alternative reading frame, which is shifted by one nucleotide relative to the normal ORF. One of the mechanistic scenarios (discussed in) involves binding of the first aminoacyl-tRNA to the post-translocated IRES mRNA frame shifted by one nucleotide (predominantly a +1 frame shift). In our structures, the IRES presents to the decoding center a pre-translocated or fully translocated ORF, rather than a +1 (more translocated) ORF, suggesting that eEF2 does not induce a highly populated fraction of +1 shifted IRES mRNAs. It is likely that alternative frame setting occurs following eEF2 release and that this depends on transient displacement of the start codon in the decoding center, allowing binding of the corresponding amino acyl-tRNA to an off-frame codon. Further structural studies involving 80S•IRES•tRNA complexes are necessary to understand the mechanisms underlying alternative reading frame selection. The presence of several translocation complexes in a single sample suggests that the structures represent equilibrium states of forward and reverse translocation of the IRES, which interconvert among each other. This is consistent with the observations that the intergenic IRESs are prone to reverse translocation. Specifically, biochemical toe-printing studies in the presence of eEF2•GTP identified IRES in a non-translocated position unless eEF1a•aa-tRNA is also present. These findings indicate that IRES translocation by eEF2 is futile: the IRES returns to the A site upon releasing eEF2•GDP unless an amino-acyl tRNA enters the A site and blocks IRES back-translocation. This contrasts with the post-translocated 2tRNA•mRNA complex, in which the classical P and E-site tRNAs are stabilized in the non-rotated ribosome after translocase release. Thus, the meta-stability of the post-translocation IRES is likely due to the absence of stabilizing structural features present in the 2tRNA•mRNA complex. In the initiation state, the IRES resembles a pre-translocation 2tRNA•mRNA complex reduced to the A/P-tRNA anticodon-stem loop and elbow in the A site and the P/E-tRNA elbow contacting the L1 stalk. Because the anticodon-stem loop of the A-tRNA is sufficient for translocation completion, we ascribe the meta-stability of the post-translocation IRES to the absence of the P/E-tRNA elements, either the ASL or the acceptor arm, or both. Furthermore, interactions of SL4 and SL5 with the 40S subunit likely contribute to stabilization of pre-translocation structures. Partitioned roles of 40S subunit rearrangements Our structures delineate the mechanistic functions for intersubunit rotation and head swivel in translocation. These functions are partitioned. Specifically, intersubunit rotation allows eEF2 entry into the A site, while the head swivel mediates PKI translocation. Various degrees of intersubunit rotation have been observed in cryo-EM studies of the 80S•IRES initiation complexes. This suggests that the subunits are capable of spontaneous rotation, as is the case for tRNA-bound pre-translocation complexes. The pre-translocation Structure I with eEF2 least advanced into the A site adopts a fully rotated conformation. Reverse intersubunit rotation from Structure I to V shifts the translocation tunnel (the tunnel between the A, P and E sites) toward eEF2, which is rigidly attached to the 60S subunit. This allows eEF2 to move into the A site. As such, reverse intersubunit rotation facilitates full docking of eEF2 in the A site. Because the histidine-diphthamide tip of eEF2 (H583, H694 and Diph699) attaches to the codon-anticodon-like helix of PKI, eEF2 appears to directly force PKI out of the A site. The head swivel allows gradual translocation of PKI to the P site, first with respect to the body and then to the head. The fully swiveled conformations of Structures II and III represent the mid-point of translocation, in which PKI relocates between the head A site and body P site. We note that such mid-states have not been observed for 2tRNA•mRNA, but their formation can explain the formation of subsequent pe/E hybrid and ap/P chimeric structures (Figure 1—figure supplement 1). Reverse swivel from Structure III to V brings the head to the non-swiveled position, restoring the A and P sites on the small subunit. The functions of eEF2 in translocation To our knowledge, our work provides the first high-resolution view of the dynamics of a ribosomal translocase that is inferred from an ensemble of structures sampled under uniform conditions. The structures, therefore, offer a unique opportunity to address the role of the elongation factors during translocation. Translocases are efficient enzymes. While the ribosome itself has the capacity to translocate in the absence of the translocase, spontaneous translocation is slow. EF-G enhances the translocation rate by several orders of magnitude, aided by an additional 2- to 50-fold boost from GTP hydrolysis. Due to the lack of structures of translocation intermediates, the mechanistic role of eEF2/EF-G is not fully understood. The 80S•IRES•eEF2 structures reported here suggest two main roles for eEF2 in translocation. As discussed above, the first role is to directly shift PKI out of the A site upon spontaneous reverse intersubunit rotation. In our structures, the tip of domain IV docks next to PKI, with diphthamide 699 fit into the minor groove of the codon-anticodon-like helix of PKI (Figure 7). This arrangement rationalizes inactivation of eEF2 by diphtheria toxin, which catalyzes ADP-ribosylation of the diphthamide (reviewed in). The enzyme ADP-ribosylates the NE2 atom of the imidazole ring, which in our structures interacts with the first two residues of the anticodon-like strand of PKI. The bulky ADP-ribosyl moiety at this position would disrupt the interaction, rendering eEF2 unable to bind to the A site and/or stalled on ribosomes in a non-productive conformation. As eEF2 shifts PKI toward the P site in the course of reverse intersubunit rotation, the 60S-attached translocase migrates along the surface of the 40S subunit, guided by electrostatic interactions. Positively-charged patches of domains II and III (R391, K394, R433, R510) and IV (K613, R617, R609, R631, K651) slide over rRNA of the 40S body (h5) and head (h18 and h33/h34), respectively. The Structures reveal hopping of the positive clusters over rRNA helices. For example, between Structures II and V, the K613/R617/R631 cluster of domain IV hops by ~19 Å (for Cα of R617) from the phosphate backbone of h33 (at nt 1261–1264) to that of the neighboring h34 (at nt 1442–1445). Thus, sliding of eEF2 involves reorganization of electrostatic, perhaps isoenergetic interactions, echoing those implied in extraordinarily fast ribosome inactivation rates by the small-protein ribotoxins and in fast protein association and diffusion along DNA. Comparison of our structures with the 80S•IRES initiation structure reveals the structural basis for the second key function of the translocase: 'unlocking' of intrasubunit rearrangements that are required for step-wise translocation of PKI on the small subunit. The unlocking model of the ribosome•2tRNA•mRNA pre-translocation complex has been proposed decades ago and functional requirement of the translocase in this process has been implicated. However, the structural and mechanistic definitions of the locked and unlocked states have remained unclear, ranging from the globally distinct ribosome conformations to unknown local rearrangements, e.g. those in the decoding center. FRET data indicate that translocation of 2tRNA•mRNA on the 70S ribosome requires a forward-and-reverse head swivel, which may be related to the unlocking phenomenon. Whereas intersubunit rotation of the pre-translocation complex occurs spontaneously, the head swivel is induced by the eEF2/EF-G translocase, consistent with requirement of eEF2 for unlocking. Structural studies revealed large head swivels in various 70S•tRNA•EF-G and 80S•tRNA•eEF2 complexes, but not in 'locked' complexes with the A site occupied by the tRNA in the absence of the translocase. Our structures suggest that eEF2 induces head swivel by 'unlocking' the head-body interactions (Figure 7). Binding of the ASL to the A site is known from structural studies of bacterial ribosomes to result in 'domain closure' of the small subunit, i.e. closer association of the head, shoulder and body domains. The domain closure 'locks' cognate tRNA in the A site via stacking on the head A site (C1274 in S. cerevisiae or C1054 in E. coli) and interactions with the body A-site nucleotides A1755 and A1756 (A1492 and A1493 in E. coli). This 'locked' state is identical to that observed for PKI in the 80S•IRES initiation structures in the absence of eEF2. Structure I demonstrates that at an early pre-translocation step, the histidine-diphthamide tip of eEF2 is wedged between A1755 and A1756 and PKI. This destabilization allows PKI to detach from the body A site upon spontaneous reverse 40S body rotation, while maintaining interactions with the head A site. Destabilization of the head-bound PKI at the body A site thus allows mobility of the head relative to the body. The histidine-diphthamide-induced disengagement of PKI from A1755 and A1756 therefore provides the structural definition for the 'unlocking' mode of eEF2 action. In summary, our structures are consistent with a model of eEF2-induced translocation in which both PKI and eEF2 passively migrate into the P and A site, respectively, during spontaneous 40S body rotation and head swivel, the latter being allowed by 'unlocking' of the A site by eEF2. Observation of different PKI conformations sampling a range of positions between the A and P sites in the presence of eEF2•GDP implies that thermal fluctuations of the 40S head domain are sufficient for translocation along the energetically flat trajectory. Insights into eEF2 association with and dissociation from the ribosome The conformational rearrangements in eEF2 from Structure I through Structure V provide insights into the mechanisms of eEF2 association with the pre-translocation ribosome and dissociation from the post-translocation ribosome. In all five structures, the GTPase domain is attached to the P stalk and the sarcin-ricin loop. In the fully-rotated pre-translocation-like Structure I, an additional interaction exists. Here, switch loop I interacts with helix 14 (415CAAA418) of the 18S rRNA. This stabilization renders the GTPase center to adopt a GTP-bound conformation, similar to those observed in other translational GTPases in the presence of GTP analogs and in the 80S•eEF2 complex bound with a transition-state mimic GDP•AlF4–. The switch loop contacts the base of A416 (invariable A344 in E. coli and A463 in H. sapiens). Mutations of residues flanking A344 in E. coli 16S rRNA modestly inhibit translation but do not specifically affect EF-G-mediated translocation. However, the effect of A344 mutation on translation was not addressed in that study, leaving the question open whether this residue is critical for eEF2/EF-G function. The interaction between h14 and switch loop I is not resolved in Structures II to V, in all of which the small subunit is partially rotated or non-rotated, so that helix 14 is placed at least 6 Å farther from eEF2 (Figure 5d). We conclude that unlike other conformations of the ribosome, the fully rotated 40S subunit of the pre-translocation ribosome provides an interaction surface, complementing the P stalk and SRL, for binding of the GTP-bound translocase. This structural basis rationalizes the observation of transient stabilization of the rotated 70S ribosome upon EF-G•GTP binding and prior to translocation. The least rotated conformation of the post-translocation Structure V suggests conformational changes that may trigger eEF2 release from the ribosome at the end of translocation. The most pronounced inter-domain rearrangement in eEF2 involves movement of domain III. In the rotated or mid-rotated Structures I through III, this domain remains rigidly associated with domain V and the N-terminal superdomain and does not undergo noticeable rearrangements. In Structure V, however, the tip of helix A of domain III is displaced toward domain I by ~5 Å relative to that in mid-rotated or fully rotated structures. This displacement is caused by the 8 Å movement of the 40S body protein uS12 upon reverse intersubunit rotation from Structure I to V (Figure 6d). We propose that the shift of domain III by uS12 initiates intra-domain rearrangements in eEF2, which unstack the β-platform of domain III from that of domain V. This would result in a conformation characteristic of free eEF2 and EF-G in which the β-platforms are nearly perpendicular. As we discuss below, Structure V captures a 'pre-unstacking' state due to stabilization of the interface between domains III and V by sordarin. Sordarin stabilizes GDP-bound eEF2 on the ribosome Sordarin is a potent antifungal antibiotic that inhibits translation. Based on biochemical experiments, two alternative mechanisms of action were proposed: sordarin either prevents eEF2 departure by inhibiting GTP hydrolysis or acts after GTP hydrolysis. Although our complex was assembled using eEF2•GTP, density maps clearly show GDP and Mg2+ in each structure (Figure 5g). Our structures therefore indicate that sordarin stalls eEF2 on the ribosome in the GDP-bound form, i.e. following GTP hydrolysis and phosphate release. The mechanism of stalling is suggested by comparison of pre-translocation and post-translocation structures in our ensemble. In all five structures, sordarin is bound between domains III and V of eEF2, stabilized by hydrophobic interactions identical to those in the isolated eEF2•sordarin complex (Figures 5g and h). In the nearly non-rotated post-translocation Structure V, the tip of domain III is shifted, however the interface between domains III and V remains unchanged, suggesting strong stabilization of this interface by sordarin. We note that Structure V is slightly more rotated than the 80S•2tRNA•mRNA complex in the absence of eEF2•sordarin, implying that sordarin interferes with the final stages of reverse rotation of the post-translocation ribosome. We propose that sordarin acts to prevent full reverse rotation and release of eEF2•GDP by stabilizing the interdomain interface and thus blocking uS12-induced disengagement of domain III from domain V. Implications for tRNA and mRNA translocation during translation Because translocation of tRNA must involve large-scale dynamics, this step has long been regarded as the most puzzling step of translation. Intersubunit rearrangements and tRNA hybrid states have been proposed to play key roles half a century ago. Despite an impressive body of biochemical, fluorescence and structural data accumulated since then, translocation remains the least understood step of elongation. The structural understanding of ribosome and tRNA dynamics has been greatly aided by a wealth of X-ray and cryo-EM structures (reviewed in). However, visualization of the eEF2/EF-G-induced translocation is confined to very early pre-EF-G-entry states and late (almost translocated or fully translocated) states, leaving most of the path from the A to the P site uncharacterized (Figure 1—figure supplement 1). Our study provides new insights into the structural understanding of tRNA translocation. First, we propose that tRNA and IRES translocations occur via the same general trajectory. This is evident from the fact that ribosome rearrangements in translocation are inherent to the ribosome and likely occur in similar ways in both cases. Furthermore, the step-wise coupling of ribosome dynamics with IRES translocation is overall consistent with that observed for 2tRNA•mRNA translocation in solution. For example, fluorescence and biochemical studies revealed that the early pre-translocation EF-G-bound ribosomes are fully rotated and translocation of the tRNA-mRNA complex occurs during reverse rotation of the small subunit, coupled with head swivel. The sequence of ribosome rearrangements during IRES translocation also agrees with that inferred from 70S•EF-G structures, including those in which the A-to-P-site translocating tRNA was not present. Specifically, an earlier translocation intermediate ribosome (TIpre) was proposed to adopt a rotated (7–9°) body and a partly rotated head (5–7.5°), in agreement with the conformation of our Structure I. The most swiveled head (18–21°) was observed in a mid-rotated ribosome (3–5°) of a later translocation intermediate TIpost, similar to the conformation of our Structure III. Overall, these correlations suggest that the intermediate locations of the elusive A-to-P-site translocating tRNA are similar to those of PKI in our structures. Second, the structures clarify the structural basis of the often-used but structurally undefined terms 'locking' and 'unlocking' with respect to the pre-translocation complex (Figure 6f). We deem the pre-translocation complex locked, because the A-site bound ASL-mRNA is stabilized by interactions with the decoding center. These interactions are maintained for the classical- and hybrid-state tRNAs in the spontaneously sampled non-rotated and rotated ribosomes, respectively. Unlocking involves separation of the codon-anticodon helix from the decoding center residues by the protruding tip of eEF2/EF-G (Figure 7), occurring in the fully rotated ribosome at an early pre-translocation step. This unlatches the head, allowing creation of hitherto elusive intermediate tRNA positions during spontaneous reverse body rotation. Third, our findings uncover a new role of the head swivel. Previous studies showed that this movement widens the constriction ('gate') between the P and E sites, thus allowing the P-tRNA passage to the E site. In addition to the 'gate-opening' role, we now show that the head swivel brings the head A site to the body P site, allowing a step-wise conveying of the codon-anticodon helix between the A and P sites. Finally, the similar populations of particles (within a 2X range) in our 80S•IRES•eEF2 reconstructions (Figure 1—figure supplement 2) suggest that the intermediate translocation states sample several energetically similar and interconverting conformations. This is consistent with the idea of a rather flat energy landscape of translocation, suggested by recent work that measured mechanical work produced by the ribosome during translocation. Our findings implicate, however, that the energy landscape is not completely flat and contains local minima for transient positions of the codon-anticodon helix between the A and P sites. The shift of the PKI with respect to the body occurs during forward head swivel in two major sub-steps of ~4 Å each (initiation complex to I, and I to II), after which PKI undergoes small shifts to settle in the body P site in Structures III, IV and V (Figure 2—source data 1). Movement of PKI relative to the head occurs during the subsequent reverse swivel in three 3–7 Å sub-steps (II to III to IV to V). It is possible that additional meta-stable but less populated states exist between the conformations we observe. We note that four of our near-atomic resolution maps comprised ~30,000 particles each, the minimum number required for a near-atomic-resolution reconstruction of the ribosome. A larger data set will therefore be necessary to reveal additional sub-states. Concluding remarks Translation of viral mRNA Our work sheds light on the dynamic mechanism of cap-independent translation by IGR IRESs, tightly coupled with the universally conserved dynamic properties of the ribosome. The cryo-EM structures demonstrate that the TSV IRES structurally and dynamically represents a chimera of the 2tRNA•mRNA translocating complex (A/P-tRNA • P/E-tRNA • mRNA). Like in the 2tRNA•mRNA translocating complex in which the two tRNAs move independently of each other, the PKI domain moves relative to the 5´-domain, causing the IRES to undergo an inchworm-walk translocation. A large structural difference between the IRES and the 2tRNA•mRNA complex exists, however, in that the IRES lacks three out of six tRNA-like domains involved in tRNA translocation. This difference likely accounts for the inefficient translocation of the IRES, which is difficult to stabilize in the post-translocation state and therefore is prone to reverse translocation. Although structurally handicapped, the TSV IRES manages to translocate by employing ribosome dynamics that are remarkably similar to that in 2tRNA•mRNA translocation. The uniformity of ribosome dynamics underscores the idea that translocation is an inherent and structurally-optimized property of the ribosome, supported also by translocation activity in the absence of the elongation factor. This property is rendered by the relative mobility of the three major building blocks, the 60S subunit and the 40S head and body, assisted by ligand-interacting extensions including the L1 stalk and the P stalk. Intergenic IRESs, in turn, represent a striking example of convergent molecular evolution. Viral mRNAs have evolved to adopt an atypical structure to employ the inherent ribosome dynamics, to be able to hijack the host translational machinery in a simple fashion. Ensemble cryo-EM Our current understanding of macromolecular machines, such as the ribosome, is often limited by a gap between biophysical/biochemical studies and structural studies. For example, Förster resonance energy transfer can provide insight into the macromolecular dynamics of an assembly at the single-molecule level but is limited to specifically labeled locations within the assembly. High-resolution crystal structures, on the other hand, can provide static images of an assembly, and the structural dynamics can only be inferred by comparing structures that are usually obtained in different experiments and under different, often non-native, conditions. Cryo-EM offers the possibility of obtaining integrated information of both structure and dynamics as demonstrated in lower-resolution studies of bacterial ribosome complexes. Recent technical advances, including direct electron detectors and image processing software, have significantly improved the resolution at which such studies can be performed. The increased resolution, need for larger datasets and more sophisticated algorithms have also led to a massive increase in the computational power required to process the data. The available computing infrastructure and computational efficiency have therefore become deciding factors in how many different structural states can be resolved. This is presumably one of the reasons why most recent studies of ribosome complexes have focused on a single high-resolution structure despite the non-uniform local resolution of the maps that likely reflects structural heterogeneity. The computational efficiency of FREALIGN has allowed us to classify a relatively large dataset (1.1 million particles) into 15 classes (Figure 1—figure supplement 2) and obtain eight near-atomic-resolution structures from it. The classification, which followed an initial alignment of all particles to a single reference, required about 130,000 CPU hours or about five to six full days on a 1000-CPU cluster. While it would clearly be impractical to perform this type of analysis on a desktop computer, one may extrapolate using Moore’s law that such practice will become routine in less than ten years. Therefore, cryo-EM has the potential to become a standard tool for uncovering detailed dynamic pathways of complex macromolecular machines. A particularly exciting application will be to infer the high-resolution temporal trajectory of a pathway from an ensemble of equilibrium states in a single sample, as described in our work, together with an analysis of samples quenched at different time points of the reaction. Materials and methods S. cerevisiae 80S ribosome preparation 80S ribosomes used in this study were prepared from Saccharomyces cerevisiae strain W303 as described previously. To obtain ribosomal subunits, purified 80S was incubated in dissociation buffer (20 mM HEPES·KOH (pH 7.5), 0.5 M KCl, 2.5 mM magnesium acetate, 2 mM dithiothreitol (DTT), and 0.5 U/μl RNasin) for 1 hr at 4°C. The dissociated subunits were then layered on sucrose gradients (10% to 30% sucrose) in the dissociation buffer and centrifuged for 15 hr at 22,000 rpm in an SW32 rotor. Fractions corresponding to 40S and 60S subunits were pooled and buffer-exchanged to subunit storage buffer containing 50 mM Tris (pH7.5), 20 mM MgCl2, 100 mM KCl, and 2 mM DTT. Purified subunits were flash-frozen in liquid nitrogen and stored in aliquots at –80°C. Taura syndrome virus IRES preparation Plasmid pUC57 (Genscript) containing the synthetic DNA encoding for nucleotides 6741–6990 of the TSV mRNA sequence was used to amplify the 250-nucleotide fragment by PCR. This DNA fragment (TSV IRES RNA) served as a template for in vitro transcription. The transcription reaction was incubated for 4 hr at 37°C, and the resulting transcription product was treated with DNase I for 30 mins at 37°C. The RNA was then extracted with acidic phenol/chloroform, gel-purified, and then ethanol precipitated with 100% ethanol, followed by an 80% ethanol wash. The resulting RNA pellet was air-dried at room temperature and suspended in RNase-free water. The TSV IRES transcription product was folded in 1X IRES refolding buffer (20 mM Potassium acetate pH 7.5 and 5 mM MgCl2), incubated at 65°C for 10 min and cooled down gradually at room temperature, prior to complex preparation for cryo-EM study. S. cerevisiae eEF2 purification C-terminally His6-tagged eEF2 was produced in yeast TKY675 cells obtained from Terri Goss Kinzy. Yeast cells were grown in 4 liters of YPD media at 27°C and 160 rpm, to A600=1.5. Yeast cell pellet (~5 g) was obtained by centrifugation and re-suspended in 20 ml of the lysis buffer (50 mM potassium phosphate pH 7.6, 1 M KCl, 1% Tween 20, 10% Glycerol, 10 mM imidazole, 0.2 mM PMSF, 1 mM DTT, and 1 tablet of Roche miniComplete protease inhibitor). The suspension was lysed with microfluidizer at 25,000 psi at 4°C, and then clarified by centrifugation twice at 30,000 × g for 20 min. The supernatant was subjected to Ni-NTA affinity chromatography using the AKTAexplorer 100 system (GE Healthcare). After lysate application onto the column, the column was washed with a five-column volume of wash buffer (50 mM potassium phosphate pH 7.6, 1 M KCl, 1% Tween 20, 10% Glycerol, 20 mM imidazole, 0.2 mM PMSF and 1 mM DTT). A gradient elution method was used to reach the final imidazole concentration of 250 mM. Eluted fractions were buffer-exchanged into buffer A (30 mM HEPES·KOH (pH 7.5), 5% glycerol, 65 mM ammonium chloride, 7 mM β–mercaptoethanol and 1 tablet of miniComplete protease inhibitor) for HiTrap SP Sepharose High Performance cation-exchange chromatography (GE Healthcare). A gradient elution method was used to reach the final salt concentration of 1 M KCl in buffer A over the 20-column volume (100 ml). The peak fraction was concentrated and buffer-exchanged into buffer A, which is also the buffer used for the subsequent size-exclusion chromatography employing Superdex 200 (GE Healthcare). Fractions corresponding to the eEF2 peak were concentrated and stored in aliquots at -20°C. 80S•TSV IRES•eEF2•GTP•sordarin complex preparation The IRES-eEF2-ribosome complex was assembled in two steps. First, refolded TSV IRES RNA (8 μM - all concentrations are specified for the final complex) was incubated with the yeast 40S small subunit (0.8 μM) for 15 min at 30°C, in the buffer containing 45 mM HEPES·KOH (pH 7.5), 10 mM MgCl2, 100 mM KCl, 2.5 mM spermine and 2 mM β–mercaptoethanol. The 60S subunit (0.8 μM) was then added and incubated for 15 min at 30°C. Subsequently, eEF2 (5 μM), sordarin (800 μM) and GTP (1 mM) were added and incubated for 15 min at 30°C. The solution was then incubated on ice for 10 min and flash-frozen in liquid nitrogen. Cryo-EM specimen preparation Quantifoil Cu 200 mesh grids (SPI Supplies, West Chester, PA) were coated with a thin layer of carbon and glow discharged for 45 s at 25 mA. 3 µL of sample with a concentration of ~0.1 µM was applied to the grid, incubated for 30 s and plunged into liquid ethane using an FEI Vitrobot Mark 2 (FEI Company, Hillsboro, OR) after blotting for 3 s at 4°C and ~85% relative humidity. Electron microscopy Cryo-EM data were collected in movie mode on an FEI Krios microscope (FEI Company, Hillsboro, OR) operating at 300 kV and equipped with a K2 Summit direct detector (Gatan Inc., Pleasanton, CA) operating in super-resolution mode with pixel size of 0.82 Å per super-resolution pixel. Each movie consisted of 50 frames collected over 18.8 s with an exposure per frame of 1.4 e-/Å2 as shown by Digital Micrograph (Gatan Inc., Pleasanton, CA), giving a total exposure of 70 e-/Å2. The defocus ranged between ~0.7 to ~2.5 µm underfocus. Image processing The gain-corrected super-resolution movie frames were corrected for anisotropic magnification using bilinear interpolation. The frames were downsampled by Fourier cropping to a pixel size of 1.64 Å. The downsampled frames were then motion-corrected and exposure filtered using Unblur. The image defocus was determined using CTFFIND4 on non-exposure-filtered images and images with excessive motion, low contrast, ice contamination or poor power spectra were removed based on visual inspection using TIGRIS (http://tigris.sourceforge.net/). 50 particles were picked manually using TIGRIS, summed and rotationally averaged to serve as a reference for correlation-based particle picking in IMAGIC. 1,105,737 two-dimensional images of ribosomes (termed 'particles') were picked automatically, extracted into 256 x 256 boxes and converted to MRC/CCP4 format with a corresponding list of micrograph numbers and defocus values for input to FREALIGN v9. The summary of procedures resulting in 3D cryo-EM maps is presented on Figure 1—figure supplement 2. FREALIGN v9 was used for refinement, classification and 3D reconstruction of all ribosome structures. Initial particle alignments were obtained by performing an angular grid search (FREALIGN mode 3) with a density map calculated from the atomic model of the non-rotated 80S ribosome bound with 2 tRNAs (PDB: 3J78). For this search, the resolution was limited to 20 Å and the resolution of the resulting reconstruction was 3.6 Å, as determined by the FSC = 0.143 threshold criterion. Four additional rounds of mode 3 with the resolution limited to 7 Å improved the resolution of the reconstruction to 3.5 Å. Starting with cycle 6, particles were classified into six classes using 21 rounds of mode 1 refinement. Inspection of the six classes suggested that several represented mixed conformations. The alignment parameters of the class containing the largest number of particles (25%) were therefore used to initialize classification into 15 classes. For this classification, particle images were downsampled by Fourier cropping to a pixel size of 3.28 Å to accelerate processing. 99 rounds of refinement and classification were performed using mode 1 with a resolution limit of 7 Å. To help separate different conformations affecting small subunit, IRES and eEF2, we used a 3D mask that included density belonging to these parts of the structure. This mask was applied in every cycle to the 3D reference structures prior to refinement and classification in 42 additional cycles. The mask was then changed to include only the head of the small subunit, IRES and eEF2, and a final 18 cycles of refinement and classification were run. We selected six out of the 15 final classes based on clear density present for IRES and eEF2 and continued all further processing with this subset of the data (312,698 particles). The six classes were grouped into three groups based on the rotational state of the small subunit, and each group was further refined and classified using between six and 36 cycles of mode 1 and particles downsampled to 1.64 Å pixel size. For this classification, FREALIGN’s focused mask feature was used to select either the region of IRES PKI (for classes showing intermediate rotation of the small subunit) or a region containing both IRES PKI and eEF2 domain 4 (for classes showing no rotation of the small subunit). This refinement and sub-classification produced eight new classes with more distinct features in the regions selected by the focused masks. These eight classes were used as starting references for a final 33 cycles of refinement and classification using mode 1 and focused mask with the radius of 45 Å covering the vicinity of the ribosomal A site. The first 26 cycles were performed using particles downsampled to 3.28 Å pixel size, followed by two cycles at a pixel size of 1.64 Å, and five cycles at a pixel size of 0.82 Å. The resolution limit for the final cycles was set at 5 Å. The resulting eight reconstructions were used for further analyses, model building and structural refinements, as described below. In parallel, to enhance resolution of the IRES 5´ domain, we performed classification and refinement of the eight classes using a mask with the radius of 50 Å covering the vicinity of the E site and L1 stalk; these maps were used for model building and confirmation of the IRES 5´ domain structure, but not for structure refinements. Among the resulting eight reconstructions, four reconstructions contained well defined PKI and eEF2 densities (I, II, IV and V) (Figure 1—figure supplement 1). PKI was poorly resolved in reconstruction III. Reconstruction VI represents the previously reported 80S•TSV IRES initiation complex in the least rotated conformation. Reconstructions VII and VIII correspond to ribosomes adopting intermediate rotational states, similar to that of Structure III, with weak density in the region of the 5’ domain of the IRES and no density for the PKI domain. To resolve heterogeneity of PKI in reconstruction III, we performed additional sub-classification of all eight classes into two or three classes each. This sub-classification did not result into different structures for the four classes of interest (I, II, IV and V), suggesting a high degree of homogeneity in the masked regions of PKI and eEF2 domain IV. Sub-classification of reconstruction III helped improve the PKI density, resulting in a 4.2 Å reconstruction. All maps were subsequently B-factor-filtered by bfactor.exe, using the B-factors of -50 to -120 Å2, as suggested by bfactor.exe for individual maps, and used for real-space structure refinements. FSC curves (Figure 1—figure supplement 3) were calculated by FREALIGN for even and odd particles half-sets. Blocres was used to calculate the local resolution of unfiltered and unmasked volumes using a box size 60 pixel, step size of 3 pixels and FSC resolution criterion (threshold 0.143). The output volumes were then colored according to the local resolution of the final reconstructions (Figure 1—figure supplement 3) using the Surface Color tool of Chimera  Model building and refinement The starting structural models were assembled using the high-resolution crystal structure of S. cerevisiae 80S ribosome, the cryo-EM structure of the 80S•TSV IRES complex and the crystal structure of the isolated eEF2•sordarin complex. The structure of the diphthamide residue of eEF2 (699) was adopted from PDB: 1ZM4. Initial domain fitting into the cryo-EM maps was performed using Chimera, followed by manual modeling of local regions into the density using Pymol and Coot. Parts of several ribosomal proteins were modeled using I-TASSER  and Phyre2 . The structural models were refined by real-space simulated-annealing refinement using atomic electron scattering factors, employing RSRef as described. Secondary-structure restrains for ribosomal proteins and base-pairing restraints for RNA molecules were employed, as described. The refined structural models closely agree with the corresponding maps, as shown by low real-space R-factors of ~0.2 to 0.27, and they have good stereochemical parameters, characterized by low deviation from ideal bond lengths and angles (Figure 1—source data 1). The maps revealed regions, which are differently resolved in Structures I to V. The most prominent difference is in the platform subdomain of the 40S subunit, which is well resolved in Structures I, IV and V but poorly resolved in Structures II and III. The following components of the 40S platform in Structures II and III lacked resolution: proteins eS1, uS11, eS26 and eL41, 18S rRNA nt 892–900, 900–918 and the 3´ end beyond nt 1792. These and other regions of low density were modeled as protein or RNA backbone. For structural comparisons, the distances and angles were calculated in Pymol and Chimera, respectively. To calculate an angle of the 40S subunit rotation between two 80S structures, the 25S rRNAs were aligned using Pymol, and the angle between 18S rRNAs was measured in Chimera. To calculate an angle of the 40S-head rotation (swivel) between two 80S structures, the 18S rRNAs of the bulk of the 40S body (18S nucleotides excluding nt 1150–1620) were aligned using Pymol, and the angle between the 18S rRNA residues 1150–1620 was measured in Chimera. Figures were prepared in Pymol and Chimera. Funding Information This paper was supported by the following grants:   to Nikolaus Grigorieff.  to Nikolaus Grigorieff.   to Andrei A Korostelev.   to Andrei A Korostelev. Additional information Competing interests NG: Reviewing editor, eLife. The other authors declare that no competing interests exist. Author contributions PDA, Collected and analyzed cryo-EM data, Drafting or revising the article. CSK, Prepared the ribosome•IRES•eEF2 complex, Built and refined structural models, Analysis and interpretation of data, Drafting or revising the article. TG, Assisted with cryo-EM data processing and analyses, Drafting or revising the article. NG, Designed the project, Assisted with cryo-EM data processing and analyses, Drafting or revising the article. AAK, Designed the project, Built and refined structural models, Analysis and interpretation of data, Drafting or revising the article. Additional files Major datasets The following datasets were generated: References Three-dimensional structure of the ribosomal translocase: elongation factor G from Thermus thermophilus Visualization of the hybrid state of trna binding promoted by spontaneous ratcheting of the ribosome Structural insights into trna dynamics on the ribosome Insights into factorless translational initiation by the trna-like pseudoknot domain of a viral IRES Ribosome structures to near-atomic resolution from thirty thousand cryo-em particles A new system for naming ribosomal proteins The structure of the eukaryotic ribosome at 3.0 A resolution Interactions of adenosine diphosphate-ribosylated elongation factor 2 with ribosomes tRNA dynamics on the ribosome during translation Translocation in protein synthesis: A hybrid structure model Structure of the ribosome with elongation factor G trapped in the pretranslocation state Structure and dynamics of the mammalian ribosomal pretranslocation complex One number does not fit all: Mapping local variations in resolution in cryo-em reconstructions Factor-independent assembly of elongation-competent ribosomes by an internal ribosome entry site located in an RNA virus that infects penaeid shrimp Restrained real-space macromolecular atomic refinement using a new resolution-dependent electron-density function Structural dynamics of ribosome subunit association studied by mixing-spraying time-resolved cryogenic electron microscopy Coordinated conformational and compositional dynamics drive ribosome translocation Structure of ef-g–ribosome complex in a pretranslocation state A primer to single-particle cryo-electron microscopy Understanding the mode of action of diphtheria toxin: A perspective on progress during the 20th century Initiation of translation in bacteria by a structured eukaryotic IRES RNA Spontaneous intersubunit rotation in single ribosomes tRNA-mrna mimicry drives translation initiation from a viral IRES Ribosomal proteins S12 and S13 function as control elements for translocation of the mrna:trna complex Dual use of GTP hydrolysis by elongation factor G on the ribosome The crystal structure of elongation factor G complexed with GDP, at 2.7 A resolution ADP-ribosylated elongation factor 2 (adp-ribosyl-ef-2) is unable to promote translocation within the ribosome mRNAs that specifically interact with eukaryotic ribosomal subunits  A new understanding of the decoding principle on the ribosome Sordarin inhibits fungal protein synthesis by blocking translocation differently to fusidic acid Ribosome structure and dynamics during translocation and termination Coot: Model-building tools for molecular graphics Antibiotics that bind to the A site of the large ribosomal subunit can induce mrna translocation Observation of intersubunit movement of the ribosome in solution using FRET mRNA translocation occurs during the second step of ribosomal intersubunit rotation Coupling of ribosomal L1 stalk and trna dynamics during translation elongation Initiation of translation by cricket paralysis virus IRES requires its translocation in the ribosome Ribosome dynamics and trna movement by time-resolved electron cryomicroscopy A ratchet-like inter-subunit reorganization of the ribosome during translocation The structure of the ribosome with elongation factor G trapped in the posttranslocational state Factor-free (\"non-enzymic\") and factor-dependent systems of translation of polyuridylic acid by escherichia coli ribosomes Mechanism of translocation in ribosomes. II. Activation of spontaneous (nonenzymic) translocation in ribosomes of Escherichia coli by p-chloromercuribenzoate Protein sliding along DNA: dynamics and structural characterization Lipid–protein interactions in double-layered two-dimensional AQP0 crystals Visualizing one-dimensional diffusion of eukaryotic DNA repair factors along a chromatin lattice Automatic estimation and correction of anisotropic magnification distortion in electron microscopes Measuring the optimal exposure for single particle cryo-EM using a 2.6 Å reconstruction of rotavirus VP6 Rotation of the head of the 30S ribosomal subunit during mrna translocation An end to 40 years of mistakes in DNA-protein association kinetics? Structural variant of the intergenic internal ribosome entry site elements in dicistroviruses and computational search for their counterparts GTP hydrolysis by EF-G synchronizes trna movement on small and large ribosomal subunits Intersubunit movement is required for ribosomal translocation The mechanism of eukaryotic translation initiation and principles of its regulation Divergent trna-like element supports initiation, elongation, and termination of protein biosynthesis Factorless ribosome assembly on the internal ribosome entry site of cricket paralysis virus Initiator met-trna-independent translation mediated by an internal ribosome entry site element in cricket paralysis virus-like insect viruses Structural rearrangements of the ribosome at the trna proofreading step Exotoxin a–eef2 complex structure indicates ADP ribosylation by ribosome mimicry EF-g-catalyzed translocation of anticodon stem-loop analogs of transfer RNA in the ribosome After the ribosome structure: How does translocation work? Structure of ratcheted ribosomes with trnas in hybrid states Elongation factor 2 as a novel target for selective inhibition of fungal protein synthesis Two crystal structures demonstrate large conformational changes in the eukaryotic ribosomal translocase The Phyre2 web portal for protein modeling, prediction and analysis Taura syndrome virus IRES initiates translation by binding its trna-mrna-like structural element in the ribosomal decoding center The electrostatic character of the ribosomal surface enables extraordinarily rapid target location by ribotoxins A conserved histidine in switch-ii of EF-G moderates release of inorganic phosphate Simulated-annealing real-space refinement as a tool in model building Structural dynamics of the ribosome Crystal structure of a 70S ribosome-trna complex reveals functional interactions and rearrangements Structural basis for translation termination on the 70S ribosome Conformational changes of elongation factor G on the ribosome during tRNA translocation Structural insights into ribosome translocation Direct measurement of the mechanical work during translocation by the ribosome Structural insights into viral ires-dependent translation mechanisms Likelihood-based classification of cryo-em images using FREALIGN Domain IV of elongation factor G from Thermus thermophilus is strictly required for translocation Intermediate states in the movement of transfer RNA in the ribosome Cramming more components onto integrated circuits Cryo-EM of ribosomal 80S complexes with termination factors reveals the translocated cricket paralysis virus IRES Structural elements in the internal ribosome entry site of plautia stali intestine virus responsible for binding with ribosomes Kinetic determination of the effects of ADP-ribosylation on the interaction of eukaryotic elongation factor 2 with ribosomes Recognition of cognate transfer RNA by the 30S ribosomal subunit Kinetically competent intermediates in the translocation step of protein synthesis Studies on the formation of trensfer ribonucleic acid-ribosome complexes. synthesis Translation elongation after assembly of ribosomes on the cricket paralysis virus internal ribosomal entry site without initiation factors or initiator trna UCSF chimera--a visualization system for exploratory research and analysis Mechanistic role of structurally dynamic regions in dicistroviridae IGR iress Control of ribosomal subunit rotation by elongation factor G Visualization of two transfer rnas trapped in transit during elongation factor g-mediated translocation Head swivel on the ribosome facilitates translocation by means of intra-subunit trna hybrid sites Structural determinants of an internal ribosome entry site that direct translational reading frame selection Alternative reading frame selection mediated by a trna-like domain of an internal ribosome entry site Hydrolysis of GTP by elongation factor G drives trna movement on the ribosome CTFFIND4: Fast and accurate defocus estimation from electron micrographs Optimal determination of particle orientation, absolute hand, and contrast loss in single-particle electron cryomicroscopy I-TASSER: a unified platform for automated protein structure and function prediction A dynamic RNA loop in an IRES affects multiple steps of elongation factor-mediated translation initiation Functional replacement of two highly conserved tetraloops in the bacterial ribosome Following movement of domain IV of elongation factor G during ribosomal translocation Translation initiation at the CUU codon is mediated by the internal ribosome entry site of an insect picorna-like virus in vitro An elongation factor g-induced ribosome rearrangement precedes trna-mrna translocation Structures of the bacterial ribosome at 3.5 A resolution Structure of the ribosome-bound cricket paralysis virus IRES RNA Structure of the 70S ribosome complexed with mrna and trna Visualization of the eef2-80s ribosome transition-state complex by cryo-electron microscopy Initial bridges between two ribosomal subunits are formed within 9.4 milliseconds, as studied by time-resolved cryo-em Domain movements of elongation factor eef2 and the eukaryotic 80S ribosome facilitate trna translocation Cryo-EM visualization of a viral internal ribosome entry site bound to human ribosomes Elongation factor G stabilizes the hybrid-state conformation of the 70S ribosome A model of the functioning ribosome: Locking and unlocking of the ribosome subparticles Rapid kinetic analysis of ef-g-dependent mrna translocation in the ribosome Structures of yeast 80S ribosome-trna complexes in the rotated and nonrotated conformations Blasticidin S inhibits translation by trapping deformed trna on the ribosome Cryo-EM structure of the mammalian eukaryotic release factor erf1-erf3-associated termination complex Structures of modified eef2 80S ribosome complexes reveal the role of GTP hydrolysis in translocation Elongation factor G bound to the ribosome in an intermediate state of translocation Locking and unlocking of ribosomal motions A new generation of the IMAGIC image processing system Structural basis of the translational elongation cycle The mechanism for activation of GTP hydrolysis on the ribosome Switch from cap- to factorless ires-dependent 0 and +1 frame translation during cellular stress and dicistrovirus infection Initiation of protein synthesis from the A site of the ribosome Binding mode of the first aminoacyl-trna in translation initiation mediated by plautia stali intestine virus internal ribosome entry site The I-TASSER Suite: protein structure and function prediction Crystal structures of ef-g-ribosome complexes trapped in intermediate states of translocation How the ribosome hands the a-site trna to the P site during ef-g-catalyzed translocation Crystal structures of complexes containing domains from two viral internal ribosome entry site (IRES) rnas bound to the 70S ribosome 10.7554/eLife.14874.059 Decision letter Subramaniam Sriram In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included. Thank you for submitting your article \"Ensemble cryo-EM uncovers inchworm-like translocation of a viral IRES through the ribosome\" for consideration by eLife. Your article has been favorably evaluated by John Kuriyan (Senior editor) and three reviewers, one of whom, Sriram Subramaniam, is a member of our Board of Reviewing Editors. The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. Summary: Type IV internal ribosome entry sites (IRESs) initiate translation without using any of the canonical eukaryotic translation initiation factors. Thus, they represent the most streamlined mode of eukaryotic translation initiation discovered. They have been studied biochemically and structurally. The current prevailing model is that these IRESs fold into a compact 2-domain structure that bind to the 40S subunit through multiple contacts. Critical interactions occur between two IRES RNA stem-loops and the head of the 40S subunit. This positions one of the IRES domains into the decoding groove. This domain (a pseudoknot, PKI) mimics the tRNA-mRNA anticodon-codon interaction, apparently first docking in the A site. The 60S subunit then joins in a GTP hydrolysis-independent step. This complex is then recognized by elongation factor 2, which catalyzes translocation of the PKI domain into the P site, allowing tRNA delivery to the A-site. Another round of translocation brings this tRNA to the P site. The mechanisms of these IRESs suggests that they can be powerful tools for understand translation in general (a feature the authors of this manuscript exploit). A rich set of biochemical and functional data have established that different parts of the IRES affect different steps, have shown similarities and differences between different type IV IRESs, and have established some of the key differences between the canonical and IRES-driven initiation mechanisms. In addition, various structures of the IRES alone or bound to the ribosome have been published, using both crystallography and cryo-EM. Until recently, the cryo-EM structures were of low-or mid-resolution. However, when combined with the higher resolution data from crystal structures and the many functional and biochemical studies, the models that resulted have been very informative and have allowed many predictions to be made. In this manuscript, the authors attack the question of IRES translocation. They present a series of cryo-EM structures of a Taura Syndrome Virus (TSV) IRES bound to 80S and eEF2, using the antibiotic sordarin. The authors interpret the set of structures as showing the trajectory of the mRNA-tRNA-mimicking PKI moving from the A to the P site. Overall, this is an impressive piece of work. It appears technically well done, the description is rich and detailed, and the conclusions are well supported and discussed. As such, it represents an important addition to the IRES field. In many ways, the mechanism that is presented is not surprising; an \"inchworm\"- like mechanism has been predicted in the literature (although never referred to as such!). However, to see it and to have detailed structures along the trajectory is very important. I will say that as the paper is written, it probably speaks as much to the mechanism of eEF2 and translocation in general as it does to IRES-specific function. Essential revisions: 1) There are few recent discoveries regarding IRES function that are not mentioned in the manuscript. As these discoveries relate directly to the interactions that the authors visualize and discuss, they should add a bit of discussion or analysis: • First, some type IV IRESs to can initiate in an alternate reading frame. Do their structures suggest how this might occur? This effect appears to relate to a base adjacent to the codon-anticodon mimic, which they have good density for. References: Au et al. (2015) PNAS 112:E6446-55, Wang et al. (2014) PloS One 9:e103601, Ren et al. (2012) PNAS 109:E630-9, Ren et al. (2014) Nucleic Acids Res. 42:9366-82. • Recent work implicates the VLR loop/loop 3 in PKI as having a role in eEF2 function: Ruehle et al., (2015) eLife), and it has been explored in manuscripts from the Jan lab. This is not mentioned or discussed. Can the authors comment on what this loop is doing and contacting and does it explain this previous work? Also, the Ruehle et al. presents biochemical data in favor of their spontaneous forward and reverse translocation that the authors allude to. 2) The interactions between the highly conserved apical loops of SL4 and 5 make critical interactions with eS5 and eS25. In addition, the IRES makes critical interaction with the L1 stalk. These regions of the type IV IRESs are very highly conserved, but no high-resolution information is known for these interactions. Was the local resolution good enough to say how binding these mysterious interactions are achieved, and perhaps how it relates to ribosome conformation, IRES conformation, etc.? 3) Related to the above, it would be interesting to see some more details of how the IRES changes conformation; not just globally, but internally. Is the resolution sufficient to see this? Any mechanistic insight? 4) By the very nature of this work, in which 5 structures at near atomic resolution are dissected, the figures are quite dense in information content and individual panels are generally quite small. In addition, the paper is quite long because of the high information content. The general reader can of course skip the detailed sections in the middle and read the Discussion, which is very clear. What seems to be missing for a general reader who wishes to dive into the \"information forest\" is an outline figure at the beginning that shows the ribosome translation cycle with various subunit motions and tRNA movements indicated. This would certainly help those who do not work in the ribosome field. 10.7554/eLife.14874.060 Author response 1) There are few recent discoveries regarding IRES function that are not mentioned in the manuscript. As these discoveries relate directly to the interactions that the authors visualize and discuss, they should add a bit of discussion or analysis: First, some type IV IRESs to can initiate in an alternate reading frame. Do their structures suggest how this might occur? This effect appears to relate to a base adjacent to the codon-anticodon mimic, which they have good density for. References: Au et al. (2015) PNAS 112:E6446-55, Wang et al. (2014) PloS One 9:e103601, Ren et al. (2012) PNAS 109:E630-9, Ren et al. (2014) Nucleic Acids Res. 42:9366-82. We agree that alternative frame selection is an interesting phenomenon andhave added a paragraph to discuss this in “IRES translocation mechanism” (third paragraph). Our structures do not directly suggest how alternate reading frame selection could occur because our data did not reveal a frame-shifted conformation of the IRES. The observation of IRES dynamics in our study indirectly suggests that an alternate (frame-shifted) codon may be transiently placed in the A site following eEF2 release, and this sampling may allow binding of an aminoacyl-tRNA to the off-frame codon. Recent work implicates the VLR loop/loop 3 in PKI as having a role in eEF2 function: Ruehle et al., (2015) eLife), and it has been explored in manuscripts from the Jan lab. This is not mentioned or discussed. Can the authors comment on what this loop is doing and contacting and does it explain this previous work? Also, the Ruehle et al. presents biochemical data in favor of their spontaneous forward and reverse translocation that the authors allude to. We have revised our Results section to address this important comment. Loop 3 connects the ASL-like and the mRNA-like regions of the PKI domain. Loop 3 of the post-translocated state (Structure V) is stabilized by interactions with the β-hairpin loop of uS7 and helix 23 of 18S rRNA in the E site, in a manner reminiscent of that for the E-site tRNA in the 80S*2tRNA*mRNA structure. In the pre-translocation states, however, loop 3 is poorly resolved in density maps. This implies conformational flexibility of loop 3, also reported by biochemical studies of unbound IGR IRESs (Jan and Sarnow, 2002; Pfingsten et al., 2007). Our structures therefore suggest that loop 3 contributes to stabilization of the post-translocation IRES, rationalizing the recent detailed biochemical study (Ruehle et al., (2015) eLife), which reported that IGR IRES mutated constructs with shortened loop 3 are defective in eEF2-catalyzed translocation. 2) The interactions between the highly conserved apical loops of SL4 and 5 make critical interactions with eS5 and eS25. In addition, the IRES makes critical interaction with the L1 stalk. These regions of the type IV IRESs are very highly conserved, but no high-resolution information is known for these interactions. Was the local resolution good enough to say how binding these mysterious interactions are achieved, and perhaps how it relates to ribosome conformation, IRES conformation, etc.? We find that the phosphate backbone of SL4 and 5 interact with the positively charged and aromatic residues of eS5 (uS7) and eS25. We have added a description of these interactions in the main text and also Supplementary Figures (Figure 3—figure supplement 2, Figure 3—figure supplement 4) to demonstrate the interactions. In addition, we find that interactions of SL4 and SL5 with the small subunit are somewhat similar to those of the L1 stalk with the small subunit of the hybrid-state 80S*tRNA structure. We have added Figure 3—figure supplement 3 to illustrate this similarity. The interactions between the IRES and the L1 stalk are less well resolved – although the density is strong, the resolution is insufficient to interpret the interactions unambiguously. We therefore refrain from detailed interpretation of L1 stalk interactions. 3) Related to the above, it would be interesting to see some more details of how the IRES changes conformation; not just globally, but internally. Is the resolution sufficient to see this? Any mechanistic insight? We now provide a more extensive discussion of IRES local interactions and conformational changes, supplemented by additional illustrations. We report the rearrangements of stem loop 3 (conserved in TSV-like IRESs of group 2: Aparavirus), which resembles a tRNA elbow, as we reported previously. Our current structures indicate that SL3 undergoes rearrangements similar to those of the translocating A-site tRNA (Figure 1—figure supplement 6). In addition, we demonstrate local rearrangements of the “bridge” between the 5’ domain and PKI domain (Figure 3—figure supplement 7). This region interacts with protein uL5 in the two most compact IRES conformations (III and IV), but not in other states. This reveals the stabilizing role of protein uL5 at the intermediate stages of IRES translocation. 4) By the very nature of this work, in which 5 structures at near atomic resolution are dissected, the figures are quite dense in information content and individual panels are generally quite small. In addition, the paper is quite long because of the high information content. The general reader can of course skip the detailed sections in the middle and read the Discussion, which is very clear. What seems to be missing for a general reader who wishes to dive into the \"information forest\" is an outline figure at the beginning that shows the ribosome translation cycle with various subunit motions and tRNA movements indicated. This would certainly help those who do not work in the ribosome field. We have reorganized the panels in most figures to make the figures less dense and increase the sizes of individual panels. We agree that a figure showing ribosome-2tRNA-mRNA and summarizing conformational differences between structures representing various translocation states would be helpful. We now include a supplementary figure, showing ribosome-2tRNAs-mRNA structures (Figure 1—figure supplement 1), which we refer to in the manuscript. We also include the views of tRNA-bound structures in the supplementary figure showing interactions of the A-site finger with the tRNAs and the IRES (Figure 3—figure supplement 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diff --git a/annotated_BioC_JSON/PMC4918759_ann.json b/annotated_BioC_JSON/PMC4918759_ann.json
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+[{"sourceid":"4918759","sourcedb":"","project":"","target":"","text":"Structures of human ADAR2 bound to dsRNA reveal base-flipping mechanism and basis for site selectivity ADARs (adenosine deaminases acting on RNA) are editing enzymes that convert adenosine (A) to inosine (I) in duplex RNA, a modification reaction with wide-ranging consequences on RNA function. Our understanding of the ADAR reaction mechanism, origin of editing site selectivity and effect of mutations is limited by the lack of high-resolution structural data for complexes of ADARs bound to substrate RNAs. Here we describe four crystal structures of the deaminase domain of human ADAR2 bound to RNA duplexes bearing a mimic of the deamination reaction intermediate. These structures, together with structure-guided mutagenesis and RNA-modification experiments, explain the basis for ADAR deaminase domain’s dsRNA specificity, its base-flipping mechanism, and nearest neighbor preferences. In addition, an ADAR2-specific RNA-binding loop was identified near the enzyme active site rationalizing differences in selectivity observed between different ADARs. Finally, our results provide a structural framework for understanding the effects of ADAR mutations associated with human disease. Introduction RNA editing reactions alter a transcript’s genomically encoded sequence by inserting, deleting or modifying nucleotides. Deamination of adenosine (A), the most common form of RNA editing in humans, generates inosine (I) at the corresponding nucleotide position. Since I base pairs with cytidine (C), it functions like guanosine (G) in cellular processes such as splicing, translation and reverse transcription. A to I editing has wide-ranging consequences on RNA function including altering miRNA recognition sites, redirecting splicing and changing the meaning of specific codons. Two different enzymes carry out A to I editing in humans; ADAR1 and ADAR2. ADAR activity is required for nervous system function and altered editing has been linked to neurological disorders such as epilepsy and Prader Willi Syndrome. In addition, mutations in the ADAR1 gene are known to cause the autoimmune disease Aicardi-Goutieres Syndrome (AGS) and the skin disorder Dyschromatosis Symmetrica Hereditaria (DSH). Hyper editing has been observed at certain sites in cancer cells, such as in the mRNA for AZIN1 (antizyme inhibitor 1). However, hypo editing also occurs in cancer-derived cell lines exemplified by reduced editing observed in the message for glioma-associated oncogene 1 (Gli1). The ADAR proteins have a modular structure with double stranded RNA binding domains (dsRBDs) and a C-terminal deaminase domain (see Fig. 1a for hADAR2 domains). ADARs efficiently deaminate specific adenosines in duplex RNA while leaving most adenosines unmodified. The mechanism of adenosine deamination requires ADAR to flip the reactive base out of an RNA double helix to access its active site. How an enzyme could accomplish this task with a duplex RNA substrate is not known. Furthermore, how an ADAR deaminase domain contributes to editing site selectivity is also unknown, since no structures of ADAR deaminase domain-RNA complexes have been reported. To address these knowledge gaps, we set out to trap the human ADAR2 deaminase domain (aa299–701, hADAR2d) bound to different duplex RNAs and solve structures for the resulting complexes using x-ray crystallography. We then evaluated the importance of protein-RNA contacts using structure-guided mutagenesis and RNA-modification experiments coupled with adenosine deamination kinetics. Results Trapping the flipped conformation The ADAR reaction involves the formation of a hydrated intermediate that loses ammonia to generate the inosine-containing product RNA (for reaction scheme see Fig. 1b). The covalent hydrate of the nucleoside analog 8-azanebularine (N) mimics the proposed high-energy intermediate (for reaction scheme see Fig. 1b). For trapping hADAR2d bound to RNA for crystallography, we incorporated 8-azanebularine into duplex RNAs shown recently to be excellent substrates for deamination by hADAR2d (for duplex sequence see Fig. 1c) (for characterization of protein–RNA complex see Supplementary Fig. 1). In addition, for one of these duplexes (Bdf2), we positioned the 8-azanebularine opposite either uridine or cytidine to mimic an A-U pair or A-C mismatch at the editing site creating a total of three different RNA substrates for structural studies (Fig. 1c). The hADAR2d protein (without RNA bound) has been previously crystallized and structurally characterized revealing features of the active site including the presence of zinc. In addition, an inositol hexakisphosphate (IHP) molecule was found buried in the core of the protein hydrogen bonded to numerous conserved polar residues. For crystallization of hADAR2d-RNA complexes, we used both the wild type (WT) deaminase domain and a mutant (E488Q) that has enhanced catalytic activity. A description of the crystallization conditions, X-ray diffraction data collection and solution of the structures can be found in Online Methods. Four protein-RNA combinations generated diffracting crystals that resulted in high-resolution structures (hADAR2d WT–Bdf2-U, hADAR2d WT–Bdf2-C, hADAR2d E488Q–Bdf2-C, hADAR2d E488Q–Gli1) (Table 1). In each of these complexes, the protein binds the RNA on one face of the duplex over ~ 20 bp using a positively charged surface near the zinc-containing active site (Fig. 2, Supplementary Fig. 2a). The large binding site (1493 Å2 RNA surface area and 1277 Å2 protein surface area buried) observed for hADAR2d is consistent with recent footprinting studies. Both strands of the RNA contact the protein with the majority of these interactions mediated through the phosphodiester-ribose backbone near the editing site (Fig. 2c, Supplementary Fig. 2 b–d). The structures show a large deviation from A-form RNA conformation at the editing site (Fig. 2, Fig. 3, Supplementary Video 1). The 8-azanebularine is flipped out of the helix and bound into the active site as its covalent hydrate where it interacts with several amino acids including V351, T375, K376, E396 and R455 (Fig. 3a, Supplementary Fig. 3a). The side chain of E396 H-bonds to purine N1 and O6. This interaction was expected given the proposed role of E396 in mediating proton transfers to and from N1 of the substrate adenosine. The 2’-hydroxyl of 8-azanebularine H-bonds to the backbone carbonyl of T375 while the T375 side chain contacts its 3’-phosphodiester. R455 and K376 help position the flipped nucleotide in the active site by fastening the phosphate backbone flanking the editing site. The R455 side chain ion pairs with the 5’-phosphodiester of 8-azanebularine while the K376 side chain contacts its 3’-phosphodiester. Lastly, the side chain of V351 provides a hydrophobic surface for interaction with the nucleobase of the edited nucleotide. RNA binding does not alter IHP binding or the H-bonding network linking IHP to the active site. ADARs use a unique mechanism to modify duplex RNA The ADAR2 base-flipping loop, bearing residue 488, approaches the RNA duplex from the minor groove side at the editing site. The side chain of this amino acid penetrates the helix where it occupies the space vacated by the flipped out base and H-bonds to the complementary strand orphaned base and to the 2’ hydroxyl of the nucleotide immediately 5’ to the editing site (Figs. 3b, 3c). In the four structures reported here, three different combinations of helix-penetrating residue and orphan base are observed (i.e. E488 + U, E488 + C and Q488 + C) and all three combinations show the same side chain and base positions (Figs. 3b, 3c, Supplementary Fig. 4a for overlay of all three). For instance, in the complex with hADAR2d E488Q and the Bdf2-C duplex, the protein recognizes an orphaned C by donating H-bonds from Nε2 to cytosine N3 and from its backbone NH to cytosine O2 (Fig. 3b). In the complex with hADAR2d WT and the Bdf2-U duplex, a similar interaction is observed with the E488 backbone NH hydrogen bonded to the uracil O2 and the E488 side chain H-bonded to the uracil N3H (Fig. 3c). Interestingly, the E488Q mutant was discovered in a screen for highly active ADAR2 mutants and this residue was suggested to be involved in base flipping given its effect on editing substrates with a fluorescent nucleobase at the editing site. ADARs react preferentially with adenosines in A•C mismatches and A-U pairs over A•A and A•G mismatches. A purine at the orphan base position (in its anti conformation) would clash with the 488 residue explaining the preference for pyrimidines here. The interaction of the 488 residue with the orphaned base is reminiscent of an interaction observed for Hha I DNA methyltransfersase (MTase), a duplex DNA modifying enzyme that also uses a base flipping mechanism to access 2’-deoxycytidine (dC) for methylation. For that enzyme, Q237 H-bonds to an orphaned dG while it fills the void left by the flipped out dC (Supplementary Fig. 4b). In addition, two glycine residues flank Q237 allowing the loop to adopt the conformation necessary for penetration into the helix. The flipping loop in ADAR2 (i.e. aa487–489) also has the helix-penetrating residue flanked by glycines. However, unlike the case of the DNA MTase that approaches the DNA from the major groove, the ADAR2 loop approaches the duplex from the minor groove side. Such an approach requires deeper penetration of the intercalating residue to access the H-bonding sites on the orphaned base, necessitating an additional conformational change in the RNA duplex. This change includes shifting of the base pairs immediately 5’ to the editing site toward the helical axis and a widening of the major groove opposite the editing site (Figs. 4a, 4b, Supplementary Video 1). In the case of the hADAR2d WT–Bdf2-U RNA, this shift is accompanied by a shearing of the U11-A13' base pair with U11 shifted further in the direction of the major groove creating an unusual U-A \"wobble\" interaction with adenine N6 and N1 within H-bonding distance to uracil N3H and O2, respectively (Fig. 4c, Supplementary Fig. 3b). This type of wobble pair has been observed before and requires either the imino tautomer of adenine or the enol tautomer of uracil. The ADAR-induced distortion in RNA conformation results in a kink in the RNA strand opposite the editing site (Fig. 4b). This kink is stabilized by interactions of the side chains of R510 and S495 with phosphodiesters in the RNA backbone of the unedited strand (Fig. 4a). Interestingly, ADAR2’s flipping loop approach from the minor groove side is like that seen with certain DNA repair glycosylases (e.g. UDG, HOGG1, and AAG) that also project intercalating residues from loops bound in the minor groove (Supplementary Fig. 5a). However, these enzymes typically bend the DNA duplex at the site of modification to allow for penetration of intercalating residues and damage recognition. While hADAR2d clearly alters the duplex conformation to gain access to the modification site from the minor groove, it does not bend the RNA duplex (Figs. 2a, 2b, 4b). Furthermore, ADARs do not modify duplex DNA. The DNA B-form helix has groove widths and depths that would prevent productive interactions with ADAR. For instance, ADAR can readily penetrate an A-form helix from the minor groove side and place the helix-penetrating residue in the space occupied by the editing site base (Supplementary Fig. 6). However, this residue cannot penetrate the minor groove enough to occupy the base position in a B-form helix (Supplementary Fig. 6). Furthermore, DNA lacks the 2’ hydroxyls that are used by ADAR for substrate recognition (Fig. 2c). Indeed, in each of the four complexes reported here, the protein contacts at least five ribose 2’ hydroxyl groups (Fig. 2c, Supplementary Fig. 2 b–d). Thus, hADAR2d uses a substrate recognition and base flipping mechanism with similarities to other known nucleic acid-modifying enzymes but uniquely suited for reaction with adenosine in the context of duplex RNA. Structures explain nearest neighbor preferences ADARs have a preference for editing adenosines with 5’ nearest neighbor U (or A) and 3’ nearest neighbor G. The ADAR2 flipping loop occupies the minor groove spanning the three base pairs that include the nearest neighbor nucleotides flanking the edited base (Figs. 3b, 3c). As described above, the base pair including the 5’ nearest neighbor U (U11-A13’ in the Bdf2 duplex) is shifted from the position it would occupy in a typical A-form helix to accommodate the loop (Fig. 4a). Also, the minor groove edge of this pair is juxtaposed to the protein backbone at G489. Modeling a G-C or C-G pair at this position (i.e. 5’ G or 5’ C) suggests a 2-amino group in the minor groove would clash with the protein at G489 (Fig. 5a, Supplementary Fig. 7c). Indeed, replacing the U-A pair adjacent to the editing site with a C-G pair in the Gli1 duplex substrate resulted in an 80% reduction in the rate of hADAR2d-catalyzed deamination (Figs. 5b, 5c). To determine whether this effect arises from an increase in local duplex stability from the C-G for U-A substitution or from the presence of the 2-amino group, we replaced the U-A pair with a U-2-aminopurine (2AP) pair. 2AP is an adenosine analog that forms a base pair with uridine of similar stability to a U-A pair, but places an amino group in the minor groove (Fig. 5b). Importantly, this substitution also resulted in an 80% reduction in rate, illustrating the detrimental effect of the amino group in the minor groove at this location. These observations suggest that hADAR2’s 5’ nearest neighbor preference for U (or A) is due to a destabilizing clash with the protein backbone at G489 that results from the presence of an amino group in the minor groove at this location for sequences with 5’ nearest neighbor G or C. However, the observed clash is not severe and the enzyme would be able to accommodate G or C 5’ nearest neighbors by slight structural perturbations, explaining why this sequence preference is not an absolute requirement. In each of the hADAR2d-RNA structures reported here, the backbone carbonyl oxygen at S486 accepts an H-bond from the 2-amino group of the G on the 3’ side of the edited nucleotide (Fig. 5d). Guanine is the only common nucleobase that presents an H-bond donor in the RNA minor groove suggesting that other nucleotides in this position would reduce editing efficiency. Indeed, mutating this base to A, C or U, while maintaining base pairing at this position, reduced the rate of deamination by hADAR2d in Gli1 mRNA model substrates (Supplementary Fig. 7 a–b). To test the importance of the amino group on the 3’ G in the hADAR2d reaction, we prepared RNA duplex substrates with purine analogs on the 3’ side of the edited A (Fig. 5e). We tested a G analog that lacks the 2-amino group (inosine, I) and one that blocks access to this amino group (N2-methylguanosine (N2MeG). In addition, we compared a 3’ A to a 3’ 2AP since 2AP could form the H-bonding interaction observed with S486. We found the substrate with a 3’ N2MeG to be unreactive to hADAR2d-catalyzed deamination confirming the importance of the observed close approach by the protein to the 3’ G 2-amino group (Fig. 5f). In addition, the substrate with a 3’ I displayed a reduced deamination rate compared to the substrate with a 3’ G suggesting the observed H-bond to the 2-amino group contributes to the 3’ nearest neighbor selectivity (Fig. 5f). This conclusion is further supported by the observation that deamination in the substrate with a 3’ 2AP is faster than in the substrate with a 3’ A (Fig. 5f). RNA-binding loops of the ADAR catalytic domain The structures reported here identify RNA-binding loops of the ADAR catalytic domain and suggest roles for several amino acids not previously known to be important for editing, either substrate binding or catalysis (Fig. 6). The side chain for R510 ion-pairs with the 3’ phosphodiester of the orphaned nucleotide (Figs. 3a, 3c). This residue is conserved in ADAR2s and ADAR1s, but is glutamine in the editing-inactive ADAR3s (Supplementary Table 1). Mutation of hADAR2d at this site to either glutamine (R510Q) or to alanine (R510A) reduced the measured deamination rate constant by approximately an order of magnitude (Fig. 6c). In addition, the contact point near the 5’ end of the unedited strand involves G593, K594 and R348, residues completely conserved in the family of ADAR2s (Fig. 2c, Supplementary Table 1). Mutation of any of these residues to alanine (G593A, K594A, R348A) substantially reduces editing activity (Fig. 6c). In addition, mutation of G593 to glutamic acid (G593E) resulted in a nearly two orders of magnitude reduction in rate, consistent with proximity of this residue to the negatively charged phosphodiester backbone of the RNA (Fig. 6c). RNA binding leads to an ordering of the 454–477 loop, which was disordered in the RNA-free hADAR2d structure (Fig. 1d, green) (Supplementary Video 2). This loop binds the RNA duplex contacting the minor groove near the editing site and inserting into the adjacent major groove (Fig. 6e). This loop sequence is conserved in ADAR2s but different in the family of ADAR1s (Fig. 6d). The substantial difference in sequence between the ADARs in this RNA-binding loop suggests differences in editing site selectivity between the two ADARs arise, at least in part, from differences in how this loop binds RNA substrates. Discussion Base flipping is a well-characterized mechanism by which nucleic acid modifying enzymes gain access to sites of reaction that are otherwise buried in base-paired structures. DNA methylases, DNA repair glycosylases and RNA loop modifying enzymes are known that flip a nucleotide out of a base pair. However, none of the structurally characterized base-flipping enzymes access their reactive sites from within a normal base-paired RNA duplex. We are aware of one other protein-induced nucleotide flipping from an RNA duplex region. Bacterial initiation factor 1 (IF1) binds to the 30S ribosomal subunit at helix 44 of 16S RNA with A1492 and A1493 flipped out of the helix and bound into protein pockets (Supplementary Fig. 5b). However, these nucleotides are located in a highly distorted and dynamic duplex region containing several mismatches and are predisposed to undergo this conformational change. Thus, this system is not illustrative of base flipping from a normal duplex and does not involve an enzyme that must carryout a chemical reaction on the flipped out nucleotide. Other RNA modification enzymes are known that flip nucleotides out of loops, even from base pairs in loop regions (pseudoU synthetase, tRNA transglycosylase, and restrictocin bound to sarcin/ricin loop of 28S rRNA) (Supplementary Fig. 5b). Because the modification sites are not flanked on both sides by normal duplex, these enzymes do not experience the same limits in approach to the substrate that ADARs do. The fact that ADARs must induce flipping from a normal duplex has implications on its preference for adenosines flanked by certain base pairs, a phenomenon that was not well understood prior to this work. In our structures, the flipped out 8-azanebularine is hydrated, mimicking the tetrahedral intermediate predicted for deamination of adenosine (Figs. 1b, 3a, Supplementary Fig. 3 a–b). Our use of 8-azanebularine, with its high propensity to form a covalent hydrate, allowed us to capture a true mimic of the tetrahedral intermediate and reveal the interactions between the deaminase active site and the reactive nucleotide. In addition, 8-azanebularine was found to adopt a 2’-endo sugar pucker with its 2’-hydroxyl H-bonded to the protein backbone carbonyl at T375. The 2’ endo conformation appears to facilitate access of the nucleobase to the zinc-bound water for nucleophilic attack at C6. Several other base-flipping enzymes stabilize the altered nucleic acid conformation by intercalation of an amino acid side chain into the space vacated by the flipped out base. For hADAR2, E488 serves this role. In the two structures with wild type hADAR2, the E488 residue and orphan base are in nearly identical positions (see Supplementary Fig. 4a for overlay). Thus, the E488 side chain directly contacts each orphan base, likely by accepting an H-bond from uracil N3H or by donating an H-bond to cytidine N3. The latter interaction requires E488 to be protonated. The pKa of E488 in the ADAR-RNA complex has not been measured, but proximity to H-bond acceptors, such as cytidine N3, and insertion between stacked nucleobases, would undoubtedly elevate this value and could lead to a substantial fraction in the protonated state at physiologically relevant pH. Since the glutamine side chain is fully protonated under physiologically relevant conditions, a rate enhancement for the E488Q mutant would be expected if the reaction requires E488 protonation. The interactions of hADAR2d with base pairs adjacent to the editing site adenosine explain the known 5’ and 3’ nearest neighbor preferences (Fig. 5). While these studies indicate the ADAR2 catalytic domain makes an important contact to the 3’ nearest neighbor G, Stefl et al. suggested the 3’ G preference arises from dsRBD binding selectivity for ADAR2. These authors reported a model for ADAR2’s dsRBDs bound to an editing substrate based on NMR data from the isolated dsRBDs (lacking the deaminase domain) and short RNA fragments derived from the GluR-B R/G site RNA. They describe an interaction wherein the 3’ G 2-amino group H-bonds to the backbone carbonyl of S258 found in the β1-β2 loop of ADAR2’s dsRBDII. It is not possible for the S486-3’G interaction we describe here and the S258-3’G interaction reported by Stefl et al. to exist in the same complex since both involve protein loops bound in the RNA minor groove at the same location. Because our structures have captured the edited nucleotide in the conformation required to access the active site, the interactions observed here are highly likely to occur during the deamination reaction at the editing site. However, since dsRBDs are known to bind promiscuously with duplex RNA, it is possible that the S258-3’G interaction found in a complex lacking the deaminase domain is not relevant to catalysis at the editing site. It is also possible that ADAR dsRBD and catalytic domain binding are sequential, with release of the dsRBD from the RNA taking place prior to catalytic domain engagement and base flipping. Aicardi-Goutieres Syndrome (AGS) and Dyschromatosis Symmetrica Hereditaria (DSH) are human diseases caused by mutations in the human ADAR1 gene and several of the disease-associated mutations are found in the deaminase domain. Given the conservation in RNA binding surface and active site residues, we expect the hADAR1 catalytic domain to bind RNA with a similar orientation of the helix found in our hADAR2d-RNA structures. When one maps the locations of the AGS-associated mutations onto the hADAR2d-RNA complex, two mutations involve residues in close proximity to the RNA (\u003c 4 Å) (Supplementary Fig. 8a). G487 of hADAR2 is found on the flipping loop near the RNA (Fig. 3b). Sequence in this loop is highly conserved among ADARs and corresponds to G1007 in hADAR1 (Supplementary Table 2). An arginine at this position would preclude close approach of the flipping loop to the RNA, preventing E1008 insertion and base flipping into the active site (Supplementary Fig. 8b). This is consistent with the observation that the G1007R mutation in hADAR1 inhibits RNA editing activity. Also, K376 forms salt bridges with both the 5’ and 3’ phosphodiesters of the guanosine on the 3’ side of the editing site (Fig. 2). The corresponding residue in hADAR1 (R892) could form similar contacts and the R892H mutation would likely alter this interaction. In summary, the structures described here establish human ADAR2 as a base-flipping enzyme that uses a unique mechanism well suited for modifying duplex RNA. In addition, this work provides a basis for understanding the role of the ADAR catalytic domain in determining editing site selectivity and additional structural context to evaluate the impact of ADAR mutations associated with human disease. Online Methods Unless otherwise stated, reagents were purchased from Fisher Scientific, Sigma-Aldrich, or Life Technologies. T4 polynucleotide kinase, T4 DNA ligase, molecular biology grade bovine serum albumin (BSA), and RNase inhibitor were purchased from New England Biolabs. γ-[32P] ATP was purchased from Perkin-Elmer Life Sciences. The Avian Myeloblastosis Virus (AMV) reverse transcriptase, deoxynucleotide triphosphate (dNTP) mix and RQ1 RNase free DNase were purchased from Promega. Pfu Ultra II was purchased from Stratagene. Dpn 1 was purchased from Invitrogen. Quickchange XL II mutagenesis kit was purchased from Agilent Technologies. RNA oligonucleotides were synthesized at the University of Utah DNA/Peptide Core Facility or purchased from GE Healthcare Dharmacon, Inc. or Sigma Aldrich. DNA oligonucleotides were purchased from Integrated DNA Technologies. Storage phosphor imaging plates from Molecular Dynamics were imaged using Molecular Dynamics 9400 Typhoon phosphorimager. Data were analyzed using Molecular Dynamics ImageQuant 5.2 software. Electrospray Ionization (ESI) mass spectrometry of oligonucleotide samples was carried out at the Campus Mass Spectrometry Facilities, UC Davis. Oligonucleotide masses were determined using Mongo Oligo Mass Calculator v2.06. Expression and purification of hADAR2 deaminase domain (hADAR2d) for crystallography Protein expression and purification were carried out by modifying a previously reported protocol. In brief, BCY123 cells were transformed with a pSc-ADAR construct encoding either hADAR2d-WT or hADAR2d-E488Q (corresponding to the deaminase domain; residues 299–701). Cells were streaked on yeast minimal media minus uracil (CM-ura) plates. A single colony was used to inoculate a 15 mL CM-ura starter culture. After shaking at 300 rpm and 30 °C overnight, 10 mL of starter culture was used to inoculate each liter of yeast growth media. After 24 h, cells were induced with the addition of 110 mL of sterile 30% galactose per liter, and protein was expressed for 5 h. Cells were collected by centrifugation and stored at −80 °C. Cells were lysed in Buffer A (20 mM Tris-HCl pH 8.0, 5% glycerol, 35 mM imidazole, 1mM BME, 0.01% Triton × 100) with 750 mM NaCl using a microfluidizer and cell lysate clarified by centrifugation (39,000 × g for 25 min). Lysate was passed over a 5 mL Ni-NTA column; washed in three steps with 20–50mL of Lysis Buffer, Wash I buffer (Buffer A + 300 mM NaCl), and Wash II buffer (Buffer A + 100 mM NaCl); and protein eluted by a 35–300 mM imidazole gradient in Wash II over 80 min at a flow rate of 1 ml/min. Fractions containing the target protein were pooled and further purified on a 2 mL GE Healthcare Lifesciences Hi-Trap Heparin HP column in the absence of BME. The 10xHis fusion protein was cleaved with an optimized ratio of 1 mg of TEV protease for each 1 mg of protein. Cleavage was carried out for 1–2 h before passing the product over another Ni-NTA column at 0.5 mL/min. The flow-through and wash were collected; dialyzed against 20 mM Tris pH 8.0, 200 mM NaCl, 5% glycerol, and 1 mM BME; and concentrated to just under 1 mL for gel filtration on a GE Healthcare HiLoad 16/600 Superdex 200 PG column. Fractions containing purified protein were pooled and concentrated to 5–7 mg/mL for crystallography trials. Purification of RNAs for crystallography The 8-azanebularine (N) phosphoramidite was synthesized as previously described and RNAs were synthesized as previously described. Single-stranded RNAs (See Supplementary Table 2 for sequences) were purified by denaturing polyacrylamide gel electrophoresis and visualized using UV shadowing. Bands were excised from the gel, crushed and soaked overnight at 4 °C in 0.5 M NH4OAc, 0.1% sodium dodecyl sulfate (SDS) and 0.1 mM EDTA. Polyacrylamide fragments were removed using a 0.2 µm filter followed by desalting on C18 Sep-Pak column. The RNA solutions were lyophilized to dryness, re-suspended in nuclease-free water, quantified by absorbance at 260 nm and stored at −70 °C. Oligonucleotide mass was confirmed by electrospray ionization mass spectrometry. Unmodified RNA stands were purchased from Dharmacon-GE Life Sciences and purified as described above. Duplex RNA was hybridized in a 1:1 ratio by heating to 95 °C for 5 min and slowly cooling to 30 °C. hADAR2d-RNA complex crystallization Crystals of hADAR2d E488Q+Bdf2-C RNA complex were grown at room temperature by the sitting drop vapor diffusion method. A solution of 0.5 µL volume containing 4.5 mg/mL protein and 70 µM of Bdf2-C 23mer RNA (1:0.7 ADAR2:RNA molar ratio) were mixed with 0.5 µL of 0.1 M MES:NaOH pH 6.5, 9% (w/v) PEG 3350, 13% glycerol, and 0.015M NAD, which was added to improve crystal growth. Crystals took several weeks to grow. A single, cube-shaped crystal about 120 µm in size was soaked briefly in a solution of mother liquor plus 30% glycerol before flash-cooling in liquid nitrogen. Data were collected via fine-phi slicing using 0.2° oscillations on beamline 24-ID-C at the Advanced Photon Source at Argonne National Laboratories in Chicago. To obtain crystals of the hADAR2d+WT:Bdf2-C RNA, an identical procedure was used as above; however, the crystallization conditions had slightly different concentrations of reagents (10% PEG 3350, 15% glycerol, 0.1 M MES:NaOH pH 6.5, no NAD). For the hADAR2d+WT:Bdf2-U construct, hanging drop vapor diffusion using 200 nL of a mixture containing 4.5 mg/mL protein and 70 µM of Bdf2-U (1:0.7 molar ratio) and 200 nL of a mother liquor (0.1 M ammonium acetate, 0.1M Bis-tris pH 5.5, 17% PEG 10,000) yielded several crystals with a morphology similar to the one described above. All wild type crystals were soaked briefly in a solution of mother liquor plus 30% glycerol before flash-cooling in liquid nitrogen. Data were collected via fine-phi slicing using 0.2° oscillations on beamline 12-2 at the Stanford Synchrotron Radiation Lightsource. Crystals of the hADAR2d E488Q+Gli1 RNA complex were grown using hanging drop vapor diffusion. A solution of volume 200 nL containing 4.5 mg/mL protein and 100 µM of Gli1 23mer RNA (1:1 ADAR2:RNA molar ratio) were mixed with 200 nL of 0.1 M MES:NaOH pH 6.5 and 12% PEG 20,000. At room temperature, a single diamond-shaped crystal about 150 µm long and 50 µm wide was observed about a week later. This crystal was soaked briefly in a solution of mother liquor plus 30% glycerol before flash-cooling in liquid nitrogen. Data were collected on beamline 12-2 at the Stanford Synchrotron Radiation Lightsource using the fine-phi splicing described above. Processing and refinement of crystallographic data Data for the E488Q Bdf2-C-bound and Gli1-bound structures were processed using XDS and scaled with Aimless (CCP4 1994). Diffraction data for hADAR2d wild type structures were processed with XDS and scaled with SCALA (Kabsch, 2010). The RNA-free hADAR2d crystal structure (PDB ID: 1ZY7) was used as a model for molecular replacement using PHENIX. The structures were refined using PHENIX including TLS parameters and Zn coordination restraints. Ideal Zn-ligand distances were determined using average distances found for similar coordination models in the PDB database. Table 1 gives the statistics in data processing and model refinement. The asymmetric unit for Gli1-bound hADAR2d E488Q includes two complexes of protein:RNA. In each of these complexes, the first 17 residues of the deaminase domain (residues 299–316) as well as a C-terminal proline (Pro701) are disordered and were therefore not included in the model. However, although the RNA-free ADAR2 structure (PDB ID: 1ZY7) lacked electron density for residues 457–475, we observed density for the backbone atoms of these residues. These residues were initially modeled as polyalanine. After a several rounds of refinement, electron density revealed the location of some side chains. Residues whose basic side chains interact with the RNA backbone are clearly defined in the final density map. Although some non-RNA-binding side chains show only weak density, backbone density is strong. As observed in the original hADAR2d RNA-free structure, inositol hexakisphosphate (IHP) was buried in the enzyme core. The asymmetric units for Bdf2-bound ADARs contain one ADAR2d:RNA complex (protein chain A) and one RNA-free ADAR2d monomer (chain D). The N-terminus of the Bdf2-bound structures include more residues than Gli1-bound, beginning at Pro305 in chain A and Thr304 for chain D in the mutant structure, and beginning at Arg307 in chain A and Thr304 or Pro305 in chain D in the wild type structures. The first few residues (in structures in which the specified residues are modeled) had weak side chain density, including residues 305 and 307 in chain A, and residues 304–307 in chain D, and are modeled in the structure as alanine. The last residue of E488Q+Bdf2-C, Pro701, had very weak electron density for both protein subunits in the asymmetric unit. Unlike the E488Q+Gli1 structure, electron density was defined better in the originally disordered loop (residues 457–475) for most residues in the Bdf2-bound structures. With the exception of Glu466 in the wildtype structures, we were able to model-build in main chain and side chain atoms for all residues of this loop in the ADAR subunit complexed to the Bdf2 RNA duplexes. In the RNA-free subunit (chain D) of E488Q+Bdf2-C, a crystal contact stabilized this flexible loop so that we were able to model in the backbone for residues 457–475, but residues 465–475 were modeled as alanine because of poorly defined side chain density. An identical crystal contact was observed in the wildtype structures. In the wt+Bdf2-C complex, density for residues 467–470 was strong enough for side chains to be included in these structures; however, side chain density was not strong for residues 465, 466, 471, 473–475, and 477. Therefore, these side chains were not included in the model. In wt+Bdf2-U, density for side chains 465, 466, 470, 471, and 473–475 was too weak to model. IHP was observed in all ADAR deaminase domains in the asymmetric unit. To model the hydrated 8-azanebularine nucleotide in all RNAs, a CCP4 dictionary file for adenosine (A) was modified to replace the 6-amino group with hydrogen, to change atom 8 to nitrogen and to include an additional hydroxyl group off carbon 6. Additionally, an energy minimization calculated idealized structure was used to determine ideal bond angles and distances for the modified base of the hydrated 8-azanebularine (unpublished data, Professor Dean Tantillo, University of California- Davis). The refinement restraint dictionary file was edited to match these parameters. Expression and purification of hADAR2d for in vitro deamination kinetics Histidine-tagged human ADAR2 deaminase domain (hADAR2d) and hADAR2d mutant proteins were expressed in S. cerevisiae strain BCY123 and purified as described above with the following modifications. Cell lysate was 0.45 µm filtered after centrifugation and loaded 3 times through 5 mL Ni-NTA Superflow (Qiagen) at 3 mL/min. Washes of 50 ml with buffer 1, 2 and 3 at 4 mL/min followed by elution with 35 mL gradient from Buffer 3 to elution buffer. Selected elution fractions from the Ni NTA column were pooled and loaded at 0.5 mL/min on 1 mL HiTrap Heparin HP column from GE. The column was washed with 10 mL of Heparin 1 buffer at 0.5 mL/min and eluted with a 12 mL gradient from Heparin 1 to Heparin 2 buffer. Selected elution fractions from the Heparin column were pooled and concentrated to \u003c300 µL in 10,000 MWCO Amicon Ultra 4 centrifugal filter at 6500 RCF and 4 °C. TEV protease cleavage and gel filtration steps were omitted. Buffer exchange was accomplished via 3 rounds of concentration to \u003c300 µL followed by addition of 3 mL of Storage buffer. After final concentration, protein concentrations were determined using BSA standards visualized by SYPRO Orange staining on SDS-polyacrylamide gels and the purified proteins were stored at −70 °C. Site-directed mutagenesis Mutagenesis of hADAR2 catalytic domain was carried out via PCR site directed mutagenesis using the primers listed in Supplementary Table 2. All primers were purchased from IDT and PAGE purified as described above but were desalted by phenol chloroform extraction, ethanol precipitation and 70% ethanol wash instead of C18 Sep-Pack. Sequences for mutant plasmids were confirmed by Sanger sequencing. Preparation of hGLI1 splint-ligated RNA Oligonucleotides were purified as described above but were desalted by phenol chloroform extraction, ethanol precipitation and 70% ethanol wash. The 3’ GLI1 top strand 12 mer RNAs were radiolabeled with [γ-32P] at the 5’ end with T4 PNK as described previously. Labeled 3’ GLI1 top strand 12 mer RNAs were ligated as previously described to give an internally labeled RNA. The splint ligation products were PAGE purified as described above. Labeled RNAs were hybridized with the complementary GLI1 bottom strand 24 mer RNA (Y is chosen based on the identity of X, see Fig. 3b) in 10 mM Tris-HCl, 0.1 mM EDTA pH 7.5 and 100 mM NaCl. See Supplementary Table 2 for RNA sequences. In vitro deamination kinetics with internally32P labeled substrates Deamination kinetics of analog containing RNAs were carried out as previously described but with the following modifications. The final reaction volume was 10 µL. Final enzyme concentration was 300 nM. Final RNA concentration was 10 nM. Final reaction conditions were: 16 mM Tris HCl pH 7.4, 3.3% glycerol, 1.6 mM EDTA, 0.003% Nonidet NP-40, 60 mM KCl, 7.1 mM NaCl, 0.5 mM DTT, 160 units/mL Rnasin, 1 µg/mL yeast tRNA. Reactions were quenched by adding 190 µL 95 °C nuclease-free water followed by incubation at 95 °C for 5 min or by 10 µL 0.5% SDS at 95 °C followed by incubation at 95 °C for 5 min. Each experiment was carried out in triplicate, and the rate constants reported in the text are average values ± standard deviations. Sequences of RNAs used to prepare internally labeled substrates are shown in Supplementary Table 2. For comparison of hADAR2-D mutants, deamination kinetics were carried out as described above with the following modifications. Final reaction conditions were 300 nM hADAR2d, 10 nM RNA, 16 mM Tris HCl pH 7.4, 3.6% glycerol, 1.6 mM EDTA, 0.003% Nonidet NP-40, 60 mM KCl, 8.6 mM NaCl, 0.5 mM DTT, 160 units/mL Rnasin, 1 µg/mL yeast tRNA. EMSA analysis of radioactively labelled RNA Duplex RNAs containing 8-azanebularine and32P labeled were prepared as previously described. Samples containing 0.25 nM RNA and different concentrations of hADAR2d E488Q (128, 64, 32, 16, 8, 4, 2, 1, 0.5, 0.25 and 0 nM) were equilibrated in 20 mM Tris-HCl, pH 7, 6% glycerol, 0.5 mM DTT, 60 mM KCl, 20 mM NaCl, 0.1 mM BME, 1.5 mM EDTA, 0.003% NP-40, 160 units/ml RNasin, 100 µg/ml BSA and 1.0 µg/ml yeast tRNA for 30 min at 30 °C. Assay and data analysis were carried out as previously described. See Supplementary Table 2 for RNA sequences. In vitro transcription of RNA A truncation of hGLI1 mRNA incorporating 81 nucleotides upstream and 65 nucleotides downstream of the edited site was transcribed and purified as previously described. 3’ Nearest neighbor mutants of hGLI1 RNA were generated by site directed mutagenesis to generate G to A, G to C and G to U nearest neighbor mutants. A second site −32 bases from the edit site was mutated to maintain the original secondary structure of the RNA. See Supplementary Table 2 for primers used for mutagenesis. Deamination kinetics of transcribed hGLI1 RNAs Deamination kinetics of transcribed RNAs were carried out as previously described but with the following modifications. Final reaction volume was 20 µL. Final enzyme concentrations was 10 nM. Final RNA concentration was 2 nM. Final reaction conditions were: 17 mM Tris HCl pH 7.4, 5.0% glycerol, 1.6 mM EDTA, 0.003% Nonidet NP-40, 60 mM KCl, 15.6 mM NaCl, 0.5 mM DTT, 160 units/mL RNasin, 1 µg/mL yeast tRNA. Reactions were quenched by adding 10 µL 0.5% SDS at 95 °C followed by incubation at 95 °C for 5 min. cDNA was generated from RNA via RT-PCR, Sanger sequenced and quantified using SeqScanner 2 software from Applied Biosystems. The kobs (min−1) of each assay was calculated as described previously. Supplementary Material Accession codes. Coordinates for the hADAR2d-RNA complexes have been deposited in the Protein Data Bank (PDB) with PDB IDs of 5ED1 and 5ED2 for the hADAR2d-E488Q–Bdf2-C RNA and hADAR2d-E488Q–Gli1 RNA complexes, respectively, and 5HP2 and 5HP3 for the hADAR2d-WT–Bdf2-U RNA and hADAR2d-WT–Bdf2-C RNA complexes, respectively. Author Contributions J.M.T., M.M.M., A.I.S. and Y.Z. purified protein. K.J.P and J.M.T. designed and purified RNA for crystallography and characterized protein/RNA binding. M.M.M. and A.I.S. conducted crystallization trials. M.M.M and A.J.F collected diffraction data and solved/refined crystal structure. J.M.T., Y.Z. and J.H. measured enzyme reaction rates. K.T. synthesized 8-azanebularane phosphoramidite. J.M.T and A.I.S conducted mutagenesis. J.M.T., M.M.M., P.A.B. and A.J.F. analyzed the structures. P.A.B. wrote the initial manuscript draft. J.M.T, M.M.M, P.A.B and A.J.F edited the manuscript. References  RNA editing by adenosine deaminases that act on RNA Functions and regulation of RNA editing by ADAR deaminases ADAR1 regulates ARHGAP26 gene expression through RNA editing by disrupting miR-30b-3p and miR-573 binding Regulation of alternative splicing by RNA editing RNA editing changes the lesion specificity for the DNA repair enzyme NEIL1 A standardized nomenclature for adenosine deaminases that act on RNA A to I RNA Editing and Human Disease Adenosine-to-inosine RNA editing and human disease Mice with altered serotonin 2C receptor RNA editing display characteristics of Prader-Willi syndrome Mutations in ADAR1 cause Aicardi-Goutieres syndrome associated with a type I interferon signature Mutations of the RNA-specific adenosine deaminase gene (DSRAD) are involved in dyschromatosis symmetrica hereditaria Seven novel mutations of the ADAR gene in Chinese families and sporadic patients with disychromatosis symmetrica hereditaria (DSH) Recoding RNA editing of AZIN1 predisposes to hepatocellular carcinoma RNA editing enters the limelight in cancer RNA editing of the GLI1 transcription factor modulates the output of Hedgehog signaling ADAR proteins: Structure and Catalytic Mechanism Genome-wide identification of human RNA editing sites by parallel DNA capturing and sequencing A transition state analogue for an RNA-editing reaction Recognition of Duplex RNA by the Deaminase Domain of the RNA editing Enyzme ADAR2 Inositol Hexakisphosphate Is Bound in the ADAR2 Core and Required for RNA editing Mechanistic Insights into edtiing-site specificity of ADARs RNA-Seq Analysis Identifies A Novel Set of Editing Substrates for Human ADAR2 Present in Saccharomyces cerevisiae Substrate recognition by ADAR1 and ADAR2 HhaI methyltransferase flips its target base out of the DNA helix HhaI DNA Methyltransferase Uses the Protruding Gln237 for Active Flipping of Its Target Cytosine Cobalt hexamine induced tautomeric shift in Z-DNA: the structure of d(CGCGCA).d(TGCGCG) in two crystal forms A nucleotide-flipping mechanism from the structure of human uracil-DNA glycosylase boudn to DNA Structural basis for recognition and repair of the endogenous mutagen 8-oxoguanine in DNA Crystal structure of a human alkylbase-DNA repair enzyme complexed to DNA: mechanisms for nucleotide flipping and base excision Recent advances in the structural mechanisms of DNA glycosylases Predicting sites of ADAR editing in double stranded RNA N2-Modified 2-aminopurine ribonucleosides as minor-groove-modulating adenosine replacements in duplex RNA Base flipping Cocrystal Structure of a tRNA psi55 Pseuduridine Synthase: Nucleotide Flipping by an RNA-modifying Enyzme Base excision repair initiation revealed by crystal structures and binding kinetics of human uracil-DNA glycosylase with DNA Crystal structures of restrictocin-inhibitor complexes with implications for RNA recognition and base flipping Crystal Structure of an Initiation Factor Bound to the 30S Ribosomal Subunit Structure, Dynamics, and Elasticity of Free 16S rRNA Helix 44 Studied by Molecular Dynamics Simulations Chemical trapping and crystal structure of a catalytic tRNA guanine transglycosylase covalent intermediate Calculation of Relative Hydration Free Energy Differences for Heteroaromatic Compounds: Use in the Design of Adenosine Deaminase and Cytidine Deaminase Inhibitors The Solution Structure of the ADAR2 dsRBM-RNA Complex Reveals a Sequence-Specific Readout of the Minor Groove Proteins binding to duplexed RNA: one motif, multiple functions The RNA-editing enzyme ADAR1 controls innate immune responses to RNA Online methods references Large-Scale Overexpression and Purification of ADARs from Saccharomyces cerevisiae for Biophysical and Biochemical Studies Matching Active Site Structure to Substrate Analog for an RNA Editing Reaction XDS Phaser crystallographic software Towards automated crystallographic structure refinement with phenix.refine Click Modification of RNA at Adenosine: Structure and Reactivity of 7-Ethynyl- and 7-Triazolyl-8-aza-7-deazaadenosine in RNA Human ADAR2 and modified RNAs for crystallography a, Domain map for human ADAR2 b, ADAR reaction showing intermediate and 8-azanebularine (N) hydrate that mimics this structure c, Duplex RNAs used for crystallization. Bdf2 duplex sequence is derived from an editing site found in S. cerevisiae Bdf2 mRNA and Gli1 duplex has sequence surrounding the human Gli1 mRNA editing site. Italics indicate nucleotides added for duplex stability. Structure of hADAR2d E488Q bound to the Bdf2-C RNA duplex at 2.75 Å resolution a, View of structure perpendicular to the dsRNA helical axis. Colors correspond to those in Figs. 1a and 1c; flipped out base N is highlighted red, zinc in grey space-filling sphere, Q488 in yellow, previously disordered aa454–477 loop in green and inositol hexakisphosphate (IHP) in space filling. A transparent surface is shown for the hADAR2d protein. b, View of structure along the dsRNA helical axis. c, Summary of the contacts between hADAR2d E488Q and the Bdf2-C RNA duplex. ADAR recognition of the flipped out and orphaned nucleotides a, Contacts to the editing site nucleotide (N) in the active site. Colors correspond to those in Figs. 1 and 2. b, Orphan nucleotide recognition in the hADAR2d E488Q–Bdf2-C complex. c, Orphan nucleotide recognition in the hADAR2d WT–Bdf2-U complex. Other ADAR-induced changes in RNA conformation a, hADAR2d shifts the position of U11-A13’ base pair from ideal A-form RNA helix (yellow). b, Overlay of Bdf2 duplex RNA and idealized A form duplex of same sequence (yellow) illustrating kink in strand and widening of major groove opposite editing site induced by hADAR2d. c, Unusual “wobble” A13’-U11 interaction in the hADAR2d WT–Bdf2-U complex shown in stick with H-bond indicated with yellow dashes and distances shown in Å. The position of this base pair in the hADAR2d E488Q–Bdf2-C duplex is shown in wire with H-bonds shown with gray dashes. Interactions with editing site nearest neighbor nucleotides a, The minor groove edge of the U11-A13’ base pair from the Bdf2 duplex approaches G489; model with a C-G pair at this position suggests a clash with the G 2-amino group b, RNA duplex substrates prepared with different 5’ nearest neighbor nucleotides adjacent to editing site indicated in red (2AP = 2-aminopurine). c, Comparison of deamination rate constants by hADAR2d at the editing site adenosine (red) for duplexes bearing different 5’ nearest neighbors; krel = kobs/(kobs for unmodified RNA). Error bars, s.d (n=3 technical replicates). d, hADAR2 S486 backbone H-bond with 3’ G 2-amino group; e, RNA duplex substrates prepared with different 3’ nearest neighbor nucleotides adjacent to editing site indicated in red (I = inosine, N2MeG = N2-methylguanosine, 2AP = 2-aminopurine). f, Comparison of deamination rate constants by hADAR2d at the editing site adenosine (red) for duplexes bearing different 3’ nearest neighbors. krel = kobs/(kobs for unmodified RNA). Error bars, s.d (n=3 technical replicates). * indicates no reaction product observed. RNA-binding loops in the ADAR catalytic domain a, hADAR2 residues that contact phosphodiester backbone near 5’ end of unedited strand. b, Location of mutations introduced at protein-RNA interface. c, Comparison of deamination rate constants of the different hADAR2d mutants (Log scale). krel = kobs for mutant/kobs for WT. Error bars, s.d (n=3 technical replicates). d, Sequence alignment of ADAR2s (A2) and ADAR1s (A1) from different organisms with different levels of conservation colored (Yellow: conserved in all ADAR1s and ADAR2s, red: conserved in ADAR2s, blue: conserved in ADAR1s. e, Interaction of the ADAR-specific RNA-binding loop near the 5’ end of the edited strand. Colors as in d, white: not conserved, flipped out base is shown in pink. Data Processing and Refinement Statistics. Complex\tADAR2-D_E488Q:BDF2-C23mer\tADAR2-D_E488Q:GLI123mer\tADAR2-D_wt:BDF2-U23mer\tADAR2-D_wt:BDF2-C23mer\t \tData Collection\t\t\t\t\t \tSynchrotron (Beamline)\tAPS (24-ID-C)\tSSRL (12-2)\tSSRL (12-2)\tSSRL (12-2)\t \tWavelength (Å)\t0.9792\t0.9795\t0.9795\t0.9795\t \tSpace Group\tP212121\tP212121\tP212121\tP212121\t \tCell dimensions\t\t\t\t\t \ta, b, c, (Å)\t82.36, 107.50, 121.10\t79.13, 81.61, 256.62\t81.32, 106.68, 120.49\t81.51, 107.21, 120.62\t \tα, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t \tResolution (Å)\t100 – 2.75 (2.82 – 2.75)\t50.0 – 2.95 (3.03 – 2.95)\t100 – 2.98 (3.06 – 2.98)\t100 – 3.09 (3.17 – 3.09)\t \tRmerge (%)\t7.0 (68.5)\t9.6 (135.1)\t14.4 (86.3)\t11.6 (68.7)\t \tCC1/2\t99.6 (66.3)\t99.7 (47.6)\t99.1 (75.3)\t99.3 (77.1)\t \tI/σ (I)\t11.57 (1.52)\t12.27 (1.16)\t10.00 (1.88)\t10.56 (1.75)\t \tCompleteness (%)\t96.5 (98.8)\t98.1 (98.9)\t97.3 (90.0)\t96.8 (89.1)\t \tRedundancy\t2.93 (3.00)\t5.19 (5.13)\t4.79 (4.56)\t3.31 (2.84)\t \tRefinement\t\t\t\t\t \t  Resolution (Å)\t2.75\t2.95\t2.98\t3.09\t \t  No of reflections (F\u003e0)\t27,153\t35,727\t21,376\t19,325\t \t  Rwork/ Rfree\t16.27/22.34\t18.79/20.75\t16.67 / 24.67\t16.29 / 23.79\t \tNo. of atoms\t\t\t\t\t \t  Protein\t6197\t6038\t6168\t6157\t \t  RNA\t973\t1950\t973\t973\t \t  Inositol Hexakisphosphate  (IHP)\t72\t72\t72\t72\t \t  Zn\t2\t2\t2\t2\t \t  Waters\t33\t0\t1\t1\t \tB factors\t\t\t\t\t \t  Protein\t68.46\t90.65\t63.63\t67.92\t \t  RNA\t88.24\t108.8\t69.70\t77.49\t \t  Inositol Hexakisphosphate  (IHP)\t47.10\t65.57\t44.23\t43.77\t \t  Zn\t48.47\t64.38\t38.25\t49.04\t \t  Waters\t48.11\tN/A\t43.66\t50.20\t \tr.m.s deviations\t\t\t\t\t \t  Bond lengths (Å)\t0.010\t0.007\t0.009\t0.010\t \t  Bond angles (°)\t1.342\t0.885\t1.359\t1.365\t 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+[{"sourceid":"4918766","sourcedb":"","project":"","target":"","text":"Mechanism of extracellular ion exchange and binding-site occlusion in the sodium-calcium exchanger Na+/Ca2+ exchangers utilize the Na+ electrochemical gradient across the plasma membrane to extrude intracellular Ca2+, and play a central role in Ca2+ homeostasis. Here, we elucidate their mechanisms of extracellular ion recognition and exchange through a structural analysis of the exchanger from Methanococcus jannaschii (NCX_Mj) bound to Na+, Ca2+ or Sr2+ in various occupancies and in an apo state. This analysis defines the binding mode and relative affinity of these ions, establishes the structural basis for the anticipated 3Na+:1Ca2+ exchange stoichiometry, and reveals the conformational changes at the onset of the alternating-access transport mechanism. An independent analysis of the dynamics and conformational free-energy landscape of NCX_Mj in different ion-occupancy states, based on enhanced-sampling molecular-dynamics simulations, demonstrates that the crystal structures reflect mechanistically relevant, interconverting conformations. These calculations also reveal the mechanism by which the outward-to-inward transition is controlled by the ion-occupancy state, thereby explaining the emergence of strictly-coupled Na+/Ca2+ antiport. Introduction Na+/Ca2+ exchangers (NCX) play physiologically essential roles in Ca2+ signaling and homeostasis. NCX catalyzes the uphill extrusion of intracellular Ca2+ across the cell membrane, by coupling this process to the downhill permeation of Na+ into the cell, with a 3 Na+ to 1 Ca2+ stoichiometry. This reaction is, however, inherently reversible, its direction being dictated only by the transmembrane electrochemical ion gradients . The mechanism of NCX proteins is therefore highly likely to be consistent with the alternating-access model of secondary-active transport. The basic functional unit for ion transport in NCX consists of ten membrane-spanning segments, comprising two homologous halves. Each of these halves contains a highly conserved region, referred to as α-repeat, known to be important for ion binding and translocation; in eukaryotic NCX, the two halves are connected by a large intracellular regulatory domain, which is absent in microbial NCX (Supplementary Fig. 1). Despite a long history of physiological and functional studies, the molecular mechanism of NCX has been elusive, owing to the lack of structural information. Our recent atomic-resolution structure of NCX_Mj from Methanococcus jannaschii provided the first view of the basic functional unit of an NCX protein. This structure shows the exchanger in an outward-facing conformation and reveals four putative ion-binding sites, denominated internal (Sint), external (Sext), Ca2+-binding (SCa) and middle (Smid), clustered in the center of the protein and occluded from the solvent (Fig. 1a-b). With similar ion exchange properties to those of its eukaryotic counterparts, NCX_Mj provides a compelling model system to investigate the structural basis for the specificity, stoichiometry and mechanism of the ion-exchange reaction catalyzed by NCX. In this study, we set out to determine the structures of outward-facing wild-type NCX_Mj in complex with Na+, Ca2+ and Sr2+, at various concentrations. These structures reveal the mode of recognition of these ions, their relative affinities, and the mechanism of extracellular ion exchange, for a well-defined, functional conformation in a membrane-like environment. An independent analysis based on molecular-dynamics simulations demonstrates that the structures capture mechanistically relevant states. These calculations also reveal how the ion occupancy state of the outward-facing exchanger determines the feasibility of the transition to the inward-facing conformation, thereby addressing a key outstanding question in secondary-active transport, namely how the transported substrates control the alternating-access mechanism. Results Extracellular Na+ binding The assignment of the four central binding sites identified in the previously reported NCX_Mj structure was hampered by the presence of both Na+ and Ca2+ in the protein crystals. To conclusively clarify this assignment, we first set out to examine the Na+ occupancy of these sites without Ca2+. Crystals were grown in 150 mM NaCl using the lipidic cubic phase (LCP) technique. The crystallization solutions around the LCP droplets were then slowly replaced by solutions containing different concentrations of NaCl and EGTA (Methods). X-ray diffraction of these soaked crystals revealed a Na+-dependent variation in the electron-density distribution at sites Sext, SCa and Sint, indicating a Na+ occupancy change (Fig. 1c). Occupancy refinement indicated two Na+ ions bind to Sint and SCa at low Na+ concentrations (Fig. 1c), with a slight preference for Sint (Table 1). Binding of a third Na+ to Sext occurs at higher concentrations, as no density was observed there at 10 mM Na+ or lower (Fig. 1c); Sext is however partially occupied at 20 mM Na+, and fully occupied at 150 mM (Fig. 1c). The Na+ occupation at SCa, compounded with the expected 3Na+:1Ca2+ stoichiometry, implies our previous assignment of the Smid site must be re-evaluated. Indeed, two observations indicate that a water molecule rather than a Na+ ion occupies Smid, as was predicted in a recent simulation study. First, the electron density at Smid does not depend significantly on the Na+ concentration. Second, the protein coordination geometry at Smid is clearly suboptimal for Na+ (Supplementary Fig. 1d). The water molecule at Smid forms hydrogen-bonds with the highly conserved Glu54 and Glu213 (Supplementary Fig. 1d), stabilizing their orientation to properly coordinate multiple Na+ ions at Sext, SCa and Sint. It can be inferred from this assignment that Glu54 and Glu213 are ionized, while Asp240, which flanks Smid (and is replaced by Asn in eukaryotic NCX) would be protonated, as indicated by the abovementioned simulation study. Na+-dependent conformational change The NCX_Mj structures in various Na+ concentrations also reveal that Na+ binding to Sext is coupled to a subtle but important conformational change (Fig. 2). When Na+ binds to Sext at high concentrations, the N-terminal half of TM7 is bent into two short helices, TM7a and TM7b (Fig. 2a). TM7b occludes the four central binding sites from the external solution, with the backbone carbonyl of Ala206 coordinating the Na+ ion (Fig. 2b-d). However, when Sext becomes empty at low Na+ concentrations, TM7a and TM7b become a continuous straight helix (Fig. 2a), and the carbonyl group of Ala206 retracts away (Fig. 2b-d). TM7a also forms hydrophobic contacts with the C-terminal half of TM6. These contacts are absent in the structure with Na+ at Sext, in which there is an open gap between the two helices (Fig. 2b). This difference is noteworthy because TM6 and TM1 are believed to undergo a sliding motion, relative to the rest of the protein, when the transporter switches to the inward-facing conformation. The straightening of TM7ab also opens up a passageway from the external solution to Sext and Smid, while SCa and Sint remain occluded (Fig. 2d). Thus, the structures at high and low Na+ concentrations represent the outward-facing occluded and partially open states, respectively. This conformational change is dependent on the Na+ occupancy of Sext and occurs when Na+ already occupies Sint and SCa. Our crystallographic titration experiment indicates that the K1/2 of this Na+-driven conformational transition is ~20 mM. At this concentration, Sext is partially occupied and the NCX_Mj crystal is a mixture of both the occluded and partially open conformations. This structurally-derived Na+ affinity agrees well with the external Na+ concentration required for NCX activation in eukaryotes. The finding that the Na+ occupancy change from 2 to 3 ions coincides with a conformational change of the transporter also provides a rationale to the Hill coefficient of the Na+-dependent activation process in eukaryotic NCX. Extracellular Ca2+ and Sr2+ binding and their competition with Na+ To determine how Ca2+ binds to NCX_Mj and competes with Na+, we first titrated the crystals with Sr2+ (Methods). Sr2+ is transported by NCX similarly to Ca2+ , and is distinguishable from Na+ by its greater electron-density intensity. Protein crystals soaked with 10 mM Sr2+ and 2.5 mM Na+ revealed a strong electron-density peak at site SCa, indicating binding of a single Sr2+ ion (Fig. 3a). The Sr2+-loaded NCX_Mj structure adopts the partially open conformation observed at low Na+ concentrations. Binding of Sr2+, however, excludes Na+ entirely. Crystal titrations with decreasing Sr2+ or increasing Na+ demonstrated that Sr2+ binds to the outward-facing NCX_Mj with low affinity, and that it can be out-competed by Na+ even at low concentrations (Supplementary Note 1 and Supplementary Fig. 2a-b). Thus, in 100 mM Na+ and 10 mM Sr2+, Na+ completely replaced Sr2+ (Fig. 3a) and reverted NCX_Mj to the Na+-loaded, fully occluded state. Similar titration experiments showed that Ca2+ and Sr2+ binding to NCX_Mj are not exactly alike The electron density distribution from crystals soaked in high Ca2+ and low Na+, indicates that Ca2+ can bind to Smid as well as SCa, with a preference for SCa (Fig. 3b). Binding of Ca2+ to both sites simultaneously is highly improbable due to their close proximity, and at least one water molecule can be discerned coordinating the ion (Fig. 3b). The partial Ca2+ occupancy at Smid is likely caused by Asp240, which flanks this site and can in principle coordinate Ca2+. Previous functional and computational studies, however, indicate Asp240 becomes protonated during transport. Indeed, in most NCX proteins Asp240 is substituted by Asn, which would likely weaken or abrogate Ca2+ binding to Smid. SCa is therefore the functional Ca2+ site. Similarly to Sr2+, Ca2+ binds with low affinity to outward-facing NCX_Mj and can be readily displaced by Na+ (Supplementary Note 1 and Supplementary Fig. 2c). This finding is consistent with physiological and biochemical data for both eukaryotic NCX and NCX_Mj indicating that the apparent Ca2+ affinity is much lower on the extracellular than the cytoplasmic side. Specifically, our crystallographic titration assay indicates Ca2+ binds with sub-millimolar affinity, in good agreement with the external apparent Ca2+ affinities deduced functionally for cardiac NCX (Km ~ 0.32 mM) and NCX_Mj (Km ~ 0.175 mM). Taken together, these crystal titration experiments demonstrate that the four binding sites in outward-facing NCX_Mj exhibit different specificity: Sint and Sext are Na+ specific whereas SCa, previously hypothesized to be Ca2+ specific, can also bind Na+, confirming our earlier simulation study, as well as Sr2+; Smid can also transiently accommodate Ca2+ but during transport Smid is most likely occupied by water. The ion-binding sites in NCX_Mj can therefore accommodate up to three Na+ ions or a single divalent ion, and occupancy by Na+ and Ca2+ (or Sr2+) are mutually exclusive, as was deduced for eukaryotic exchangers. A structure of NCX_Mj without Na+ or Ca2+ bound An apo state of outward-facing NCX_Mj is likely to exist transiently in physiological conditions, despite the high amounts of extracellular Na+ (~150 mM) and Ca2+ (~2 mM). We were able to determine an apo-state structure of NCX_Mj, by crystallizing the protein at lower pH and in the absence of Na+ (Methods). This structure is similar to the partially open structure with two Na+ or either one Ca2+ or one Sr2+ ion, with two noticeable differences. First, TM7ab along with the extracellular half of the TM6 and TM1 swing further away from the protein core (Fig. 3c), resulting in a slightly wider passageway into the binding sites. Second, Glu54 and Glu213 side chains rotate away from the binding sites and appear to form hydrogen-bonds with residues involved in ion coordination in the fully Na+-loaded structure (Fig. 3d). Although the binding sites are thus fully accessible to the external solution (Fig. 3e), the lack of electron density therein indicates no ions or ordered solvent molecules. This apo structure might therefore represent the unloaded, open state of outward-facing NCX_Mj. Alternatively, this structure might capture a fully protonated state of the transporter, to which Na+ and Ca2+ cannot bind. Such interpretation would be consistent with the computer simulations reported below. Indeed, transport assays of NCX_Mj have shown that even in the presence of Na+ or Ca2+, low pH inactivates the transport cycle. Ion occupancy determines the free-energy landscape of NCX_Mj That secondary-active transporters are able to harness an electrochemical gradient of one substrate to power the uphill transport of another relies on a seemingly simple principle: they must not transition between outward- and inward-open conformations unless in two precise substrate occupancy states. NCX must be loaded either with 3 Na+ or 1 Ca2+, and therefore functions as an antiporter; symporters, by contrast, undergo the alternating-access transition only when all substrates and coupling ions are concurrently bound, or in the apo state. The reason why only specific occupancy states permit this transition in a given system, thereby determining its biological function, remains unclear. To examine this central question, we sought to characterize the conformational free-energy landscape of NCX_Mj and to examine its dependence on the ion-occupancy state, using molecular dynamics (MD) simulations. This computational analysis was based solely on the published structure of NCX_Mj, independently of the crystallographic studies described above. As it happens, the results confirm that the structures now available are representing interconverting states of the functional cycle of NCX_Mj, while revealing how the alternating-access mechanism is controlled by the ion-occupancy state. A series of exploratory MD simulations was initially carried out to examine what features of the NCX_Mj structure might depend on the ion-binding sites occupancy. Specifically, we first simulated the outward-occluded form, in the ion configuration we previously predicted, now confirmed by the high-Na+ crystal structure described above (Fig. 1b). That is, Na+ ions occupy Sext, SCa, and Sint, while D240 is protonated and a water molecule occupies Smid. The Na+ ion at Sext was then relocated from the site to the bulk solution (Methods), and this system was then allowed to evolve freely in time. The Na+ ions at SCa and Sint were displaced subsequently, and an analogous simulation was then carried out. These initial simulations revealed noticeable changes in the transporter, consistent with those observed in the new crystal structures. The most notable change upon displacement of Na+ from Sext was the straightening of TM7ab (Fig. 4a). When 3 Na+ ions are bound, TM7ab primarily folds as two distinct, non-collinear α-helical fragments, owing to the loss of the backbone carbonyl-amide hydrogen-bonds between F202 and A206, and T203 and F207 (Fig. 4b). This distortion occludes Sext from the exterior (Fig. 4d, 4h-i) and appears to be induced by the Na+ ion itself, which pulls the carbonyl group of A206 into its coordination sphere (Fig. 4g). With Sext empty, however, TM7ab forms a canonical α-helix (Fig. 4a-b, 4g), thereby creating an opening between TM3 and TM7, which in turn allows water molecules from the external solution to reach into Sext (Fig. 4e, 4h-i), i.e. the transporter is no longer occluded. Displacement of Na+ from SCa and Sint induces further changes (Fig. 4c). The most noticeable is an increased separation between TM7 and TM2 (Fig. 4f), previously brought together by concurrent backbone interactions with the Na+ ion at SCa (Fig. 4d-e). TM1 and TM6 also slide further towards the membrane center, relative to the outward-occluded state (Fig. 4c). Together, these changes open a second aqueous channel leading directly into SCa and Sint (Fig. 4f, Fig. 4h-i). The transporter thus becomes fully outward-open. To more rigorously characterize the influence of the ion-occupancy state on the conformational dynamics of the exchanger, we carried out a series of enhanced-sampling MD calculations designed to reversibly simulate the transition between the outward-occluded and fully outward-open states, and thus quantify the free-energy landscape encompassing these states (Methods). As above, we initially examined three occupancy states, namely with Na+ in Sext, SCa and Sint, with Na+ only at SCa and Sint, and without Na+. These calculations demonstrate that the Na+ occupancy state of the transporter has a profound effect on its conformational free-energy landscape. When all Na+ sites are occupied, the global free-energy minimum corresponds to a conformation in which the ions are maximally coordinated by the protein (Fig. 5a, 5c); TM7ab is bent and packs closely with TM2 and TM3, and so the binding sites are occluded from the solvent (Fig. 5b). At a small energetic cost, however, the transporter can adopt a metastable ‘half-open’ conformation in which TM7ab is completely straight and Sext is open to the exterior (Fig. 5a, 5b). The Na+ ion at Sext remains fully coordinated, but an ordered water molecule now mediates its interaction with A206:O, relieving the strain on the F202:O–A206:N hydrogen-bond (Fig. 5c). This semi-open conformation is nearly identical to that found to be the most probable when Na+ occupies only SCa and Sint (2 × Na+, Fig. 5a), demonstrating that binding (or release) of Na+ to Sext occurs in this metastable conformation. Interestingly, this doubly occupied state can also access conformations in which the second aqueous channel mentioned above, i.e. leading to SCa between TM7 and TM2 and over the gating helices TM1 and TM6, also becomes open (Fig. 5b-c). Crucially, though, the free-energy landscape for this partially occupied state demonstrates that the occluded conformation is no longer energetically feasible (Fig. 5a). Displacement of the two remaining Na+ ions from SCa and Sint further reshapes the free-energy landscape of the transporter (No ions, Fig. 5a), which now can only adopt a fully open state featuring the two aqueous channels (Fig. 5b-c). The transition to the occluded state in this apo state is again energetically unfeasible. From a mechanistic standpoint, it is satisfying to observe how the open and semi-open states are each compatible with two different Na+ occupancies, explaining how sequential Na+ binding to energetically accessible conformations (prior to those binding events) progressively reshape the free-energy landscape of the transporter; by contrast, the occluded conformation is forbidden unless the Na+ occupancy is complete. This processivity is logical since three Na+ ions are involved, but also implies that in the Ca2+-bound state, which includes a single ion, the transporter ought to be able to access all three major conformations, i.e. the outward-open state, in order to release (or re-bind) Ca2+, but also the occluded conformation, and thus the semi-open intermediate, in order to transition to the inward-open state. By contrast, occupancy by H+, which as mentioned are not transported, might be compatible with a semi-open state as well as with the fully open conformation, but should not be conducive to occlusion. To assess this hypothesis, we carried out enhanced-sampling simulations for the Ca2+ and H+-bound states of outward-facing NCX_Mj analogous to those described above for Na+ (see Supplementary Note 2 and Supplementary Fig. 3-4 for details on how the structures of the Ca2+-bound state was predicted). The calculated free-energy landscape for Ca2+-bound NCX_Mj confirms the hypothesis outlined above (1 × Ca2+, Fig. 6a): consistent with the fact that NCX_Mj transports a single Ca2+, the occluded, dehydrated conformation is one of the major energetic minima, but clearly the exchanger can also adopt the semi-open and open states that would be required for Ca2+ release and Na+ entry, via either of the aqueous access channels that lead to Sext and SCa (Fig. 6b-c). By contrast, protonation of Glu54 and Glu213 makes the occluded conformation energetically unfeasible, consistent with the fact that NCX_Mj does not transport protons; in this H+-bound state, though, the exchanger can adopt the semi-open conformation captured in the low pH, apo crystal structure (2 × H+, Fig. 6a-c). Taken together, this systematic computational analysis of outward-facing NCX_Mj clearly demonstrates that the alternating-access and ion-recognition mechanisms in this Na+/Ca2+ exchanger are coupled through the influence that the bound ions have on the free-energy landscape of the protein, which in turn determines whether or not the occluded conformation is energetically feasible. This occluded conformation, which is a necessary intermediate between the outward and inward-open states, and which entails the internal dehydration of the protein, is only attainable upon complete occupancy of the binding sites. Discussion The alternating-access hypothesis implicitly dictates that the switch between outward- and inward-open conformations of a given secondary-active transporter must not occur unless the appropriate type and number of substrates are recognized. This control mechanism is functionally crucial, as it precludes the backflow of the species that is transported uphill, and also prevents the dissipation of the driving electrochemical gradients. It is however also non-trivial: antiporters, for example, do not undergo the alternating-access transition without a cargo, but this is precisely how membrane symporters reset their transport cycles. Similarly puzzling is that a given antiporter will undergo this transition upon recognition of substrates of different charge, size and number. Yet, when multiple species are to be co-translocated, by either an antiporter or a symporter, partial occupancies must not be conducive to the alternating-access switch. Here, we have provided novel insights into this intriguing mechanism of conformational control through structural studies and quantitative molecular simulations of a Na+/Ca2+ exchanger. Specifically, our studies of NCX_Mj reveal the mechanism of forward ion exchange (Fig. 7). The internal symmetry of outward-facing NCX_Mj and the inward-facing crystal structures of several Ca2+/H+ exchangers indicate that the alternating-access mechanism of NCX proteins entails a sliding motion of TM1 and TM6 relative to the rest of the transporter. Here, we demonstrate that conformational changes in the extracellular region of the TM2-TM3 and TM7-TM8 bundle precede and are necessary for the transition, and are associated with ion recognition and/or release. The most apparent of these changes involves the N-terminal half of TM7 (TM7ab); together with more subtle displacements in TM2 and TM3, this change in TM7ab correlates with the opening and closing of two distinct aqueous channels leading into the ion-binding sites from the extracellular solution. Interestingly, the bending of TM7 associated with the occlusion of the ion-binding sites also unlocks its interaction with TM6, and thus enables TM6 and TM1 to freely slide to the inward-facing conformation. We anticipate that the intracellular ion-exchange process involves analogous conformational changes. The crystal structures of NCX_Mj reported here, with either Na+, Ca2+, Sr2+ or H+ bound, capture the exchanger in different conformational states. These states can only represent a subset among all possible, but they ought to reflect inherent preferences of the transporter, modulated by the experimental conditions. For example, in the crystal of NCX_Mj in LCP, the extracellular half of the gating helices (TM6 and TM1) form a lattice contact, which might ultimately restrict the degree of opening of the ion-binding sites in some cases (e.g. in the apo, low pH structure). Nonetheless, the calculated free-energy landscapes, derived without knowledge of the experimental data, reassuringly confirm that the crystallized structures correspond to mechanistically relevant, interconverting states. The simulations also demonstrate how this landscape is drastically re-shaped upon each ion-binding event. Indeed, we show that it is the presence or absence of the occluded state in this landscape that explains the antiport function of NCX_Mj and its 3Na+:1Ca2+ stoichiometry. We posit that a similar principle might govern the alternating-access mechanism in other transporters; that is, we anticipate that for both symporters and antiporters, it is the feasibility of the occluded state, encoded in the protein conformational free-energy landscape and its dependence on substrate binding, that ultimately explains their specific coupling mechanisms. In multiple ways, our findings provide an explanation for, existing functional, biochemical and biophysical data for both NCX_Mj and its eukaryotic homologues. The striking quantitative agreement between the ion-binding affinities inferred from our crystallographic titrations and the Km and K1/2 values previously deduced from functional assays has been discussed above. Consistent with that finding, mutations that have been shown to inactivate or diminish the transport activity of NCX_Mj and cardiac NCX perfectly map to the first ion-coordination shell in our NCX_Mj structures (Supplementary Fig. 4c-d). The crystallographic data also provides the long-sought structural basis for the ‘two-site’ model proposed to describe competitive cation binding in eukaryotic NCX, underscoring the relevance of these studies of NCX_Mj as a prototypical Na+/Ca2+ exchanger. Specifically, our crystal titrations suggest that, during forward Na+/Ca2+ exchange, sites Sint and SCa, which Ca2+ and Na+ compete for, can be grouped into one; Na+ binding to these sites does not require high Na+ concentrations, and two Na+ ions along with a water molecule (at Smid) are sufficient to displace Ca2+, explaining the Hill coefficient of ~2 for Na+-dependent inhibition of Ca2+ fluxes. The Sext site, by contrast, might be thought as an activation site for inward Na+ translocation, since this is where the third Na+ ion binds at high Na+ concentration, enabling the transition to the occluded state. Interestingly, binding of Ca2+ to Smid appears to be also possible, but available evidence indicates that this event transiently blocks the exchange cycle. Indeed, structures of NCX_Mj bound to Cd2+ or Mn2+, both of which inhibit transport, show these ions at Smid; by contrast, Sr2+ binds only to SCa, and accordingly, is transported by NCX similarly to calcium. Lastly, our theory that occlusion of NCX_Mj is selectively induced upon Ca2+ or Na+ recognition is consonant with a recent analysis of the rate of hydrogen-deuterium exchange (HDX) in NCX_Mj, in the presence or absence of these ions, in conditions that favor outward-facing conformations. Specifically, saturating amounts of Ca2+ or Na+ resulted in a noticeable slowdown in the HDX rate for extracellular portions of the α-repeat helices. We interpret these observations as reflecting that the solvent accessibility of the protein interior is diminished upon ion recognition, consistent with our finding that opening and closing of extracellular aqueous pathways to the ion-binding sites depend on ion occupancy state. In addition, the increased compactness of the protein tertiary structure in the occluded state would also slow down the dynamics of the secondary-structure elements, and thus further reduce the HDX rate. Our data would also explain the observation that the reduction in the HDX rate is comparable for Na+ and Ca2+, as well as the finding that the degree of deuterium incorporation remains non-negligible even under saturating ion concentrations. As the calculated free-energy landscapes show, Na+ and Ca2+ induce the occlusion of the transporter in a comparable manner, and yet the ion-bound states retain the ability to explore conformations that are partially or fully open to the extracellular solution, precisely so as to be able to unload and re-load the substrates. Methods Protein expression, purification and crystallization NCX_Mj was expressed, purified and crystallized as previously described. Briefly, the NCX_Mj gene with a C-terminal hexa-histidine tag was subcloned into the pQE60 vector and expressed in Escherichia coli BL21(DE3)plysS. Harvested cells were homogenized and incubated in buffer containing 50 mM HEPES pH 7.2, 50 mM NaCl, 12 mM KCl, 10 mM CaCl2, 40 mM DDM. After incubation at room temperature (RT) for 3.5 hours, the supernatant was collected by centrifugation and loaded onto a Talon Co2+ affinity column (Clontech). The non-specifically bound contaminates on the column were washed with buffer containing 50 mM HEPES pH 7.2, 50 mM NaCl, 12 mM KCl, 10 mM CaCl2, 15 mM imidazole, and 1 mM DDM. The bound NCX_Mj was eluted by increasing the imidazole concentration to 300 mM. The eluate was treated with thrombin to remove the hexa-histidine tag and dialyzed against 20 mM HEPES pH 7.2, 50 mM NaCl, 12 mM KCl, 10 mM CaCl2, and 1 mM DDM at RT overnight. After overnight digestion the sample was loaded onto a second Co2+ affinity column to remove any free hexa-histidine tag and contaminant proteins. NCX_Mj in the flow-through was collected and further purified by gel filtration using a Superdex-200 (10/300) column (GE Healthcare) in 20 mM HEPES pH 7.2, 50 mM NaCl, 12 mM KCl, 10 mM CaCl2 and 0.5 mM DDM. The purified protein was then concentrated to 40 mg/ml for crystallization. Native NCX_Mj was crystallized using the lipidic cubic phase (LCP) technique, as previously described. Concentrated NCX_Mj was first reconstituted into 1-oleoyl-rac-glycerol (Sigma) in a protein:lipid weight ratio of 1:1.5, using the two-syringe method. Protein-laden LCP droplets of 35 nL were dispensed onto Corning 96-well protein-crystallization plates and overlaid with 5 μL of precipitant solution containing 40-42% PEG 400, 100 mM MES pH 6.5, 100 mM NaAc. Crystals were observed after 48 hours and grew to full size after 2 weeks. The native crystals belong to space group P212121 with a cell dimension of a=49.5Å, b=72.9Å and c=96.2Å, and contain one subunit per asymmetric unit. As the LCP droplet accounts for less than 1% of the total crystallization volume, the salt composition in the crystallization condition was determined mainly by the overlaying solution, and estimated to have 150 mM Na+ (from MES buffer and NaAc) and 30 μM Ca2+ (from LCP droplet). In these concentration conditions (high Na+ and low Ca2+) Ca2+ does not bind to NCX_Mj (as shown in our crystallographic titration experiments) and thus this native crystal structure represents NCX_Mj in 150 mM Na+. The native crystals were used in all subsequent titration experiments to define low-Na+, Ca2+ and Sr2+-loaded structures. To obtain the apo crystal form, the protein was first purified in a solution containing 20 mM Hepes-Tris pH 7.2, 100 mM NMDG, 10 mM CaCl2 and 0.5 mM DDM. The crystals were obtained in LCP with crystallization solution containing 200 mM KAc, pH 4.0, 35% PEG400. The apo NCX_Mj crystals belong to space group C2 with a cell dimension of a=164.2Å, b=46.8Å, c=97.0 Å and β=106.2°, and contain two protein subunits per asymmetric unit. Crystal titrations Once the native crystals reached their full size, the crystallization solutions overlaying lipid/protein droplets were gradually replaced by titration solutions through multiple steps of solution exchange. In general, 2-3 μL of existing crystallization solutions (normally in 5 μL) were replaced by the same volume of titration solutions, followed by overnight equilibration. The same procedures were repeated 6-10 times until the ion components in the crystal drops reached the targeted concentrations. For titration experiments to define concentration-dependent Na+ binding, the titration solutions contained 100 mM MES-Tris pH 6.5, 44% PEG400, 10 mM EGTA and a 100 mM mixture of NaAc and CsAc, in the following proportions: 100 mM CsAc; 90 mM CsAc and 10 mM NaAc; 80 mM CsAc and 20 mM NaAc; and 100 mM NaAc. Note that Cs+ does not bind NCX proteins and is commonly used as a Na+ substituent to maintain the ionic strength of the solutions. As complete removal of Na+ would deteriorate the crystals, we had to maintain a minimum Na+ concentration of about 2.5 mM in the crystal drops. The final Na+ concentrations in this set of titration experiments were about 2.5, 10, 20 and 100 mM, respectively. It is worth noting that the observed Na+-dependent conformational change occurs while the proteins are in crystal form and embedded in lipid. In the titration experiments carried out to define the mode of divalent cation binding and competition with Na+, the soaking solutions contained 100 mM MES-Tris pH 6.5, 44% PEG400, 100 mM mixture of CsAc and NaAc and various concentrations of XCl2, where X=Ca2+ or Sr2+, in the following proportions: 100 mM CsAc and 10 mM XCl2; 100 mM CsAc and 1 mM XCl2; 100 mM CsAc and 0.1 mM XCl2; 90 mM CsAc, 10mM NaAc and 10mM XCl2; and 100 mM NaAc and 10 mM XCl2. After multiple steps of solution exchanges, the final soaking conditions contained 0.1, 1, or 10 mM of X2+ together with 2.5 mM Na+; or 10 mM X2+ together with 2.5, 10 or 100 mM Na+. Data collection and structure determination After soaking crystals were mounted on 100-μm Mitegen Microloops and frozen in liquid nitrogen. All diffraction data were collected at the Advanced Photon Source (APS) GM/CA-CAT beamlines 23ID-B or 23ID-D using a beam size of 35 μm × 50 μm. Data were processed and scaled using HKL2000 and the structures were determined by molecular replacement in PHASER using our previously published NCX_Mj structure (PDB code 3V5U) as a search model. Model building was completed using COOT and structure refinement was performed with PHENIX. The data sets from crystals soaked in solutions containing 2.5 to 100 mM Na+ were collected using an X-ray wavelength of 1.033Å; the crystal grown with 150 mM Na+, and those soaked with Ca2+ and Sr2+ solutions, were obtained with a wavelength of 0.9793 Å. Lastly, the data from the crystal grown at low pH with no Na+ or Ca2+ were collected with a 2.0-Å wavelength beam. The resulting statistics for data collection and refinement are shown in Tables 2-4. All structure figures were prepared in PyMOL (The PyMOL Molecular Graphics System, Version 1.5.0.4 Schrödinger, LLC.). The ion passageways in low- and high-Na+ structures as well as the apo state were analyzed using the program CAVER. Due to the variation in diffraction resolution and intensity among crystals, ion-occupancy comparisons were made on the basis of the diffraction data obtained in the titration experiments scaled against a common reference data before map calculation. The NCX_Mj crystal obtained with 2.5 mm Na+ only was used as the reference. Conventional molecular dynamics (MD) simulations Conventional (i.e. not enhanced) MD simulations were carried out with NAMD 2.7-2.9  at constant temperature (298 K), pressure (1 atm), and membrane surface area (~69 Å2 per lipid),and with periodic boundaries in all directions. All calculations used the standard CHARMM27/CMAP force field, except for NBFIX corrections for the interaction between carboxylate-oxygens and Na+ interaction or Ca2+ (Supplementary Note 3, Supplementary Fig. 5). Electrostatic interactions were calculated using PME with a real-space cut-off of 12 Å; the same cut-off distance was used for all van der Waals interactions. Five ion-occupancy states of the transporter were considered, namely with 3 Na+, 2 Na+, 2 H+ or 1 Ca2+, and with no ions bound; in all cases Asp240 is protonated. For the 3×Na+ state, we reanalyzed a 200-ns trajectory of NCX_Mj reported previously. NCX_Mj had been embedded in a POPC lipid membrane using GRIFFIN. The initial configuration of the 2×Na+ state was generated from an equilibrated configuration of 3×Na+ state, from which the Na+ ion at Sext was displaced by means of a slow alchemical transformation that annihilates the bound ion and recreates it in the bulk solution (in the same simulation box). The resulting 2×Na+ state was simulated for 250 ns. Similarly, the state with no Na+ bound was generated from an equilibrated configuration of the 2×Na+ state, from which the remaining Na+ ions were displaced; this state was again simulated for 250 ns. For the 2×H+ state, an initial configuration was generated from an equilibrated configuration of the 3×Na+ state, by gradually annihilating the Na+ ions from the binding sites and creating protonated E54 and E213 side chains; concurrently, acetic acid molecules in the bulk solution (in the same simulation box) were deprotonated and Na+ ions introduced. A second initial configuration of the 2×H+ state was obtained from an equilibrated configuration of the simulation with no ions bound, by slowly transforming deprotonated E54 and E213 into their protonated form, while doing the opposite to acetic acid molecules in the bulk solution. These two initial configurations of the 2×H+ state were then equilibrated for 800 ns. All annihilation/creation simulations were carried out using the FEP module of NAMD; and comprised 32-50 intermediate simulations of 400 ps each, for each transformation. A soft-core van der Waals potential with a radius-shifting coefficient of 2 Å2 was used. The annihilated Na+ ions were confined within their corresponding binding sites using flat-bottom distance restraints. Specifically, the Na+ in Sext was concurrently maintained within 4 Å of E54:Cδ, A206:C, S77:Cβ, T209:Cβ and S210:Cβ. The Na+ ions in SCa and Sint were concurrently kept within 4 Å of the E213:Cδ and A47:C, respectively. The Na+ ions and acetic acid molecules in the bulk solution were kept at a distance greater than 37 Å from the membrane center. Finally, the initial configuration of the Ca2+ state was generated on the basis of the published NCX_Mj X-ray structure by placing Ca2+ in the SCa site and two water molecules coordinating Ca2+ at and near the Smid site, so as to satisfy the expected coordination geometry (see Supplementary Note 2, Supplementary Fig. 3-4). This configuration was initially equilibrated through a series of simulations in which RMSD-based restraints of gradually diminishing strength were applied to the protein Cα atoms as well the side-chains involved in Ca2+ coordination. A 250 ns equilibration was then carried without any restraints. Enhanced-sampling MD simulations Free-energy landscapes were calculated using Bias-Exchange Well-Tempered Metadynamics (BE-WT-MetaD), using GROMACS4.5.5/PLUMED. The force-field and simulation conditions were equivalent to those employed in the unbiased MD simulations. The accumulated simulation time for each of the ion-occupancy states studied was 1.6 μs. Each of these calculations consisted of 16 concurrent, interdependent simulations (or replicas); in 15 of these replicas, a WT-MetaD biasing potential was applied on different subsets of collective variables, as specified below, while the remaining replica was unbiased. Attempts to exchange coordinate configurations among replicas were made every 2-5 ps, using the Metropolis criterion. The inputs for each calculation were equilibrated configurations extracted from the unbiased MD simulations. The choice of collective variables to be biased in the BE-WT-MetaD simulations was also based on analysis of the unbiased MD trajectories (Fig. 4g-i). Specifically, to enhance the reversible opening and closing of the water channels reaching from the extracellular bulk solution into either the Sext or the SCa binding sites, we employed the following time-dependent collective variable (Fig. 4h-i):  where ri denotes the distance between the oxygen atom of each water molecule in the system i and the center of the binding site considered (V1 for Sext,V2 for SCa), and β is 10-100 nm. When the binding site was occupied, the ion was used to define its center. If the site was empty, its center was defined as the center-of-mass of the oxygen atoms coordinating the ion if bound. Bound water molecules at or near Smid (coordinating the Na+ or Ca2+ ions) were not considered. To enhance the reversible formation and disruption of selected backbone hydrogen bonds in TM7ab (Fig. 4g), we used an analogous collective variable:  In this case, the index i denotes atoms P202:O and T203:O, while the index j denotes atoms A206:N and P207:N. To preclude the artificial unraveling of TM7ab driven by this bias, an upper-bound V3max equal to 0.7 nm was imposed with a boundary potential of the form k (V3(t) – V3max) if V3(t) \u003e V3max, where k = 105 nm−4 kJ/mol. In addition, to control the bending and straightening of TM7ab more globally, we used the following path-collective variables:  where d1 and d2 denote the mean-square-differences between the conformation of TM7ab and either the straight or bent conformations, respectively, and λ = 100 nm−2. Note that V 4 is by definition confined between a lower (V4 ~ 1) and upper bound (V4 ~ 2); to confine the exploration of V5, an upper value V5max of 0.020-0.025 nm2 was imposed with boundary potential of the form k (V5(t) – V5max) if V5(t) \u003e V5max, where k = 1011 nm−8 kJ/mol. The mean-square-differences d1 and d2 comprise the backbone atoms of residues 198-211 as well as the side chain carbon atoms of residues P207 and L211. A boundary potential was also applied to confine the ions and water molecule bound to Sext, SCa, Sint and Smid to their corresponding binding sites. Specifically, the variable confined was:  where ri denotes the distance between the ion and each of its coordinating oxygen atoms; r0 was set to 0.24 nm for the Na+ ions, and to 0.30 nm for Ca2+. Note that the upper-bound value of Vc is, by definition, approximately the coordination number in the bound state, whereas Vc becomes 0 as the ion becomes unbound. For the Na+ ion bound to Sext and Sint, therefore, a lower bound value Vcmin = 4.3 was imposed with potential of the form k (Vc(t) – Vcmin) when Vc(t) \u003c Vcmin, where k = 2500 kJ/mol. An analogous restraint was used for the Na+ ion at SCa, with Vcmin = 4.75. Similarly, for the Ca2+ ion at SCa a lower bound value Vcmin = 7.4 was imposed with a potential of the form k (Vc(t) – Vcmin) when Vc(t) \u003c Vcmin, with k = 400 kJ/mol. Note that these restraints do not perturb the chemical structure of the ion-coordination sphere when the ion is bound, i.e. Vc(t) \u003e Vcmin. The displacement of the bound water molecules in the ion-coordination sphere by equivalent water molecules in the solvent was prevented similarly. The specific sets of collective variables biased in each of the replica WT-MetaD simulations, as well as further details on the biasing potentials introduced, are specified in Supplementary Table 1. Derivation of conformational free-energy landscapes To translate the data gathered in the BE-WT-MetaD simulations into conformational free-energy landscapes, we sought to identify a low-dimensional representation of the data that is nevertheless also intuitive and representative. We ultimately settled on two structure-based descriptors of the degree of opening of the each of the aqueous channels leading to the ion-binding sites (Fig. 4), defined as:  where rij denotes a set of pairwise distances for specific Cα atoms in the protein, for a given simulation snapshot. For S1, index i refers to the Cα of residues 198 to 209 in TM7, while index j refers to those in residues 66 to 80 and 290 to 297, in TM3 and TM10, respectively. For S2, index i refers to the Cα of residues 51 to 64 in TM2, while index j refers to those in residues 177 to 193 and 198 to 209, in TM6 and TM7, respectively. Therefore, S1 describes the effective separation between TM7 and TM3/TM10, on the extracellular half of the protein, and thus reports on the accessibility to the Sext site. Analogously, S2 measures the separation between TM2 and TM6-TM7, also on the extracellular side, and thus reports on the accessibility to SCa. The conformational free energy of NCX_Mj as a function of S1 and S2 was then computed for each ion-occupancy state separately (Figs. 5a, 6a). These landscapes were obtained through reweighting of the biased probability distribution from the BE-WT-MetaD sampling, using the WHAM method; through this approach we combine the statistics gathered in all replicas, and can consider alternate free-energy projections. To correct the landscape calculated for the Ca2+-bound state (Fig. 6a) on account of the excess amount of charge transferred from the ion to the protein (Supplementary Note 4, Supplementary Fig. 6), we reprocessed all the sampling obtained during the original BE-WT-MetaD simulations, introducing in the WHAM equations a re-weighting factor w for each configuration X:  where U denotes the ‘uncorrected’ CHARMM27/NBFIX potential-energy function, and Uc denotes the corrected function. To calculate Uc (X), the charge of Ca2+ was reduced to +1.8e from its standard value of +2e, and the difference was distributed among the surrounding protein residues (as specified in Supplementary Note 4). To minimally alter the charge-distribution used in the original CHARMM27 force-field, the charge added to each protein atom was proportional to the absolute value of its uncorrected charge. The statistical errors for all free-energy landscapes are provided in Supplementary Fig. 7. Derivation of representative structures and water-density maps Representative structures and water-density iso-surfaces (Fig. 5b-c, Fig. 6b-c) were derived for each BE-WT-MetaD simulation by clustering all sampling in the multi-dimensional space of V1, V2, and V4 (Eq. 1-3), plus a descriptor S3 of the proximity between TM6-TM7 and TM2-TM3/TM10, on the extracellular side of the protein. More precisely:  where r0 = 7.5 Å, and rij denotes a specific set of pairwise Cα distances, for a given simulation snapshot. Similarly to S1 and S2 (Eq. 6), index i refers to Cα atoms in the extracellular halves of TM6 and TM7, while index j refers to Cα atoms in the extracellular halves of TM2, TM3, and TM10. We thus obtained ~2,000 clusters for each of the simulation systems (using RMSD cut-off values of 1.3 Å for V1 and V2, 0.1 Å for V4; and 6.25 Å for S3). Using the WHAM equations, we calculated the relative free energy of each of these clusters, and then identified the major basins in this space with the MCL method (with p = 1.4.) Water occupancy maps were calculated for each of these major free-energy basins, only using the sampling gathered by the unbiased replicas. Supplementary Material Author Contributions J.L. and Y.J. designed the experimental studies and analyzed the resulting data. J.L., C.L. and Y.H. performed the experimental research. F.M and J.D.F.G. designed the computational research and analyzed the corresponding data. F.M. performed the computational work. J.L., Y.J. F.M. and J.D.F.G. wrote the paper. The authors declare no competing financial interests. Accession codes Atomic coordinates and structural factors have been deposited in the Protein Data Bank, with accession numbers 5HWX, 5HWY, 5HXC, 5HXE, 5HYA, 5HXH, 5HXS and 5HXR, as specified in Tables 2-4. References Sodium/calcium exchange: its physiological implications. Sodium/calcium exchanger: influence of metabolic regulation on ion carrier interactions. Calcium signaling. Calcium signalling: dynamics, homeostasis and remodelling. Unitary cardiac Na+, Ca2+ exchange current magnitudes determined from channel- like noise and charge movements of ion transport. Charge movement during Na+ translocation by native and cloned cardiac Na+/Ca2+ exchanger. The stoichiometry of the cardiac sodium-calcium exchange system. Sodium-calcium exchange and calcium-calcium exchange in internally dialyzed squid giant axons. Na/Ca exchange in barnacle muscle cells has a stoichiometry of 3 Na+/1 Ca2+. Na-Ca exchange current in mammalian heart cells. Multiple transport modes of the cardiac Na+/Ca2+ exchanger. Simple allosteric model for membrane pumps. Regulation and deregulation of cardiac Na+-Ca2+ exchange in giant excised sarcolemmal membrane patches. Initial localization of regulatory regions of the cardiac sarcolemmal Na+-Ca2+ exchanger. Sodium-calcium exchange: a molecular perspective. Mutation of amino acid residues in the putative transmembrane segments of the cardiac sarcolemmal Na+-Ca2+ exchanger. Toward a topological model of the NCX1 exchanger. Transmembrane segment packing of the Na+/Ca2+ exchanger investigated with chemical cross-linkers. The cation/Ca2+ exchanger superfamily: phylogenetic analysis and structural implications. Structural insight into the ion-exchange mechanism of the sodium/calcium exchanger. Functional asymmetry of bidirectional Ca2+-movements in an archaeal sodium- calcium exchanger (NCX_Mj). Sodium recognition by the Na+/Ca2+ exchanger in the outward-facing conformation. Effects of internal and external cations and of ATP on sodium- calcium and calcium-calcium exchange in squid axons. Effects of divalent and trivalent cations on Na+-Ca2+ exchange in cardiac sarcolemmal vesicles. Asymmetrical properties of the Na-Ca exchanger in voltage-clamped, internally dialyzed squid axons under symmetrical ionic conditions. Steady-state and dynamic properties of cardiac sodium-calcium exchange. Ion and voltage dependencies of the transport cycle. Sodium-calcium exchange current. Dependence on internal Ca2+ and Na+ and competitive binding of external Na+ and Ca2+. Structural basis for alternating access of a eukaryotic calcium/proton exchanger. Structural basis for the counter-transport mechanism of a H+/Ca2+ exchanger. Crystal structure of Ca2+/H+ antiporter protein YfkE reveals the mechanisms of Ca2+ efflux and its pH regulation. Asymmetric preorganization of inverted pair residues in the sodium-calcium exchanger. The Na+/Ca2+ exchanger NCX1 has oppositely oriented reentrant loop domains that contain conserved aspartic acids whose mutation alters its apparent Ca2+ affinity. The cardiac Na+-Ca2+ exchanger has two cytoplasmic ion permeation pathways. Mutational analysis of the alpha-1 repeat of the cardiac Na+-Ca2+ exchanger. Competitive interactions of sodium and calcium with the sodium-calcium exchange system of cardiac sarcolemmal vesicles. Crystallizing membrane proteins using lipidic mesophases. Processing of X-ray diffraction data collected in oscillation mode. Phaser crystallographic software. Coot: model-building tools for molecular graphics. PHENIX: a comprehensive Python-based system for macromolecular structure solution. CAVER: a new tool to explore routes from protein clefts, pockets and cavities. Scalable molecular dynamics with NAMD. All-atom empirical potential for molecular modeling and dynamics studies of proteins. Extending the treatment of backbone energetics in protein force fields: Limitations of gas-phase quantum mechanics in reproducing protein conformational distributions in molecular dynamics simulations. GRIFFIN: A versatile methodology for optimization of protein-lipid interfaces for membrane protein simulations. A bias-exchange approach to protein folding. Metadynamics with adaptive gaussians. GROMACS 4: Algorithms for highly efficient, load-balanced, and scalable molecular simulation. PLUMED: A portable plugin for free-energy calculations with molecular dynamics. From A to B in free energy space. The Weighted Histogram Analysis Method for free-energy calculations on biomolecules .1. The method. METAGUI: A VMD interface for analyzing metadynamics and molecular dynamics simulations. Two-state dynamics of the SH3-SH2 tandem of Abl kinase and the allosteric role of the N-cap. Peptide folding: When simulation meets experiment. Faraldo-Gómez, J.D. Atomic-resolution dissection of the energetics and mechanism of isomerization of hydrated ATP-Mg through the SOMA string method. An efficient algorithm for large-scale detection of protein families. Na+ binding to outward-facing NCX_Mj. (a) Overall structure of native outward-facing NCX_Mj from crystals grown in 150 mM Na+. N- and C-terminal halves are colored yellow and cyan, respectively. Colored spheres represent the bound Na+ (green) and water (red). (b) Structural details and definition of the four central binding sites. Only residues flanking these sites are shown for clarity (same for all other figures). The electron density (grey mesh, 1.9 Å Fo-Fc ion omit map contoured at 4σ) at Smid was modeled as water (red sphere) and those at Sext, SCa and Sint as Na+ ions (green spheres). Further details are shown in Supplementary Fig. 1. (c) Concentration-dependent change in Na+ occupancy (see also Table 1). All Fo – Fc ion-omit maps are calculated to 2.4 Å and contoured at 3σ for comparison. The displacement of A206 reflects the [Na+]-dependent conformational change from the partially open to the occluded state (observed at low and high Na+ concentrations, respectively). At 20 mM Na+, both conformations co-exist. No significant changes were observed in the side-chains involved in ion or water coordination at the SCa, Sint and Smid sites. Na+-occupancy dependent conformational change in NCX_Mj. (a) Superimposition of the NCX_Mj crystal structures obtained in high (100 mM, cyan cylinders) and low (10 mM, brown cylinders) Na+ concentrations. (b) Close-up view of the interface between TM6 and TM7ab in the NCX_Mj structures obtained at high and low Na+ concentrations (top and lower panels, respectively). Residues forming van-der-Waals contacts in the structure at low Na+ concentration are shown in detail. (c) Close-up view of the Na+-binding sites. The vacant Sext site in the structure at low Na+ concentration is indicated with a white sphere. Residues surrounding this site are also indicated; note A206 (labeled in red) coordinates Na+ at Sext via its backbone carbonyl oxygen. (d) Extracellular solvent accessibility of the ion binding sites in the structures at high and low [Na+]. Putative solvent channels are represented as light-purple surfaces. Divalent cation binding and apo structure of NCX_Mj. (a) A single Sr2+ (dark blue sphere) binds at SCa in crystals titrated with 10 mM Sr2+ and 2.5 mM Na+ (see also Supplementary Fig. 2). Residues involved in Sr2+ coordination are labeled. There are no significant changes in the side-chains involved in ion coordination, relative to the Na+-bound state. T50 and T209 (labeled in red) coordinate Sr2+ through their backbone carbonyl-oxygen atoms. High Na+ concentration (100 mM) completely displaces Sr2+ and reverts NCX_Mj to the occluded state (right panel). The contour level of the Fo – Fc omit map of the structure at high Na+ concentration was lowered (to 4σ) so as to visualize the density from Na+ ions and H2O. (b) Ca2+ (tanned spheres) binds either to SCa or Smid in crystals titrated with 10 mM Ca2+ and 2.5 mM Na+ (see also Supplementary Fig. 2). The relative occupancies are 55% and 45%, respectively. (c) Superimposition of NCX_Mj structures obtained at low Na+ concentration (10 mM) and pH 6.5 (brown) and in the absence of Na+ and pH 4 (light green), referred to as apo state. (d) Close-up view of the ion-binding sites in the apo (or high H+) state. The side chains of E54 and E213 from the low Na+ structure are also shown (light brown) for comparison. White spheres indicate the location Sint, Smid SCa. (e) Extracellular solvent accessibility of the ion-binding sites in apo NCX_Mj. Spontaneous changes in the structure of outward-occluded, fully Na+-occupied NCX_Mj (PDB code 3V5U) upon sequential displacement of Na+. (a) Representative simulation snapshots of NCX_Mj (Methods) with Na+ bound at Sext, SCa and Sint (orange cartoons, green spheres) and with Na+ bound only at SCa and Sint (marine cartoons, yellow spheres) (b) Close-up of the backbone of the N-terminal half of TM7 (TM7ab), in the same Na+ occupancy states depicted in (a). Black lines indicate (i, i + 4) carbonyl-amide pairs along the helix; specific O-N distances are indicated, in Å (magenta). (c) Representative simulation snapshots (same as above) with Na+ bound at SCa and Sint (marine cartoons, yellow spheres) and without any Na+ bound (grey cartoons). (d) Close-up of the ion-binding region in the fully Na+-occupied state. Approximate distances between TM2, TM3 and TM7 are indicated in Å. (e) Close-up of the ion-binding region in the partially Na+-occupied state. (f) Close-up of the ion-binding region in the Na+-free state. (g-i) Analytical descriptors of the changes just described, calculated from the simulations of each Na+-occupancy state depicted in panels (a-f). These descriptors were employed as collective variables in the Bias-Exchange Metadynamics simulations (Methods). (g) Probability distributions of an analytical descriptor of the backbone hydrogen-bonding pattern in TM7ab (Eq. 2). (h) Mean value (with standard deviation) of a quantitative descriptor of the solvent accessibility of the Sext site (Eq. 1). (i) Mean value (with standard deviation) of a quantitative descriptor of the solvent accessibility of the SCa site (Eq. 1). Thermodynamic basis for the proposed mechanism of substrate control of the alternating-access transition of NCX. (a) Calculated conformational free-energy landscapes for outward-facing NCX_Mj, for two different Na+-occupancy states, and for a state with no ions bound. The free energy is plotted as a function of two coordinates, each describing the degree of opening of the aqueous channels leading to the Sext and SCa sites, respectively (see Methods). Contours correspond to 1 kcal/mol intervals. Black circles map the X-ray structures of NCX_Mj obtained at high and low Na+ concentration, as well as that at low pH, reported in this study. (b) Density isosurfaces for water molecules within 12 Å of the ion-binding region (grey volumes), for each of the major conformational free-energy minima in each ion-occupancy state. Na+ ions are shown as green spheres. The two inverted-topology repeats in the transporter structure (transparent cartoons) are colored differently (TM1-5, orange; TM6-10, marine). (c) Close-up views of the ion-binding region in the same conformational free-energy minima. Key residues involved in Na+ and water coordination (W) are highlighted (sticks, black lines). The water-density maps in (b) is shown here as a grey mesh. Note D240 is protonated, while E54 and E213 are ionized. Thermodynamic basis for the proposed mechanism of substrate control of the alternating-access transition of NCX. (a) Calculated free-energy landscapes for outward-facing NCX_Mj, for the Ca2+ and the fully protonated state. The free energy is plotted as in Fig. 5. For Ca2+, a map is shown in which a correction for the charge-transfer between the ion and the protein is introduced, alongside the uncorrected map (see Supplementary Notes 3-4 and Supplementary Fig. 5-6). The uncorrected map overstabilizes the open state relative to the semi-open and occluded because it also overestimates the cost of dehydration of the ion, once it is bound to the protein (this effect is negligible for Na+). Black circles map the crystal structures obtained at high Ca2+ concentration and at low pH (or high H+) reported in this study. (b) Water-density isosurfaces analogous to those in Fig. 5 are shown for each of the major conformational free-energy minima in the free-energy maps. The Ca2+ ion is shown as a red sphere; the protein is shown as in Fig. 5. (c) Close-up views of the ion-binding region in the same conformational free-energy minima. Key residues involved in Ca2+ and water coordination (W) are highlighted (sticks, black lines). The water-density maps in (b) are shown here as a grey mesh. In the occluded state with Ca2+ bound, helix TM7ab bends in the same way as in the fully occupied Na+ state, as the carbonyl of Ala206 forms a hydrogen-bonding interaction with Ser210. Structural mechanism of extracellular forward ion exchange in NCX. The carbonyl groups of Ala47 (on TM2b) and Ala206 (on TM7b), and the side chains of Glu54 (on TM2c) and Glu213 (on TM7c) are highlighted; these are four of the key residues for ion chelation and conformational changes. The green open cylinders represent the gating helices TM1 and TM6. Asterisks mark the states whose crystal structures have been determined in this study. These states and their connectivity can also be deduced from the calculated free-energy landscapes, which also reveal a Ca2+-loaded outward-facing occluded state, and an unloaded, fully open state. Concentration-dependent Na+ occupancy of outward-facing NCX_Mj Sites \\ [Na+]\t2.5 mM\t10 mM\t20 mM\t100 mM\t150 mM\t \tSint\t0.84\t0.92\t0.99\t0.96\t1.00\t \tSCa\t0.75\t0.87\t0.97\t1.00\t1.00\t \tSext\t-\t-\t0.59\t0.79\t0.93\t \t\t \tTotal\t1.58\t1.79\t2.55\t2.75\t2.93\t \t\t \tConformation\tPartially open\tPartially open\tMixed\tOccluded\tOccluded\t \t Data collection and refinement statistics for the NCX_Mj structures obtained from crystals soaked with varying amounts of Na+, and no Ca2+, and at low pH and no Na+ or Ca2+. [Na+]\t2.5 mM PDB 5HWX\t10 mM PDB 5HWY\t20 mM PDB 5HXC\t100 mM PDB 5HXE\t150 mM PDB 5HYA\t0 mM PDB 5HXH\t \tData collection\t\t\t\t\t\t\t \tSpace group\tP212121\tC2\t \tCell dimensions\t\t\t\t\t\t\t \ta, b, c (Å)\t49.70, 72.28, 95.78\t46.21, 71.97, 95.63\t49.75, 72.56, 95.78\t49.77, 72.85, 96.36\t49.49, 72.88, 96.21\t164.18, 46.83, 96.96\t \tα, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 106.20, 90\t \tResolution (Å)\t2.40 (2.44-2.40)\t2.10 (2.14-2.10)\t2.10 (2.14-2.10)\t2.28 (2.32-2.28)\t1.90 (1.93-1.90)\t2.80 (2.85-2.80)\t \tRsym (%)\t9.4 (99.4)\t9.3 (69.8)\t10.5 (99.6)\t9.9 (56.3)\t8.6 (88.0)\t10.5 (94.3)\t \tI/σI\t24.2 (1.8)\t24.3 (2.7)\t20.6 (1.9)\t15.9 (2.8)\t32.9 (2.4)\t17.7 (1.2)\t \tCC1/2\t(0.625)\t(0.819)\t(0.602)\t(0.722)\t(0.786)\t(0.561)\t \tCompleteness (%)\t99.8 (99.1)\t99.9 (100)\t99.9 (99.8)\t96.4 (98.6)\t98.9 (97.8)\t99.9 (99.9)\t \tRedundancy\t6.9 (6.7)\t6.1 (6.0)\t7.1 (7.0)\t3.4 (3.3)\t9.3 (8.9)\t7.1 (5.7)\t \tRefinement\t\t\t\t\t\t\t \tResolution (Å)\t50-2.4\t50-2.1\t50-2.1\t50-2.3\t50-1.9\t50-2.80\t \tNo. reflections\t13977\t19254\t20739\t15767\t27923\t21489\t \tRwork/Rfree\t0.21/0.25\t0.19/0.22\t0.19/0.23\t0.19/0.24\t0.179/0.207\t0.20/0.26\t \tNo. atoms\t\t\t\t\t\t\t \tProtein\t2206\t2274\t2366\t2229\t2229\t4410\t \tLigand/Ion\t56/3\t154/2\t161/5\t162/6\t257/4\t121/2\t \tWater\t18\t36\t67\t67\t100\t39\t \tB-factors\t\t\t\t\t\t\t \tProtein\t53.25\t34.28\t34.91\t39.75\t26.05\t42.98\t \tLigand/Ion\t62.94/47.70\t55.86/31.75\t55.25/36.47\t58.24/38.93\t46.11/22.13\t54.29/63.47\t \tWater\t58.12\t41.16\t46.97\t49.74\t37.80\t33.17\t \tR.m.s deviations\t\t\t\t\t\t\t \tBond lengths (Å)\t0.004\t0.006\t0.008\t0.003\t0.006\t0.003\t \tBond angles (°)\t0.819\t0.915\t1.269\t1.024\t0.966\t0.705\t \t Values in parenthesis are for highest resolution shell. 5% of the data was used in the Rfree calculation. ‘Ligand’ atoms are from lipids, PEG400 and acetates. Data collection and refinement statistics for the NCX_Mj structures obtained from crystals soaked with varying amounts of Na+ and Sr2+. [Sr2+] / [Na+]\t10 mM / 2.5 mM PDB 5HXS\t1 mM / 2.5 mM N/A*\t0.1 mM / 2.5 mM N/A*\t10 mM / 10 mM N/A*\t10 mM / 100 mM N/A#\t \tData collection\t\t\t\t\t\t \tSpace group\tP212121\t \tCell dimensions\t\t\t\t\t\t \ta, b, c (Å)\t49.52, 72.35, 96.00\t49.76, 72.62, 95.62\t49.80, 72.27, 94.84\t49.67, 72.46, 96.43\t49.88, 72.43, 95.91\t \tα, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t \tResolution (Å)\t2.80 (2.85-2.80)\t2.90 (2.95-2.90)\t2.54 (2.58-2.54)\t2.30 (2.34-2.30)\t2.50 (2.54-2.50)\t \tRsym (%)\t12.3 (88.0)\t12.3 (91.0)\t11.9 (88.5)\t13.7 (95.8)\t10.9 (96.8)\t \tI/σI\t19.0 (1.8)\t19.2 (1.6)\t21.5 (1.9)\t18.4 (1.6)\t19.7 (1.8)\t \tCC1/2\t(0.674)\t(0.686)\t(0.768)\t(0.420)\t(0.767)\t \tCompleteness (%)\t98.8 (90.4)\t94.9 (96.1)\t99.9 (100.0)\t99.8 (99.9)\t99.6 (99.7)\t \tRedundancy\t6.5 (5.4)\t6.7 (6.9)\t7.1 (6.8)\t6.2 (4.5)\t5.9 (6.0)\t \tRefinement\t\t \tResolution (Å)\t50-2.80\t50-2.90\t50-2.54\t50-2.3\t50-2.5\t \tNo. reflections\t8927\t7611\t11483\t16488\t12665\t \tRwork/Rfree\t0.22/0.27\t0.22/0.27\t0.23/0.26\t0.20/0.25\t0.21/0.24\t \tNo. atoms\t\t\t\t\t\t \tProtein\t2227\t2249\t2217\t2271\t2223\t \tLigand/Ion\t110/2\t133/3\t155/3\t101/3\t104/6\t \tWater\t10\t2\t5\t34\t17\t \tB-factors\t\t\t\t\t\t \tProtein\t61.55\t67.82\t60.83\t46.42\t56.75\t \tLigand/Ion\t74.56/63.32\t85.56/70.47\t80.33/90.41\t64.79/38.25\t71.27/53.50\t \tWater\t59.22\t61.31\t68.58\t51.39\t55.07\t \tR.m.s deviations\t\t\t\t\t\t \tBond lengths (Å)\t0.004\t0.005\t0.005\t0.007\t0.003\t \tBond angles (°)\t0.887\t1.076\t0.963\t1.057\t0.733\t \t Values in parenthesis are for highest resolution shell. 5% of the data was used in the Rfree calculation. ‘Ligand’ atoms are from lipids, PEG400 and acetates. These structures are virtually identical to that resolved with 10 mM Sr2+ (PDB 5HXS), except for the weakened electron-density signal for the divalent ion, and were therefore not deposited in the PDB. This structure is virtually identical to that resolved with 100 mM Na+ without Ca2+ or Sr2+ (Table 1, PDB 5HXE) and was therefore not deposited in the PDB. Data collection and refinement statistics for the NCX_Mj structures obtained from crystals soaked with varying amounts of Na+ and Ca2+. [Ca2+] / [Na+]\t10 mM / 2.5 mM PDB 5HXR\t1 mM / 2.5 mM N/A*\t0.1 mM / 2.5 mM N/A*\t10 mM / 10 mM N/A*\t \tData collection\t\t\t\t\t \tSpace group\tP212121\t \tCell dimensions\t\t\t\t\t \ta, b, c (Å)\t49.70, 72.52, 96.94\t49.80, 72.26, 95.80\t49.48, 72.47, 96.30\t49.88, 72.22, 96.10\t \tα, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t \tResolution (Å)\t2.45 (2.49-2.45)\t2.65 (2.70-2.65)\t2.40 (2.44-2.40)\t2.20 (2.24-2.20)\t \tRsym (%)\t11.8 (94.2)\t11.1 (92.1)\t11.2 (91.3)\t10.3 (99.1)\t \tI/σI\t24.4 (1.6)\t20.9 (1.6)\t20.4 (1.6)\t22.4 (1.8)\t \tCC1/2\t(0.611)\t(0.696)\t(0.632)\t(0.549)\t \tCompleteness (%)\t99.7 (100.0)\t99.9 (100.0)\t99.2 (100.0)\t99.7 (100.0)\t \tRedundancy\t7.8 (7.8)\t7.0 (6.5)\t6.9 (7.0)\t7.1 (7.1)\t \tRefinement\t\t\t\t\t \tResolution (Å)\t50-2.45\t50-2.65\t50-2.40\t50-2.2\t \tNo. reflections\t12996\t10548\t13736\t18080\t \tRwork/Rfree\t0.22/0.26\t0.22/0.28\t0.20/0.26\t0.19/0.24\t \tNo. atoms\t\t\t\t\t \tProtein\t2211\t2225\t2228\t2284\t \tLigand/Ion\t114/3\t130/3\t162/3\t164/4\t \tWater\t16\t20\t42\t57\t \tB-factors\t\t\t\t\t \tProtein\t64.97\t71.22\t53.50\t45.16\t \tLigand/Ion\t78.98/60.53\t90.03/72.90\t70.67/45.53\t69.09/42.92\t \tWater\t68.57\t81.02\t64.12\t55.45\t \tR.m.s deviations\t\t\t\t\t \tBond lengths (Å)\t0.002\t0.004\t0.002\t0.004\t \tBond angles (°)\t0.630\t1.143\t0.686\t0.650\t \t Values in parenthesis are for highest resolution shell. 5% of the data was used in the Rfree calculation. ‘Ligand’ atoms are from lipids, PEG400 and acetates. These structures are virtually identical to that resolved with 10 mM Ca2+ and 2.5 mM Na+ (PDB 5HXR), except for the weakened electron-density signal for the divalent ion, and were therefore not deposited in the 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+[{"sourceid":"4919469","sourcedb":"","project":"","target":"","text":"Investigation of the Interaction between Cdc42 and Its Effector TOCA1 Transducer of Cdc42-dependent actin assembly protein 1 (TOCA1) is an effector of the Rho family small G protein Cdc42. It contains a membrane-deforming F-BAR domain as well as a Src homology 3 (SH3) domain and a G protein-binding homology region 1 (HR1) domain. TOCA1 binding to Cdc42 leads to actin rearrangements, which are thought to be involved in processes such as endocytosis, filopodia formation, and cell migration. We have solved the structure of the HR1 domain of TOCA1, providing the first structural data for this protein. We have found that the TOCA1 HR1, like the closely related CIP4 HR1, has interesting structural features that are not observed in other HR1 domains. We have also investigated the binding of the TOCA HR1 domain to Cdc42 and the potential ternary complex between Cdc42 and the G protein-binding regions of TOCA1 and a member of the Wiskott-Aldrich syndrome protein family, N-WASP. TOCA1 binds Cdc42 with micromolar affinity, in contrast to the nanomolar affinity of the N-WASP G protein-binding region for Cdc42. NMR experiments show that the Cdc42-binding domain from N-WASP is able to displace TOCA1 HR1 from Cdc42, whereas the N-WASP domain but not the TOCA1 HR1 domain inhibits actin polymerization. This suggests that TOCA1 binding to Cdc42 is an early step in the Cdc42-dependent pathways that govern actin dynamics, and the differential binding affinities of the effectors facilitate a handover from TOCA1 to N-WASP, which can then drive recruitment of the actin-modifying machinery. Introduction The Ras superfamily of small GTPases comprises over 150 members that regulate a multitude of cellular processes in eukaryotes. The superfamily can be divided into five families based on structural and functional similarities: Ras, Rho, Rab, Arf, and Ran. All members share a well defined core structure of ∼20 kDa known as the G domain, which is responsible for guanine nucleotide binding. It is this guanine nucleotide binding that underlies their function as molecular switches, controlling a vast array of signaling pathways. These molecular switches cycle between active, GTP-bound, and inactive, GDP-bound, states with the help of auxiliary proteins. The guanine nucleotide exchange factors mediate formation of the active state by promoting the dissociation of GDP, allowing GTP to bind. The GTPase-activating proteins stimulate the rate of intrinsic GTP hydrolysis, mediating the return to the inactive state (reviewed in Ref.). The overall conformation of small G proteins in the active and inactive states is similar, but they differ significantly in two main regions known as switch I and switch II. These regions are responsible for “sensing” the nucleotide state, with the GTP-bound state showing greater rigidity and the GDP-bound state adopting a more relaxed conformation (reviewed in Ref.). In the active state, G proteins bind to an array of downstream effectors, through which they exert their extensive roles within the cell. The structures of more than 60 small G protein·effector complexes have been solved, and, not surprisingly, the switch regions have been implicated in a large proportion of the G protein-effector interactions (reviewed in Ref.). However, because each of the 150 members of the superfamily interacts with multiple effectors, there are still a huge number of known G protein-effector interactions that have not yet been studied structurally. The Rho family comprises 20 members, of which three, RhoA, Rac1, and Cdc42, have been relatively well studied. The role of these three proteins in the coordination of the actin cytoskeleton has been examined extensively. RhoA acts to rearrange existing actin structures to form stress fibers, whereas Rac1 and Cdc42 promote de novo actin polymerization to form lamellipodia and filopodia, respectively. A number of RhoA and Rac1 effector proteins, including the formins and members of the protein kinase C-related kinase (PRK)6 family, along with Cdc42 effectors, including the Wiskott-Aldrich syndrome (WASP) family and the transducer of Cdc42-dependent actin assembly (TOCA) family, have also been linked to the pathways that govern cytoskeletal dynamics. Cdc42 effectors, TOCA1 and the ubiquitously expressed member of the WASP family, N-WASP, have been implicated in the regulation of actin polymerization downstream of Cdc42 and phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2). N-WASP exists in an autoinhibited conformation, which is released upon PI(4,5)P2 and Cdc42 binding or by other factors, such as phosphorylation. Following their release, the C-terminal regions of N-WASP are free to interact with G-actin and a known nucleator of actin assembly, the Arp2/3 complex. The importance of TOCA1 in actin polymerization has been demonstrated in a range of in vitro and in vivo studies, but the exact role of TOCA1 in the many pathways involving actin assembly remains unclear. The most widely studied role of TOCA1 is in membrane invagination and endocytosis, although it has also been implicated in filopodia formation, neurite elongation, transcriptional reprogramming via nuclear actin, and interaction with ZO-1 at tight junctions. A role in cell motility and invasion has also been established. TOCA1 comprises an N-terminal F-BAR domain, a central homology region 1 (HR1) domain, and a C-terminal SH3 domain. The F-BAR domain is a known dimerization, membrane-binding, and membrane-deforming module found in a number of cell signaling proteins. The TOCA1 SH3 domain has many known binding partners, including N-WASP and dynamin. The HR1 domain has been directly implicated in the interaction between TOCA1 and Cdc42, representing the first Cdc42-HR1 domain interaction to be identified. Other HR1 domains studied so far, including those from the PRK family, have been found to bind their cognate Rho family G protein-binding partner with high specificity and affinities in the nanomolar range. The structures of the PRK1 HR1a domain in complex with RhoA and the HR1b domain in complex with Rac1 show that the HR1 domain comprises an anti-parallel coiled-coil that interacts with its G protein binding partner via both helices. Both of the G protein switch regions are involved in the interaction. The coiled-coil fold is shared by the HR1 domain of the TOCA family protein, CIP4, and, based on sequence homology, by TOCA1 itself. These HR1 domains, however, show specificity for Cdc42, rather than RhoA or Rac1. How different HR1 domain proteins distinguish their specific G protein partners remains only partially understood, and structural characterization of a novel G protein-HR1 domain interaction would add to the growing body of information pertaining to these protein complexes. Furthermore, the biological function of the interaction between TOCA1 and Cdc42 remains poorly understood, and so far there has been no biophysical or structural insight. The interactions of TOCA1 and N-WASP with Cdc42 as well as with each other have raised questions as to whether the two Cdc42 effectors can interact with a single molecule of Cdc42 simultaneously. There is some evidence for a ternary complex between Cdc42, N-WASP, and TOCA1, but there was no direct demonstration of simultaneous contacts between the two effectors and a single molecule of Cdc42. Nonetheless, the substantial difference between the structures of the G protein-binding regions of the two effectors is intriguing and implies that they bind to Cdc42 quite differently, providing motivation for investigating the possibility that Cdc42 can bind both effectors concurrently. WASP interacts with Cdc42 via a conserved, unstructured binding motif known as the Cdc42- and Rac-interactive binding region (CRIB), which forms an intermolecular β-sheet, expanding the anti-parallel β2 and β3 strands of Cdc42. In contrast, the TOCA family proteins are thought to interact via the HR1 domain, which may form a triple coiled-coil with switch II of Rac1, like the HR1b domain of PRK1. Here, we present the solution NMR structure of the HR1 domain of TOCA1, providing the first structural data for this protein. We also present data pertaining to binding of the TOCA HR1 domain to Cdc42, which is the first biophysical description of an HR1 domain binding this particular Rho family small G protein. Finally, we investigate the potential ternary complex between Cdc42 and the G protein-binding regions of TOCA1 and N-WASP, contributing to our understanding of G protein-effector interactions as well as the roles of Cdc42, N-WASP, and TOCA1 in the pathways that govern actin dynamics. Experimental Procedures Expression Constructs The Xenopus tropicalis TOCA1 HR1 domain (residues 330–426 and N-terminally extended constructs as indicated) were amplified from cDNA (TOCA1 accession number NM_001005148) and cloned into pGEX-6P-1 (GE Healthcare) or pGEX-HisP. The HR1 domain of human CIP4 (residues 388–481) was amplified from IMAGE clone 3532036, the Xenopus laevis FBP17 HR1 domain (residues 385–486) from IMAGE clone 5514481, and the X. tropicalis N-WASP G protein-binding domain (GBD) (residues 197–255) from IMAGE clone 5379332, and all were cloned into pGEX-6P-1. The resulting constructs express the proteins as N-terminal GST fusions with a 3C protease-cleavable tag, with pGEX-HisP expressing an additional C-terminal His6 tag. Human Cdc42Δ7Q61L and full-length Cdc42 were cloned into pGEX-2T (GE Healthcare) and pGEX-6P-1, respectively. A C-terminally extended construct of TOCA1 comprising residues 330–545 was cloned into pMAT10-P.7 The resulting construct expresses TOCA1 330–545 as an N-terminal His-MBP fusion protein with a 3C protease-cleavable tag. Full-length X. tropicalis TOCA1, TOCA1 F-BAR (residues 1–287), and TOCA1 ΔSH3 (residues 1–480) were PCR-amplified from a cDNA clone (IMAGE 5157175) and cloned into pET-His6-SNAP using FseI and AscI sites that had been incorporated into the primers to create His-SNAP-TOCA1 proteins. Protein Expression GST fusion proteins (HR1 domains and Cdc42) were expressed in E. coli BL21 cells (Invitrogen). Stationary cultures were diluted 1:10 and grown at 37 °C until an A600 of ∼0.8 was reached and then induced with 0.1 mm isopropyl-β-d-thiogalactopyranoside for 20 h at 20 °C. The GST-N-WASP GBD construct was expressed in E. coli BL21-CodonPlus®-RIL (Agilent Technologies). The proteins were purified using glutathione-agarose beads (Sigma) and eluted from the beads by cleavage of the GST tag with 3C protease (HR1 domains, N-WASP GBD, and full-length Cdc42Q61L) or thrombin (Novagen, Cdc42Δ7Q61L) prior to gel filtration on a 16/60 S75 column (GE Healthcare). His-MBP-HR1-SH3 was purified using nickel-nitrilotriacetic acid-agarose beads (Life Technologies) prior to cleavage with 3C protease and gel filtration. Full-length TOCA1, TOCA1 F-BAR, and TOCA1 ΔSH3 were expressed from pET-His6-SNAP in BL21 pLysS, grown at 37 °C until an A600 of ∼0.6 was reached, and induced with 0.3 mm isopropyl-β-d-thiogalactopyranoside overnight at 19 °C. Proteins were coupled to nickel-nitrilotriacetic acid-agarose (Qiagen), eluted using increasing concentrations of imidazole, and further purified by gel filtration using a 16/60 S200 column (GE Healthcare). All protein concentrations were determined by amino acid analysis (Protein and Nucleic Acid Chemistry Facility, Department of Biochemistry, University of Cambridge). Nucleotide Exchange For NMR experiments, Cdc42 was nucleotide-exchanged for the non-hydrolyzable GTP analogue GMPPNP (Sigma) as described previously. For scintillation proximity assays (SPAs), Cdc42 was loaded with [3H]GTP using [8-3H]GTP (PerkinElmer Life Sciences), as described previously. The protein was confirmed as full-length using mass spectrometry (PNAC facility, Department of Biochemistry, University of Cambridge). SPAs For direct assays, GST-PAK, GST-ACK, or His-tagged TOCA1 constructs were attached to a fluoromicrosphere via an anti-GST or anti-His antibody in the presence of Cdc42Δ7Q61L·[3H]GTP. Binding curves were fitted using a direct binding isotherm to obtain Kd values and their curve-fitting errors for the G protein-effector interactions. For competition assays, free ACK GBD, TOCA1 HR1, TOCA1 HR1SH3, or N-WASP GBD was titrated into a mixture of 30 nm Cdc42Δ7Q61L·[3H]GTP and 30 nm GST-ACK immobilized on a fluoromicrosphere as above. Data were fitted to competition binding isotherms to obtain Kd values and curve-fitting errors, as described previously. NMR Spectroscopy The NMR experiments and resonance assignments of the HR1 domain are described. The NMR experiments were carried out with 0.9 mm 13C/15N-labeled HR1 domain in 20 mm sodium phosphate, pH 7.5, 150 mm NaCl, 5 mm MgCl2, 5 mm DTT, 10% D2O. Distance restraints were derived from a 15N-separated NOESY (100-ms mixing time) recorded on a Bruker DRX500 and a 13C-separated NOESY (100-ms mixing time) recorded on an Avance AV600. NMR data were processed using AZARA (W. Boucher, University of Cambridge) and analyzed using ANALYSIS. Structure Calculation Structures were calculated iteratively using CNS version 1.0 interfaced to Aria version 2.3.1. The PROSLQ force field was used for non-bonded parameters. Backbone torsion angles were estimated from CA, CO, CB, N, and HA chemical shifts using TALOS-N. The “strong” φ and ψ restraints were included with an error of ±2 S.D. values of the averaged TALOS-N predictions. Dihedral angle predictions for residues 323–340 were weak, so no restraints were included for this region. NMR Titrations All of the 15N and 13C HSQCs were recorded at 25 °C in 50 mm sodium phosphate, pH 5.5, 25 mm NaCl, 5 mm MgCl2, 5 mm DTT, 10% D2O on a Bruker DRX500. 15N-HR1 HSQC experiments were recorded on 0.2 mm 15N-HR1 domain with HR1/Cdc42·GMPPNP ratios of 1:0, 1:0.25, 1:0.5, 1:1, and 1:4. Experiments were recorded on 0.27 mm 15N-Cdc42·GMPPNP at Cdc42/HR1 ratios of 1:0, 1:0.25, 1:0.5, and 1:2.2. The 15N HSQC titrations with N-WASP were recorded on 0.6 mm 15N-HR1 domain or 0.15 mm 15N-Cdc42 at the ratios indicated in the figures. Chemical Shift Mapping The chemical shift changes, δ, were calculated using the equation,  where δ(1H) and δ(15N) are the chemical shift changes for the 1H and 15N dimensions, respectively. Residues that had disappeared were assigned a δ value larger than the maximum calculated δ for the data set, and residues that were too overlapped to be reliably assigned in the complex spectra were assigned δ = 0. The residues that had shifted more than the mean chemical shift change across the spectrum were classed as significant and were filtered for solvent accessibility using NACCESS. Residues with \u003c50% solvent accessibility were considered to be buried and unavailable for binding. Pyrene Actin Assays Pyrene actin assays were carried out as described previously. Xenopus high speed supernatant was used at 5 mg/ml and supplemented with 0.12 mg/ml pyrene actin as described previously. TOCA1 HR1 domain or N-WASP CRIB domain was added at the concentrations indicated. Liposomes were made, using methods described previously, from 60% phosphatidylcholine, 30% phosphatidylserine, and 10% PI(4,5)P2 to 2 mm final lipid concentration. All of the lipids used were natural brain or liver lipids from Avanti Polar Lipids. The assays were initiated by the addition of 5 μl of liposomes per 200 μl of reaction mix. Results Cdc42-TOCA1 Binding TOCA1 was identified in Xenopus extracts as a protein necessary for Cdc42-dependent actin assembly and was shown to bind to Cdc42·GTPγS but not to Cdc42·GDP or to Rac1 and RhoA. Given its homology to other Rho family binding modules, it is likely that the HR1 domain of TOCA1 is sufficient to bind Cdc42. The C. elegans TOCA1 orthologues also bind to Cdc42 via their consensus HR1 domain. The HR1 domains from the PRK family bind their G protein partners with a high affinity, exhibiting a range of submicromolar dissociation constants (Kd) as low as 26 nm. A Kd in the nanomolar range was therefore expected for the interaction of the TOCA1 HR1 domain with Cdc42. We generated an X. tropicalis TOCA1 HR1 domain construct encompassing residues 330–426. This region comprises the complete HR1 domain based on secondary structure predictions and sequence alignments with another TOCA family member, CIP4, whose structure has been determined. The interaction between [3H]GTP·Cdc42 and a C-terminally His-tagged TOCA1 HR1 domain construct was investigated using SPA. The binding isotherm for the interaction is shown in Fig. 1A, together with the Cdc42-PAK interaction as a positive control. The binding of TOCA1 HR1 to Cdc42 was unexpectedly weak, with a Kd of \u003e1 μm. It was not possible to estimate the Kd more accurately using direct SPA experiments, because saturation could not be reached due to nonspecific signal at higher protein concentrations. The TOCA1 HR1-Cdc42 interaction is low affinity.\nA, curves derived from direct binding assays in which the indicated concentrations of Cdc42Δ7Q61L·[3H]GTP were incubated with 30 nm GST-PAK or HR1-His6 in SPAs. The SPA signal was corrected by subtraction of control data with no GST-PAK or HR1-His6. The data were fitted to a binding isotherm to give an apparent Kd and are expressed as a percentage of the maximum signal; B and C, competition SPA experiments were carried out with the indicated concentrations of ACK GBD (B) or HR1 domain (C) titrated into 30 nm GST-ACK and either 30 nm Cdc42Δ7Q61L·[3H]GTP or full-length Cdc42Q61L·[3H]GTP. The Kd values derived for the ACK GBD with Cdc42Δ7 and full-length Cdc42 were 0.032 ± 0.01 and 0.011 ± 0.01 μm, respectively. The Kd values derived for the TOCA1 HR1 with Cdc42Δ7 and full-length Cdc42 were 6.05 ± 1.96 and 5.39 ± 1.69 μm, respectively. It was possible that the low affinity observed was due to negative effects of immobilization of the HR1 domain, so other methods were employed, which utilized untagged proteins. Isothermal titration calorimetry was carried out, but no heat changes were observed at a range of concentrations and temperatures (data not shown), suggesting that the interaction is predominantly entropically driven. Other G protein-HR1 domain interactions have also failed to show heat changes in our hands.7 Infrared interferometry with immobilized Cdc42 was also attempted but was unsuccessful for both TOCA1 HR1 and for the positive control, ACK. The affinity was therefore determined using competition SPAs. A complex of a GST fusion of the GBD of ACK, which binds with a high affinity to Cdc42, with radiolabeled [3H]GTP·Cdc42 was preformed, and the effect of increasing concentrations of untagged TOCA1 HR1 domain was examined. Competition of GST-ACK GBD bound to [3H]GTP·Cdc42 by free ACK GBD was used as a control and to establish the value of background counts when Cdc42 is fully displaced. The data were fitted to a binding isotherm describing competition. Free ACK competed with itself with an affinity of 32 nm, similar to the value obtained by direct binding of 23 nm. The TOCA1 HR1 domain also fully competed with the GST-ACK but bound with an affinity of 6 μm (Fig. 1, B and C), in agreement with the low affinity observed in the direct binding experiments. The Cdc42 construct used in the binding assays has seven residues deleted from the C terminus to facilitate purification. These residues are not generally required for G protein-effector interactions, including the interaction between RhoA and the PRK1 HR1a domain. In contrast, the C terminus of Rac1 contains a polybasic sequence, which is crucial for Rac1 binding to the HR1b domain from PRK1. As the observed affinity between TOCA1 HR1 and Cdc42 was much lower than expected, we reasoned that the C terminus of Cdc42 might be necessary for a high affinity interaction. The binding experiments were repeated with full-length [3H]GTP·Cdc42, but the affinity of the HR1 domain for full-length Cdc42 was similar to its affinity for truncated Cdc42 (Kd ≈ 5 μm; Fig. 1C). Thus, the C-terminal region of Cdc42 is not required for maximal binding of TOCA1 HR1. Another possible explanation for the low affinities observed was that the HR1 domain alone is not sufficient for maximal binding of the TOCA proteins to Cdc42 and that the other domains are required. Indeed, GST pull-downs performed with in vitro translated human TOCA1 fragments had suggested that residues N-terminal to the HR1 domain may be required to stabilize the HR1 domain structure. Furthermore, both BAR and SH3 domains have been implicated in interactions with small G proteins (e.g. the BAR domain of Arfaptin2 binds to Rac1 and Arl1), while an SH3 domain mediates the interaction between Rac1 and the guanine nucleotide exchange factor, β-PIX. TOCA1 dimerizes via its F-BAR domain, which could also affect Cdc42 binding, for example by presenting two HR1 domains for Cdc42 interactions. Various TOCA1 fragments (Fig. 2A) were therefore assessed for binding to full-length Cdc42 by direct SPA. The isolated F-BAR domain showed no binding to full-length Cdc42 (Fig. 2B). Full-length TOCA1 and ΔSH3 TOCA1 bound with micromolar affinity (Fig. 2B), in a similar manner to the isolated HR1 domain (Fig. 1A). The HR1-SH3 protein could not be purified to homogeneity as a fusion protein, so it was assayed in competition assays after cleavage of the His tag. This construct competed with GST-ACK GBD to give a similar affinity to the HR1 domain alone (Kd = 4.6 ± 4 μm; Fig. 2C). Taken together, these data suggest that the TOCA1 HR1 domain is sufficient for maximal binding and that this binding is of a relatively low affinity compared with many other Cdc42·effector complexes. The Cdc42-HR1 interaction is of low affinity in the context of full-length protein and in TOCA1 paralogues.\nA, diagram illustrating the TOCA1 constructs assayed for Cdc42 binding. Domain boundaries are derived from secondary structure predictions; B, binding curves derived from direct binding assays, in which the indicated concentrations of Cdc42Δ7Q61L·[3H]GTP were incubated with 30 nm GST-ACK or His-tagged TOCA1 constructs, as indicated, in SPAs. The SPA signal was corrected by subtraction of control data with no fusion protein. The data were fitted to a binding isotherm to give an apparent Kd and are expressed as a percentage of the maximum signal. C–E, representative examples of competition SPA experiments carried out with the indicated concentrations of the TOCA1 HR1-SH3 construct titrated into 30 nm GST-ACK and 30 nm Cdc42Δ7Q61L·[3H]GTP (C) or HR1CIP4 (D) or HR1FBP17 (E) titrated into 30 nm GST-ACK and 30 nm Cdc42FLQ61L·[3H]GTP. The low affinity of the TOCA1 HR1-Cdc42 interaction raised the question of whether the other known Cdc42-binding TOCA family proteins, FBP17 and CIP4, also bind weakly. The HR1 domains from FBP17 and CIP4 were purified and assayed for Cdc42 binding in competition SPAs, analogous to those carried out with the TOCA1 HR1 domain. The affinities of both the FBP17 and CIP4 HR1 domains were also in the low micromolar range (10 and 5 μm, respectively) (Fig. 2, D and E), suggesting that low affinity interactions with Cdc42 are a common feature within the TOCA family. Structure of the TOCA1 HR1 Domain Because the TOCA1 HR1 domain was sufficient for maximal Cdc42-binding, we used this construct for structural studies. Initial experiments were performed with TOCA1 residues 324–426, but we observed that the N terminus was cleaved during purification to yield a new N terminus at residue 330 (data not shown). We therefore engineered a construct comprising residues 330–426 to produce the minimal, stable HR1 domain. Backbone and side chain resonances were assigned as described. 2,778 non-degenerate NOE restraints were used in initial structure calculations (1,791 unambiguous and 987 ambiguous), derived from three-dimensional 15N-separated NOESY and 13C-separated NOESY experiments. There were 1,845 unambiguous NOEs and 757 ambiguous NOEs after eight iterations. 100 structures were calculated in the final iteration; the 50 lowest energy structures were water-refined; and of these, the 35 lowest energy structures were analyzed. Table 1 indicates that the HR1 domain structure is well defined by the NMR data. Experimental restraints and structural statistics a \u003cSA\u003e, the average root mean square deviations for the ensemble ± S.D. b \u003cSA\u003ec, values for the structure that is closest to the mean. The structure closest to the mean is shown in Fig. 3A. The two α-helices of the HR1 domain interact to form an anti-parallel coiled-coil with a slight left-handed twist, reminiscent of the HR1 domains of CIP4 (PDB code 2KE4) and PRK1 (PDB codes 1CXZ and 1URF). A sequence alignment illustrating the secondary structure elements of the TOCA1 and CIP4 HR1 domains and the HR1a and HR1b domains from PRK1 is shown in Fig. 3B. The structure of the TOCA1 HR1 domain.\nA, the backbone trace of the 35 lowest energy structures of the HR1 domain overlaid with the structure closest to the mean is shown alongside a schematic representation of the structure closest to the mean. Flexible regions at the N and C termini (residues 330–333 and 421–426) are omitted for clarity. B, a sequence alignment of the HR1 domains from TOCA1, CIP4, and PRK1. The secondary structure was deduced using Stride, based on the Ramachandran angles, and is indicated as follows: gray, turn; yellow, α-helix; blue, 310 helix; white, coil. C, a close-up of the N-terminal region of TOCA1 HR1, indicating some of the NOEs defining its position with respect to the two α-helices. Dotted lines, NOE restraints. D, a close-up of the interhelix loop region showing some of the contacts between the loop and helix 1. NOEs are indicated with dotted lines. All structural figures were generated using PyMOL. In the HR1a domain of PRK1, a region N-terminal to helix 1 forms a short α-helix, which packs against both helices of the HR1 domain. This region of TOCA1 HR1 (residues 334–340) is well defined in the family of structures (Fig. 3A) but does not form an α-helix. It instead forms a series of turns, defined by NOE restraints observed between residues separated by one (residues 332–334, 333–335, etc.) or two (residues 337–340) residues in the sequence and the φ and ψ angles, assessed using Stride. These turns cause the chain to reverse direction, allowing the N-terminal segment (residues 334–340) to contact both helices of the HR1 domain. Long range NOEs were observed linking Leu-334, Glu-335, and Asp-336 with Trp-413 of helix 2, Leu-334 with Lys-409 of helix 2, and Phe-337 and Ser-338 with Arg-345, Arg-348, and Leu-349 of helix 1. These contacts are summarized in Fig. 3C. The two α-helices of TOCA1 HR1 are separated by a long loop of 10 residues (residues 380–389) that contains two short 310 helices (residues 381–383 and 386–389). Interestingly, side chains of residues within the loop region point back toward helix 1; for example, there are numerous distinct NOEs between the side chains of Asn-380 and Met-383 of the loop region and Tyr-377 and Val-376 of helix 1 (Fig. 3D). The backbone NH and CHα groups of Gly-384 and Asp-385 also show NOEs with the side chain of Tyr-377. Mapping the TOCA1 and Cdc42 Binding Interfaces The HR1TOCA1-Cdc42 interface was investigated using NMR spectroscopy. A series of 15N HSQC experiments was recorded on 15N-labeled TOCA1 HR1 domain in the presence of increasing concentrations of unlabeled Cdc42Δ7Q61L·GMPPNP to map the Cdc42-binding surface. A comparison of the 15N HSQC spectra of free HR1 and HR1 in the presence of excess Cdc42 shows that although some peaks were shifted, several were much broader in the complex, and a considerable subset had disappeared (Fig. 4A). This behavior cannot be explained by the increase in molecular mass (from 12 to 33 kDa) when Cdc42 binds and is more likely to be due to conformational exchange. This leads to broadening of the peaks so that they are not detectable. Overall chemical shift perturbations (CSPs) were calculated for each residue, whereas those that had disappeared were assigned a shift change of 0.2 (Fig. 4B). A peak that disappeared or had a CSP above the mean CSP for the spectrum was considered to be significantly affected. Mapping the binding surface of Cdc42 onto the TOCA1 HR1 domain.\nA, the 15N HSQC of 200 μm TOCA1 HR1 domain is shown in the free form (black) and in the presence of a 4-fold molar excess of Cdc42Δ7Q61L·GMPPNP (red). Expansions of two regions are shown with peak assignments, showing backbone amides in fast or intermediate exchange. B, CSPs were calculated as described under “Experimental Procedures” and are shown for backbone and side chain NH groups. The mean CSP is marked with a red line. Residues that disappeared in the presence of Cdc42 were assigned a CSP of 0.2 but were excluded when calculating the mean CSP and are indicated with open bars. Those that were not traceable due to spectral overlap were assigned a CSP of zero and are marked with an asterisk below the bar. Residues with affected side chain CSPs derived from 13C HSQCs are marked with green asterisks above the bars. Secondary structure elements are shown below the graph. C, a schematic representation of the HR1 domain. Residues with significantly affected backbone or side chain chemical shifts when Cdc42 bound and that are buried are colored dark blue, whereas those that are solvent-accessible are colored yellow. Residues with significantly affected backbone and side chain groups that are solvent-accessible are colored red. A close-up of the binding region is shown, with affected side chain heavy atoms shown as sticks. D, the G protein-binding region is marked in red onto structures of the HR1 domains as indicated. 15N HSQC shift mapping experiments report on changes to amide groups, which are mainly inaccessible because they are buried inside the helices and are involved in hydrogen bonds. Therefore, 13C HSQC and methyl-selective SOFAST-HMQC experiments were also recorded on 15N,13C-labeled TOCA1 HR1 to yield more information on side chain involvement. The affected CH groups underwent significant line broadening, similarly to the NH peaks. Side chains whose CH groups disappeared in the presence of Cdc42 are marked on the graph in Fig. 4B with green asterisks. TOCA1 residues whose signals were affected by Cdc42 binding were mapped onto the structure of TOCA1 HR1 (Fig. 4C). The changes were localized to one end of the coiled-coil, and the binding site appeared to include residues from both α-helices and the loop region that joins them. Residues outside of this region were not significantly affected, indicating that there was no widespread conformational change. The residues in the interhelical loop and helix 1 that contact each other (Fig. 3D) show shift changes in their backbone NH and side chains in the presence of Cdc42. For example, the side chain of Asn-380 and the backbones of Val-376 and Tyr-377 were significantly affected but are all buried in the free TOCA1 HR1 structure, indicating that local conformational changes in the loop may facilitate complex formation. The chemical shift mapping data indicate that the G protein-binding region of the TOCA1 HR1 domain is broadly similar to that of the CIP4 and PRK1 HR1 domains (Figs. 3B and 4D). The corresponding 15N and 13C NMR experiments were also recorded on 15N-Cdc42Δ7Q61L·GMPPNP or 15N/13C -Cdc42Δ7Q61L·GMPPNP in the presence of unlabeled HR1 domain. The overall CSP was calculated for each residue. As was the case when labeled HR1 was observed, several peaks were shifted in the complex, but many disappeared, indicating exchange on an unfavorable, millisecond time scale (Fig. 5A). Detailed side chain data could not be obtained for all residues due to spectral overlap, but constant time 13C HSQC and methyl-selective SOFAST-HMQC experiments provided further information on certain well resolved side chains (marked with green asterisks in Fig. 5B). Mapping the binding surface of the HR1 domain onto Cdc42.\nA, the 15N HSQC of Cdc42Δ7Q61L·GMPPNP is shown in its free form (black) and in the presence of excess TOCA1 HR1 domain (1:2.2, red). Expansions of two regions are shown, with most peaks in fast or intermediate exchange. B, CSPs are shown for backbone NH groups. The red line indicates the mean CSP, plus one S.D. Residues that disappeared in the presence of Cdc42 were assigned a CSP of 0.1 and are indicated with open bars. Those that were not traceable due to overlap are marked with an asterisk. Residues with disappeared peaks in 13C HSQC experiments are marked on the chart with green asterisks. Secondary structure elements are indicated below the graph. C, the residues with significantly affected backbone and side chain groups are highlighted on an NMR structure of free Cdc42Δ7Q61L·GMPPNP; those that are buried are colored dark blue, whereas those that are solvent-accessible are colored red. Residues with either side chain or backbone groups affected are colored blue if buried and yellow if solvent-accessible. Residues without information from shift mapping are colored gray. The flexible switch regions are circled. As many of the peaks disappeared, the mean chemical shift change was relatively low, so a threshold of the mean plus one S.D. value was used to define a significant CSP. Residues that disappeared were also classed as significantly affected. Parts of the switch regions (Fig. 5, B and C) are invisible in NMR spectra recorded on free Cdc42 due to conformational exchange. These switch regions become visible in Cdc42 and other small G protein·effector complexes due to a decrease in conformational freedom upon complex formation. The switch regions of Cdc42 did not, however, become visible in the presence of the TOCA1 HR1 domain. Indeed, Ser-30 of switch I and Arg-66, Arg-68, Leu-70, and Ser-71 of switch II are visible in free Cdc42 but disappear in the presence of the HR1 domain. This suggests that the switch regions are not rigidified in the HR1 complex and are still in conformational exchange. Nevertheless, mapping of the affected residues onto the NMR structure of free Cdc42Δ7Q61L·GMPPNP (Fig. 5C)8 shows that, although they are relatively widespread compared with changes in the HR1 domain, in general, they are on the face of the protein that includes the switches. Although the binding interface may be overestimated, this suggests that the switch regions are involved in binding to TOCA1. Modeling the Cdc42·TOCA1 HR1 Complex The Cdc42·HR1TOCA1 complex was not amenable to full structural analysis due to the weak interaction and the extensive exchange broadening seen in the NMR experiments. HADDOCK was therefore used to perform rigid body docking based on the structures of free HR1 domain and Cdc42 and ambiguous interaction restraints derived from the titration experiments described above. Residues with significantly affected resonances and more than 50% solvent accessibility were defined as active. Passive residues were defined automatically as those neighboring active residues. The orientation of the HR1 domain with respect to Cdc42 cannot be definitively concluded in the absence of unambiguous distance restraints; hence, HADDOCK produced a set of models in which the HR1 domain contacts the same surface on Cdc42 but is in various orientations with respect to Cdc42. The cluster with the lowest root mean square deviation from the lowest energy structure is assumed to be the best model. By these criteria, in the best model, the HR1 domain is in a similar orientation to the HR1a domain of PRK1 bound to RhoA and the HR1b domain bound to Rac1. A representative model from this cluster is shown in Fig. 6A alongside the Rac1-HR1b structure (PDB code 2RMK) in Fig. 6B. Model of Cdc42·HR1 complex.\nA, a representative model of the Cdc42·HR1 complex from the cluster closest to the lowest energy model produced using HADDOCK. Residues of Cdc42 that are affected in the presence of the HR1 domain but are not in close proximity to it are colored in red and labeled. B, structure of Rac1 in complex with the HR1b domain of PRK1 (PDB code 2RMK). C, sequence alignment of RhoA, Cdc42 and Rac1. Contact residues of RhoA and Rac1 to PRK1 HR1a and HR1b, respectively, are colored cyan. Residues of Cdc42 that disappear or show chemical shift changes in the presence of TOCA1 are colored cyan if also identified as contacts in RhoA and Rac1 and yellow if they are not. Residues equivalent to Rac1 and RhoA contact sites but that are invisible in free Cdc42 are gray. D, regions of interest of the Cdc42·HR1 domain model. The four lowest energy structures in the chosen HADDOCK cluster are shown overlaid, with the residues of interest shown as sticks and labeled. Cdc42 is shown in cyan, and TOCA1 is shown in purple. A sequence alignment of RhoA, Cdc42, and Rac1 is shown in Fig. 6C. The RhoA and Rac1 contact residues in the switch regions are invisible in the spectra of Cdc42, but they are generally conserved between all three G proteins. Several Cdc42 residues identified by chemical shift mapping are not in close contact in the Cdc42·TOCA1 model (Fig. 6A). Some of these can be rationalized; for example, Thr-24Cdc42, Leu-160Cdc42, and Lys-163Cdc42 all pack behind switch I and are likely to be affected by conformational changes within the switch, while Glu-95Cdc42 and Lys-96Cdc42 are in the helix behind switch II. Other residues that are affected in the Cdc42·TOCA1 complex but that do not correspond to contact residues of RhoA or Rac1 (Fig. 6C) include Gln-2Cdc42, Lys-16Cdc42, Thr-52Cdc42, and Arg-68Cdc42. Lys-16Cdc42 is unlikely to be a contact residue because it is involved in nucleotide binding, but the others may represent specific Cdc42-TOCA1 contacts. In the model, these side chains are involved in direct contacts (Fig. 6D). Competition between N-WASP and TOCA1 From the known interactions and effects of the proteins in biological systems, it has been suggested that TOCA1 and N-WASP could bind Cdc42 simultaneously. Studies in CHO cells indicated that a Cdc42·N-WASP·TOCA1 complex existed because FRET was observed between RFP-TOCA1 and GFP-N-WASP, and the efficiency was decreased when an N-WASP mutant was used that no longer binds Cdc42. An overlay of the HADDOCK model of the Cdc42·HR1TOCA1 complex and the structure of Cdc42 in complex with the GBD of the N-WASP homologue, WASP (PDB code 1CEE), shows that the HR1 and GBD binding sites only partly overlap, and, therefore, a ternary complex remained possible (Fig. 7A). Interestingly, the presence of the TOCA1 HR1 would not prevent the core CRIB of WASP from binding to Cdc42, although the regions C-terminal to the CRIB that are required for high affinity binding of WASP would interfere sterically with the TOCA1 HR1. A basic region in WASP including three lysines (residues 230–232), N-terminal to the core CRIB, has been implicated in an electrostatic steering mechanism, and these residues would be free to bind in the presence of TOCA1 HR1 (Fig. 7A). The N-WASP GBD displaces the TOCA1 HR1 domain.\nA, the model of the Cdc42·TOCA1 HR1 domain complex overlaid with the Cdc42-WASP structure. Cdc42 is shown in green, and TOCA1 is shown in purple. The core CRIB region of WASP is shown in red, whereas its basic region is shown in orange and the C-terminal region required for maximal affinity is shown in cyan. A semitransparent surface representation of Cdc42 and WASP is shown overlaid with the schematic. B, competition SPA experiments carried out with indicated concentrations of the N-WASP GBD construct titrated into 30 nm GST-ACK or GST-WASP GBD and 30 nm Cdc42Δ7Q61L·[3H]GTP. C, Selected regions of the 15N HSQC of 145 μm Cdc42Δ7Q61L·GMPPNP with the indicated ratios of the TOCA1 HR1 domain, the N-WASP GBD, or both, showing that the TOCA HR1 domain does not displace the N-WASP GBD. D, selected regions of the 15N HSQC of 600 μm TOCA1 HR1 domain in complex with Cdc42 in the absence and presence of the N-WASP GBD, showing displacement of Cdc42 from the HR1 domain by N-WASP. An N-WASP GBD construct was produced, and its affinity for Cdc42 was measured by competition SPA (Fig. 7B). The Kd that was determined (37 nm) is consistent with the previously reported affinity. Unlabeled N-WASP GBD was titrated into 15N-Cdc42Δ7Q61L·GMPPNP, and the backbone NH groups were monitored using HSQCs (Fig. 7C). Unlabeled HR1TOCA1 was then added to the Cdc42·N-WASP complex, and no changes were seen, suggesting that the N-WASP GBD was not displaced even in the presence of a 5-fold excess of HR1TOCA1. These experiments were recorded at sufficiently high protein concentrations (145 μm Cdc42, 145 μm N-WASP GBD, 725 μm TOCA1 HR1 domain) to be far in excess of the Kd values of the individual interactions (TOCA1 Kd ≈ 5 μm, N-WASP Kd = 37 nm). A comparison of the HSQC experiments recorded on 15N-Cdc42 alone, in the presence of TOCA1 HR1, N-WASP GBD, or both, shows that the spectra in the presence of N-WASP and in the presence of both N-WASP and TOCA1 HR1 are identical (Fig. 7C). Furthermore, 15N-TOCA1 HR1 was monitored in the presence of unlabeled Cdc42Δ7Q61L·GMPPNP (1:1) before and after the addition of 0.25 and 1.0 eq of unlabeled N-WASP GBD. The spectrum when N-WASP and TOCA1 were equimolar was identical to that of the free HR1 domain, whereas the spectrum in the presence of 0.25 eq of N-WASP was intermediate between the TOCA1 HR1 free and complex spectra (Fig. 7D). When in fast exchange, the NMR signal represents a population-weighted average between free and bound states, so the intermediate spectrum indicates that the population comprises a mixture of free and bound HR1 domain. Hence, a third, intermediate state that includes all three proteins is unlikely. Again, the experiments were recorded on protein samples far in excess of the individual Kd values (600 μm each protein). These data indicate that the HR1 domain is displaced from Cdc42 by N-WASP and that a ternary complex comprising TOCA1 HR1, N-WASP GBD, and Cdc42 is not formed. Taken together, the data in Fig. 7, C and D, indicate unidirectional competition for Cdc42 binding in which the N-WASP GBD displaces TOCA1 HR1 but not vice versa. To extend these studies to a more complex system and to assess the ability of TOCA1 HR1 to compete with full-length N-WASP, pyrene actin assays were employed. These assays, described in detail elsewhere, were carried out using pyrene actin-supplemented Xenopus extracts into which exogenous TOCA1 HR1 domain or N-WASP GBD was added, to assess their effects on actin polymerization. Actin polymerization in all cases was initiated by the addition of PI(4,5)P2-containing liposomes. Actin polymerization triggered by the addition of PI(4,5)P2-containing liposomes has previously been shown to depend on TOCA1 and N-WASP. Endogenous N-WASP is present at ∼100 nm in Xenopus extracts, whereas TOCA1 is present at a 10-fold lower concentration than N-WASP. The addition of the isolated N-WASP GBD significantly inhibited the polymerization of actin at concentrations as low as 100 nm and completely abolished polymerization at higher concentrations (Fig. 8). The GBD presumably acts as a dominant negative, sequestering endogenous Cdc42 and preventing endogenous full-length N-WASP from binding and becoming activated. The addition of the TOCA1 HR1 domain to 100 μm had no significant effect on the rate of actin polymerization or maximum fluorescence. This is consistent with endogenous N-WASP, activated by other components of the assay, outcompeting the TOCA1 HR1 domain for Cdc42 binding. Actin polymerization downstream of Cdc42·N-WASP·TOCA1 is inhibited by excess N-WASP GBD but not by the TOCA1 HR1 domain. Fluorescence curves show actin polymerization in the presence of increasing concentrations of N-WASP GBD or TOCA1 HR1 domain as indicated. Maximal rates of actin polymerization derived from the linear region of the curves are represented in bar charts below. Error bars, S.E. Discussion The Cdc42-TOCA1 Interaction The TOCA1 HR1 domain alone is sufficient for Cdc42 binding in vitro, yet the affinity of the TOCA1 HR1 domain for Cdc42 is remarkably low (Kd ≈ 5 μm). This is over 100 times lower than that of the N-WASP GBD (Kd = 37 nm) and considerably lower than other known G protein-HR1 domain interactions. The polybasic tract within the C-terminal region of Cdc42 does not appear to be required for binding to TOCA1, which is in contrast to the interaction between Rac1 and the HR1b domain of PRK1 but more similar to the PRK1 HR1a-RhoA interaction. A single binding interface on both the HR1 domain and Cdc42 can be concluded from the data presented here. Furthermore, the interfaces are comparable with those of other G protein-HR1 interactions (Fig. 4), and the lowest energy model produced in rigid body docking resembles previously studied G protein·HR1 complexes (Fig. 6). It seems, therefore, that the interaction, despite its relatively low affinity, is specific and sterically similar to other HR1 domain-G protein interactions. The TOCA1 HR1 domain is a left-handed coiled-coil comparable with other known HR1 domains. A short region N-terminal to the coiled-coil exhibits a series of turns and contacts residues of both helices of the coiled-coil (Fig. 3). The corresponding sequence in CIP4 also includes a series of turns but is flexible, whereas in the HR1a domain of PRK1, the equivalent region adopts an α-helical structure that packs against the coiled-coil. The contacts between the N-terminal region and the coiled-coil are predominantly hydrophobic in both cases, but sequence-specific contacts do not appear to be conserved. This region is distant from the G protein-binding interface of the HR1 domains, so the structural differences may relate to the structure and regulation of these domains rather than their G protein interactions. The interhelical loops of TOCA1 and CIP4 differ from the same region in the HR1 domains of PRK1 in that they are longer and contain two short stretches of 310-helix. This region lies within the G protein-binding surface of all of the HR1 domains (Fig. 4D). TOCA1 and CIP4 both bind weakly to Cdc42, whereas the HR1a domain of PRK1 binds tightly to RhoA and Rac1, and the HR1b domain binds to Rac1. The structural features shared by TOCA1 and CIP4 may therefore be related to Cdc42 binding specificity and the low affinities. In free TOCA1, the side chains of the interhelical region make extensive contacts with residues in helix 1. Many of these residues are significantly affected in the presence of Cdc42, so it is likely that the conformation of this loop is altered in the Cdc42 complex. These observations therefore provide a molecular mechanism whereby mutation of Met383-Gly384-Asp385 to Ile383-Ser384-Thr385 abolishes TOCA1 binding to Cdc42. The lowest energy model produced by HADDOCK using ambiguous interaction restraints from the titration data resembled the NMR structures of RhoA and Rac1 in complex with their HR1 domain partners. Some speculative conclusions can be made based on this model. For example, Phe-56Cdc42, which is not visible in free Cdc42 or Cdc42·HR1TOCA1, is close to the TOCA1 HR1 (Fig. 6A). Phe-56Cdc42, which is a Trp in both Rac1 and RhoA (Fig. 6C), is thought to pack behind switch I when Cdc42 interacts with ACK, maintaining the switch in a binding-competent orientation. This residue has also been identified as important for Cdc42-WASP binding. Phe-56Cdc42 is therefore likely to be involved in the Cdc42-TOCA1 interaction, probably by stabilizing the position of switch I. Some residues that are affected in the Cdc42·HR1TOCA1 complex but do not correspond to contact residues of RhoA or Rac1 (Fig. 6C) may contact HR1TOCA1 directly (Fig. 6D). Gln-2Cdc42, which has also been identified as a contact residue in the Cdc42·ACK complex, contacts Val-376TOCA1 and Asn-380TOCA1 in the model and disrupts the contacts between the interhelical loop and the first helix of the TOCA1 coiled-coil. Thr-52Cdc42, which has also been identified as making minor contacts with ACK, falls near the side chains of HR1TOCA1 helix 1, particularly Lys-372TOCA1, whereas the equivalent position in Rac1 is Asn-52Rac1. N52T is one of a combination of seven residues found to confer ACK binding on Rac1 and so may represent a specific Cdc42-effector contact residue. The position equivalent to Lys-372TOCA1 in PRK1 is Glu-58HR1a or Gln-151HR1b. Thr-52Cdc42-Lys-372TOCA1 may therefore represent a specific Cdc42-HR1TOCA1 contact. Arg-68Cdc42 of switch II is positioned close to Glu-395TOCA1 (Fig. 6D), suggesting a direct electrostatic contact between switch II of Cdc42 and helix 2 of the HR1 domain. The equivalent Arg in Rac1 and RhoA is pointing away from the HR1 domains of PRK1. The importance of this residue in the Cdc42-TOCA1 interaction remains unclear, although its mutation reduces binding to RhoGAP, suggesting that it can be involved in Cdc42 interactions. The solution structure of the TOCA1 HR1 domain presented here, along with the model of the HR1TOCA1·Cdc42 complex is consistent with a conserved mode of binding across the known HR1 domain-Rho family interactions, despite their differing affinities. The weak binding prevented detailed structural and thermodynamic studies of the complex. Nonetheless, structural studies of the TOCA1 HR1 domain, combined with chemical shift mapping, have highlighted some potentially interesting differences between Cdc42-HR1TOCA1 and RhoA/Rac1-HR1PRK1 binding. We have previously postulated that the inherent flexibility of HR1 domains contributes to their ability to bind to different Rho family G proteins, with Rho-binding HR1 domains displaying increased flexibility, reflected in their lower melting temperatures (Tm) and Rac binders being more rigid. The Tm of the TOCA1 HR1 domain is 61.9 °C (data not shown), which is the highest Tm that we have measured for an HR1 domain thus far. As such, the ability of the TOCA1 HR1 domain to bind to Cdc42 (a close relative of Rac1 rather than RhoA) fits this trend. An investigation into the local motions, particularly in the G protein-binding regions, may offer further insight into the differential specificities and affinities of G protein-HR1 domain interactions. Significance of a Weak, Transient Interaction The low affinity of the Cdc42-HR1TOCA1 interaction is consistent with a tightly spatially and temporally regulated pathway, requiring combinatorial signals leading to a series of coincident weak interactions that elicit full activation. The HR1 domains from other TOCA family members, CIP4 and FBP17, also bind at low micromolar affinities to Cdc42, so the low affinity interaction appears to be commonplace among this family of HR1 domain proteins, in contrast to the PRK family. Weak, transient protein-protein interactions are functionally significant in several systems; for example, the binding of adaptor proteins to protein cargo during the formation of clathrin-coated vesicles in endocytosis involves multiple interactions of micromolar affinity. The low affinity of the HR1TOCA1-Cdc42 interaction in the context of the physiological concentration of TOCA1 in Xenopus extracts (∼10 nm) suggests that binding between TOCA1 and Cdc42 is likely to occur in vivo only when TOCA1 is at high local concentrations and membrane-localized and therefore in close proximity to activated Cdc42. Evidence suggests that the TOCA family of proteins are recruited to the membrane via an interaction between their F-BAR domain and specific signaling lipids. For example, electrostatic interactions between the F-BAR domain and the membrane are required for TOCA1 recruitment to membrane vesicles and tubules, and TOCA1-dependent actin polymerization is known to depend specifically on PI(4,5)P2. Furthermore, the isolated F-BAR domain of FBP17 has been shown to induce membrane tubulation of brain liposomes and BAR domain proteins that promote tubulation cluster on membranes at high densities. Once at the membrane, high local concentrations of TOCA1 could exceed the Kd of F-BAR dimerization (likely to be comparable with that of the FCHo2 F-BAR domain (2.5 μm)) and that of the Cdc42-HR1TOCA1 interaction. Cdc42-HR1TOCA1 binding would then be favorable, as long as coincident activation of Cdc42 had occurred, leading to stabilization of TOCA1 at the membrane and downstream activation of N-WASP. It has been postulated that WASP and N-WASP exist in equilibrium between folded (inactive) and unfolded (active) forms, and the affinity of Cdc42 for the unfolded WASP proteins is significantly enhanced. The unfolded, high affinity state of WASP is represented by a short peptide, the GBD, which binds with a low nanomolar affinity to Cdc42. In contrast, the best estimate of the affinity of full-length WASP for Cdc42 is low micromolar. In the inactive state of WASP, the actin- and Arp2/3-binding VCA domain contacts the GBD, competing for Cdc42 binding. The high affinity of Cdc42 for the unfolded, active form pushes the equilibrium in favor of (N-)WASP activation. Binding of PI(4,5)P2 to the basic region just N-terminal to the GBD further favors the active conformation. A substantial body of data has illuminated the complex regulation of WASP/N-WASP proteins, and current evidence suggests that these allosteric activation mechanisms and oligomerization combine to regulate WASP activity, allowing the synchronization and integration of multiple potential activation signals (reviewed in Ref.). Our data are easily reconciled with this model. We envisage that TOCA1 is first recruited to the appropriate membrane in response to PI(4,5)P2 via its F-BAR domain, where the local increase in concentration favors F-BAR-mediated dimerization of TOCA1. Cdc42 is activated in response to co-incident signals and can then bind to TOCA1, further stabilizing TOCA1 at the membrane. TOCA1 can then recruit N-WASP via an interaction between its SH3 domain and the N-WASP proline-rich region. The recruitment of N-WASP alone and of the N-WASP·WIP complex by TOCA1 and FBP17 has been demonstrated. WIP inhibits the activation of N-WASP by Cdc42, an effect that is reversed by TOCA1. It may therefore be envisaged that WIP and TOCA1 exert opposing allosteric effects on N-WASP, with TOCA1 favoring the unfolded, active conformation of N-WASP and increasing its affinity for Cdc42. TOCA1 may also activate N-WASP by effective oligomerization because clustering of TOCA1 at the membrane following coincident interactions with PI(4,5)P2 and Cdc42 would in turn lead to clustering of N-WASP, in addition to pushing the equilibrium toward the unfolded, active state. In a cellular context, full-length TOCA1 and N-WASP are likely to have similar affinities for active Cdc42, but in the unfolded, active conformation, the affinity of N-WASP for Cdc42 dramatically increases. Our binding data suggest that TOCA1 HR1 binding is not allosterically regulated, and our NMR data, along with the high stability of TOCA1 HR1, suggest that there is no widespread conformational change in the presence of Cdc42. As full-length TOCA1 and the isolated HR1 domain bind Cdc42 with similar affinities, the N-WASP-Cdc42 interaction will be favored because the N-WASP GBD can easily outcompete the TOCA1 HR1 for Cdc42. A combination of allosteric activation by PI(4,5)P2, activated Cdc42 and TOCA1, and oligomeric activation implemented by TOCA1 would lead to full activation of N-WASP and downstream actin polymerization. In such an array of molecules localized to a discrete region of the membrane, it is plausible that WASP could bind to a second Cdc42 molecule rather than displacing TOCA1 from its cognate Cdc42. Our NMR and affinity data, however, are consistent with displacement of the TOCA1 HR1 by the N-WASP GBD. Furthermore, TOCA1 is required for Cdc42-mediated activation of N-WASP·WIP, implying that it may not be possible for Cdc42 to bind and activate N-WASP prior to TOCA1-Cdc42 binding. The commonly used MGD → IST (Cdc42-binding deficient) mutant of TOCA1 has a reduced ability to activate the N-WASP·WIP complex, further indicating the importance of the Cdc42-HR1TOCA1 interaction prior to downstream activation of N-WASP. In light of this, we favor an “effector handover” scheme whereby TOCA1 interacts with Cdc42 prior to N-WASP activation, after which N-WASP displaces TOCA1 from its bound Cdc42 in order to be fully activated rather than binding a second Cdc42 molecule. Potentially, the TOCA1-Cdc42 interaction functions to position N-WASP and Cdc42 such that they are poised to interact with high affinity. The concomitant release of TOCA1 from Cdc42 while still bound to N-WASP presumably enhances the ability of TOCA1 to further activate N-WASP·WIP-induced actin polymerization. There is an advantage to such an effector handover, in that N-WASP would only be robustly recruited when F-BAR domains are already present. Hence, actin polymerization cannot occur until F-BAR domains are poised for membrane distortion. Our model of the Cdc42·HR1TOCA1 complex indicates a mechanism by which such a handover could take place (Fig. 9) because it shows that the effector binding sites only partially overlap on Cdc42. The lysine residues thought to be involved in an electrostatic steering mechanism in WASP-Cdc42 binding are conserved in N-WASP and would be able to interact with Cdc42 even when the TOCA1 HR1 domain is already bound. It has been postulated that the initial interactions between this basic region and Cdc42 could stabilize the active conformation of WASP, leading to high affinity binding between the core CRIB and Cdc42. The region C-terminal to the core CRIB, required for maximal affinity binding, would then fully displace the TOCA1 HR1. A simplified model of the early stages of Cdc42·N-WASP·TOCA1-dependent actin polymerization.\nStep 1, TOCA1 is recruited to the membrane via its F-BAR domain and/or Cdc42 interactions. F-BAR oligomerization is expected to occur following membrane binding, but a single monomer is shown for clarity. Step 2, N-WASP exists in an inactive, folded conformation. The TOCA1 SH3 domain interacts with N-WASP, causing an activatory allosteric effect. The HR1TOCA1-Cdc42 and SH3TOCA1-N-WASP interactions position Cdc42 and N-WASP for binding. Step 3, electrostatic interactions between Cdc42 and the basic region upstream of the CRIB initiate Cdc42·N-WASP binding. Step 4, the core CRIB binds with high affinity while the region C-terminal to the CRIB displaces the TOCA1 HR1 domain and increases the affinity of the N-WASP-Cdc42 interaction further. The VCA domain is released for downstream interactions, and actin polymerization proceeds. WH1, WASP homology 1 domain; PP, proline-rich region; VCA, verprolin homology, cofilin homology, acidic region. In conclusion, the data presented here show that the TOCA1 HR1 domain is sufficient for Cdc42 binding in vitro and that the interaction is of micromolar affinity, lower than that of other G protein-HR1 domain interactions. The analogous HR1 domains from other TOCA1 family members, FBP17 and CIP4, also exhibit micromolar affinity for Cdc42. A role for the TOCA1-, FBP17-, and CIP4-Cdc42 interactions in the recruitment of these proteins to the membrane therefore appears unlikely. Instead, our findings agree with earlier suggestions that the F-BAR domain is responsible for membrane recruitment. The role of the Cdc42-TOCA1 interaction remains somewhat elusive, but it may serve to position activated Cdc42 and N-WASP to allow full activation of N-WASP and as such serve to couple F-BAR-mediated membrane deformation with N-WASP activation. We envisage a complex interplay of equilibria between free and bound, active and inactive Cdc42, TOCA family, and WASP family proteins, facilitating a tightly spatially and temporally regulated pathway requiring numerous simultaneous events in order to achieve appropriate and robust activation of the downstream pathway. Our data are therefore easily reconciled with the dynamic instability models described in relation to the formation of endocytic vesicles and with the current data pertaining to the complex activation of WASP/N-WASP pathways by allosteric and oligomeric effects. It is clear from the data presented here that TOCA1 and N-WASP do not bind Cdc42 simultaneously and that N-WASP is likely to outcompete TOCA1 for Cdc42 binding. We therefore postulate an effector handover mechanism based on current evidence surrounding WASP/N-WASP activation and our model of the Cdc42·HR1TOCA1 complex. The displacement of the TOCA1 HR1 domain from Cdc42 by N-WASP may represent a unidirectional step in the pathway of Cdc42·N-WASP·TOCA1-dependent actin assembly. Author Contributions J. R. W. generated constructs and proteins, set up NMR experiments, analyzed NMR data, and performed binding experiments; D. N. set up NMR experiments; H. M. F. generated longer TOCA clones and proteins; J. L. G. supervised the pyrene actin assays; D. O. supervised the protein binding assays; and H. R. M. performed NMR experiments and analyzed NMR data. J. R. W., D. O., and H. R. M. wrote the paper with input from all authors. The authors declare that they have no conflicts of interest with the contents of this article. The atomic coordinates and structure factors (code 5FRG) have been deposited in the Protein Data Bank (http://wwpdb.org/). D. Owen, unpublished data. H. R. Mott and D. Owen, unpublished data. PRK protein kinase C related kinase WASP Wiskott-Aldrich syndrome protein TOCA transducer of Cdc42-dependent actin assembly protein N-WASP neural Wiskott-Aldrich syndrome protein PI(4,5)P2 phosphatidylinositol 4,5-bisphosphate HR1 homology region 1 F-BAR Fes/CIP4 homology BAR SH3 Src homology 3 CRIB Cdc42- and Rac-interactive binding CIP4 Cdc42-interacting protein 4 MBP maltose-binding protein GBD G protein binding domain SPA scintillation proximity assay PAK p21-activated kinase ACK activated Cdc42-associated kinase HSQC heteronuclear single quantum correlation GMPPNP guanosine 5′-[β,γ-imido] triphosphate GTPγS guanosine 5′-3-O-(thio)triphosphate GBD G protein-binding domain CSP chemical shift perturbation PDB Protein Data Bank. 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filopodial formation to endocytosis Drosophila Cip4/Toca-1 integrates membrane trafficking and actin dynamics through WASP and SCAR/WAVE Requirements for F-BAR proteins TOCA-1 and TOCA-2 in actin dynamics and membrane trafficking during Caenorhabditis elegans oocyte growth and embryonic epidermal morphogenesis Cdc42 interaction with N-WASP and Toca-1 regulates membrane tubulation, vesicle formation and vesicle motility: implications for endocytosis Self-assembly of filopodia-like structures on supported lipid bilayers Phosphoinositides and membrane curvature switch the mode of actin polymerization via selective recruitment of toca-1 and Snx9 Coordination between the actin cytoskeleton and membrane deformation by a novel membrane tubulation domain of PCH proteins is involved in endocytosis A TOCA/CDC-42/PAR/WAVE functional module required for retrograde endocytic recycling Regulation of neuronal morphology by Toca-1, an F-BAR/EFC protein that induces plasma membrane invagination Nuclear actin polymerization is required for transcriptional reprogramming of Oct4 by oocytes A complex of ZO-1 and the BAR-domain protein TOCA-1 regulates actin assembly at the tight junction Transducer of Cdc42-dependent actin assembly promotes epidermal growth factor-induced cell motility and invasiveness Transducer of Cdc42-dependent actin assembly promotes breast cancer invasion and metastasis Dynamin and the actin cytoskeleton cooperatively regulate plasma membrane invagination by BAR and F-BAR proteins Structure and analysis of FCHo2 F-BAR domain: a dimerizing and membrane recruitment module that effects membrane curvature The structural basis of Rho effector recognition revealed by the crystal structure of human RhoA complexed with the effector domain of PKN/PRK1 Molecular dissection of the interaction between the small G proteins Rac1 and RhoA and protein kinase C-related kinase 1 (PRK1) Mutational analysis reveals a single binding interface between RhoA and its effector, PRK1 Differential binding of RhoA, RhoB, and RhoC to protein kinase C-related kinase (PRK) isoforms PRK1, PRK2, and PRK3: PRKs have the highest affinity for RhoB The Rac1 polybasic region is required for interaction with its effector PRK1 The NMR structure of the TC10- and Cdc42-interacting domain of CIP4 A conserved binding motif defines numerous candidate target proteins for both Cdc42 and Rac GTPases Structure of Cdc42 in complex with the GTPase-binding domain of the “Wiskott-Aldrich syndrome” protein Residues in Cdc42 that specify binding to individual CRIB effector proteins The structure of binder of Arl2 (BART) reveals a novel G protein binding domain: implications for function Delineation of the Cdc42/Rac-binding domain of p21-activated kinase The IQGAP1-Rac1 and IQGAP1-Cdc42 interactions: interfaces differ between the complexes 1H, 13C and 15N resonance assignments of the Cdc42-binding domain of TOCA1 The CCPN data model for NMR spectroscopy: development of a software pipeline ARIA2: automated NOE assignment and data integration in NMR structure calculation Protein backbone and side chain torsion angles predicted from NMR chemical shifts using artificial neural networks  Triggering actin polymerization in Xenopus egg extracts from phosphoinositide-containing lipid bilayers Activation of the WAVE complex by coincident signals controls actin assembly Structure of the small G protein Cdc42 bound to the GTPase-binding domain of ACK Structural basis for membrane binding specificity of the Bin/Amphiphysin/Rvs (BAR) domain of Arfaptin-2 determined by Arl1 GTPase Targeting and activation of Rac1 are mediated by the exchange factor β-Pix STRIDE: a web server for secondary structure assignment from known atomic coordinates of proteins SOFAST-HMQC experiments for recording two-dimensional heteronuclear correlation spectra of proteins within a few seconds The RalB-RLIP76 complex reveals a novel mode of Ral-effector interaction The HADDOCK web server for data-driven biomolecular docking The Cdc42/Rac interactive binding region motif of the Wiskott Aldrich syndrome protein (WASP) is necessary but not sufficient for tight binding to Cdc42 and structure formation An electrostatic steering mechanism of Cdc42 recognition by Wiskott-Aldrich syndrome proteins Double mutant cycle thermodynamic analysis of the hydrophobic Cdc42-ACK protein-protein interaction Single-molecule dynamics reveals cooperative binding-folding in protein recognition Specificity determinants on Cdc42 for binding its effector protein ACK Transient protein-protein interactions: structural, functional, and network properties Transient protein-protein interactions Integrating molecular and network biology to decode endocytosis Evolving nature of the AP2 α-appendage hub during clathrin-coated vesicle endocytosis Phosphatidylinositol-(4,5)-bisphosphate regulates sorting signal recognition by the clathrin-associated adaptor complex AP2 The nucleotide switch in Cdc42 modulates coupling between the GTPase-binding and allosteric equilibria of Wiskott-Aldrich syndrome protein A two-state allosteric model for autoinhibition rationalizes WASP signal integration and targeting Direct binding of the verprolin-homology domain in N-WASP to actin is essential for cytoskeletal 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+[{"sourceid":"4937829","sourcedb":"","project":"","target":"","text":"Visualizing chaperone-assisted protein folding Challenges in determining the structures of heterogeneous and dynamic protein complexes have greatly hampered past efforts to obtain a mechanistic understanding of many important biological processes. One such process is chaperone-assisted protein folding, where obtaining structural ensembles of chaperone:substrate complexes would ultimately reveal how chaperones help proteins fold into their native state. To address this problem, we devised a novel structural biology approach based on X-ray crystallography, termed Residual Electron and Anomalous Density (READ). READ enabled us to visualize even sparsely populated conformations of the substrate protein immunity protein 7 (Im7) in complex with the E. coli chaperone Spy. This study resulted in a series of snapshots depicting the various folding states of Im7 while bound to Spy. The ensemble shows that Spy-associated Im7 samples conformations ranging from unfolded to partially folded and native-like states, and reveals how a substrate can explore its folding landscape while bound to a chaperone. High-resolution structural models of protein-protein interactions are critical for obtaining mechanistic insights into biological processes. However, many protein-protein interactions are highly dynamic, making it difficult to obtain high-resolution data. Particularly challenging are interactions of intrinsically or conditionally disordered sections of proteins with their partner proteins. Recent advances in X-ray crystallography and NMR spectroscopy continue to improve our ability to analyze biomolecules that exist in multiple conformations. X-ray crystallography has historically provided valuable information on small-scale conformational changes, but observing large-amplitude heterogeneous conformational changes often falls beyond the reach of current crystallographic techniques. NMR can theoretically be used to determine heterogeneous ensembles, but in practice, this proves to be very challenging. Despite the importance of understanding how proteins fold into their native state within the cell, our knowledge about this critical process remains limited. It is clear that molecular chaperones aid in protein folding. However, exactly how they facilitate the folding process is still being debated. Structural characterization of chaperone-assisted protein folding likely would help bring clarity to this question. Structural models of chaperone-substrate complexes have recently begun to provide information as to how a chaperone can recognize its substrate. However, the impact that chaperones have on their substrates, and how these interactions affect the folding process remain largely unknown. For most chaperones, it is still unclear whether the chaperone actively participates in and affects the folding of the substrate proteins, or merely provides a suitable microenvironment enabling the substrate to fold on its own. This is a truly fundamental question in the chaperone field, and one that has eluded the community largely because of the highly dynamic nature of the chaperone-substrate complexes. To address this question, we investigated the ATP-independent Escherichia coli periplasmic chaperone Spy. Spy prevents protein aggregation and aids in protein folding under various stress conditions, including treatment with tannin and butanol. We originally discovered Spy by its ability to stabilize the protein-folding model Im7 in vivo and recently demonstrated that Im7 folds while associated with Spy. The crystal structure of Spy revealed that it forms a thin α-helical homodimeric cradle. Crosslinking and genetic experiments suggested that Spy interacts with substrates somewhere on its concave side. By using a novel X-ray crystallography-based approach to model disorder in crystal structures, we have now determined the high-resolution ensemble of the dynamic Spy:Im7 complex. This work provides a detailed view of chaperone-mediated protein folding and shows how substrates like Im7 find their native fold while bound to their chaperones. RESULTS Crystallizing the Spy:Im7 complex We reasoned that to obtain crystals of complexes between Spy (domain boundaries in Supplementary Fig. 1) and its substrate proteins, our best approach was to identify crystallization conditions that yielded Spy crystals in the presence of protein substrates but not in their absence. We therefore screened crystallization conditions for Spy with four different substrate proteins: a fragment of the largely unfolded bovine α-casein protein, wild-type (WT) E. coli Im7, an unfolded variant of Im7 (L18A L19A L37A), and the N-terminal half of Im7 (Im76-45), which encompasses the entire Spy-binding portion of Im7. We found conditions in which all four substrates co-crystallized with Spy, but in which Spy alone did not yield crystals. Subsequent crystal washing and dissolution experiments confirmed the presence of the substrates in the co-crystals (Supplementary Fig. 2). The crystals diffracted to ~1.8 Å resolution. We used Spy:Im76-45 selenomethionine crystals for phasing with single-wavelength anomalous diffraction (SAD) experiments, and used this solution to build the well-ordered Spy portions of all four complexes. However, modeling of the substrate in the complex proved to be a substantial challenge, as the electron density of the substrate was discontinuous and fragmented. Even the minimal binding portion of Im7 (Im76-45) showed highly dispersed electron density (Fig. 1a). We hypothesized that the fragmented density was due to multiple, partially occupied conformations of the substrate bound within the crystal. Such residual density is typically not considered usable by traditional X-ray crystallography methods. Thus, we developed a new approach to interpret the chaperone-bound substrate in multiple conformations. READ: a strategy to visualize heterogeneous and dynamic biomolecules To determine the structure of the substrate portion of these Spy:substrate complexes, we conceived of an approach that we term READ, for Residual Electron and Anomalous Density. We split this approach into five steps: (1) By using a well-diffracting Spy:substrate co-crystal, we first determined the structure of the folded domain of Spy and obtained high quality residual electron density within the dynamic regions of the substrate. (2) We then labeled individual residues in the flexible regions of the substrate with the strong anomalous scatterer iodine, which serves to locate these residues in three-dimensional space using their anomalous density. (3) We performed molecular dynamics (MD) simulations to generate a pool of energetically reasonable conformations of the dynamic complex and (4) applied a sample-and-select algorithm to determine the minimal set of substrate conformations that fit both the residual and anomalous density. (5) Finally, we validated the ensemble using multiple statistical tests. Importantly, even though we only labeled a subset of the residues in the flexible regions of the substrate with iodine, the residual electron density can provide spatial information on many of the other flexible residues. These two forms of data are therefore complementary: by labeling individual residues, one can locate them to specific points in space. The electron density then allowed us to connect the labeled residues of the substrate by confining the protein chain within regions of detectable density. In this way, the two forms of data together were able to describe multiple conformations of the substrate within the crystal. As described in detail below, we developed the READ method to uncover the ensemble of conformations that the Spy-binding domain of Im7 (i.e., Im76-45) adopts while bound to Spy. However, we believe that READ will prove generally applicable to visualizing heterogeneous and dynamic complexes that have previously escaped detailed structural analysis. Collecting READ data for the Spy:Im76-45 complex To apply the READ technique to the folding mechanism employed by the chaperone Spy, we selected Im76-45 for further investigation because NMR data suggested that Im76-45 could recapitulate unfolded, partially folded, and native-like states of Im7 (Supplementary Fig. 3). Moreover, binding experiments indicated that Im76-45 comprises the entire Spy-binding region. To introduce the anomalous scatterer iodine, we replaced eight Im76-45 residues with the non-canonical amino acid 4-iodophenylalanine (pI-Phe). Its strong anomalous scattering allowed us to track the positions of these individual Im76-45 residues one at a time, potentially even if the residue was found in several locations in the same crystal. We then co-crystallized Spy and the eight Im76-45 peptides, each of which harbored an individual pI-Phe substitution at one distinct position, and collected anomalous data for all eight Spy:Im76-45 complexes (Fig. 1B, Supplementary Table 1 Supplementary Dataset 1, and Supplementary Table 2). Consistent with our electron density map, we found that the majority of anomalous signals emerged in the cradle of Spy, implying that this is the likely Im7 substrate binding site. Consistent with the fragmented density, however, we observed multiple iodine positions for seven of the eight substituted residues. Together, these results indicated that the Im7 substrate binds Spy in multiple conformations. READ sample-and-select procedure To determine the structural ensemble that Im76-45 adopts while bound to Spy, we combined the residual electron density and the anomalous signals from our pI-Phe substituted Spy:Im76-45 complexes. To generate an accurate depiction of the chaperone-substrate interactions, we devised a selection protocol based on a sample-and-select procedure employed in NMR spectroscopy. This procedure iteratively constructs structural ensembles and then compares them to the experimental data. During each round of the selection, a genetic algorithm alters the ensemble and its agreement to the experimental data is re-evaluated. If successful, the selection identifies the smallest group of specific conformations that best fits the residual electron density and anomalous signals. The READ sample-and-select algorithm is diagrammed in Fig. 2. Prior to performing the selection, we generated a large and diverse pool of chaperone-substrate complexes using coarse-grained MD simulations in a pseudo-crystal environment (Fig. 2 and Supplementary Fig. 4). The coarse-grained simulations are based on a single-residue resolution model for protein folding and were extended here to describe Spy-Im76-45 binding events (Online Methods). The initial conditions of the binding simulations are not biased toward a particular conformation of the substrate or any specific chaperone-substrate interaction (Online Methods). Im76-45 binds and unbinds to Spy throughout the simulations. This strategy allows a wide range of substrate conformations to interact with the chaperone. From the MD simulations, we extracted ~10,000 diverse Spy:Im76-45 complexes to be used by the ensuing selection. Each complex within this pool comprises one Spy dimer bound to a single Im76-45 substrate. This pool was then used by the selection algorithm to identify the minimal ensemble that best satisfies both the residual electron and anomalous crystallographic data. The anomalous scattering portion of the selection uses our basic knowledge of pI-Phe geometry: the iodine is separated from its respective Cα atom in each coarse-grained conformer by 6.5 Å. The selection then picks ensembles that best reproduce the collection of iodine anomalous signals. Simultaneously, it uses the residual electron density to help choose ensembles. To make the electron density selection practical, we needed to develop a method to rapidly evaluate the agreement between the selected sub-ensembles and the experimental electron density on-the-fly during the selection procedure. To accomplish this task, we generated a compressed version of the experimental 2mFo−DFc electron density map for use in the selection. This process provided us with a target map that the ensuing selection tried to recapitulate. To reduce the extent of 3D space to be explored, this compressed map was created by only using density from regions of space significantly sampled by Im76-45 in the Spy:Im76-45 MD simulations. For each of the ~10,000 complexes in the coarse-grained MD pool, the electron density at the Cα positions of Im76-45 was extracted and used to construct an electron density map (Online Methods). These individual electron density maps from the separate conformers could then be combined (Fig. 2) and compared to the averaged experimental electron density map as part of the selection algorithm. This approach allowed us to simultaneously use both the iodine anomalous signals and the residual electron density in the selection procedure. The selection resulted in small ensembles from the MD pool that best fit the READ data (Fig. 1c,d). Before analyzing the details of the Spy:Im76-45 complex, we first engaged in a series of validation tests to verify the ensemble and selection procedure (Supplementary Note 1, Figures 1c,d, Supplemental Figures 5-7). Combined, these validation tests confirmed that the selection procedure and selected six-member ensemble recapitulate the experimental data. Of note, the final six-membered ensemble was the largest ensemble that could simultaneously decrease the RFree and pass the 10-fold cross-validation test. This ensemble depicts the conformations that the substrate Im76-45 adopts while bound to the chaperone Spy (Fig. 3 Supplementary Movie 1, and Table 1). Folding and interactions of Im7 while bound to Spy Our results showed that by using this novel READ approach, we were able to obtain structural information about the dynamic interaction of a chaperone with its substrate protein. We were particularly interested in finding answers to one of the most fundamental questions in chaperone biology—how does chaperone binding affect substrate structure and vice versa. By analyzing the individual structures of the six-member ensemble of Im76-45 bound to Spy, we observed that Im76-45 takes on several different conformations while bound. We found these conformations to be highly heterogeneous and to include unfolded, partially folded, and native-like states (Fig. 3). The ensemble primarily encompasses Im76-45 laying diagonally within the Spy cradle in several different orientations, but some conformations traverse as far as the tips or even extend over the side of the cradle (Figs. 3,4a). We constructed a contact map of the complex, which shows the frequency of interactions for chaperone-substrate residue pairs (Fig. 4). We found that the primary interaction sites on Spy reside at the N and C termini (Arg122, Thr124, and Phe29) as well as on the concave face of the chaperone (Arg61, Arg43, Lys47, His96, and Met46). The Spy-contacting residues comprise a mixture of charged, polar, and hydrophobic residues. Surprisingly, we noted that in the ensemble, Im76-45 interacts with only 38% of the hydrophobic residues in the Spy cradle, but interacts with 61% of the hydrophilic residues in the cradle. This mixture suggests the importance of both electrostatic and hydrophobic components in binding the Im76-45 ensemble. With respect to the substrate, we observed that nearly every residue in Im76-45 is in contact with Spy (Fig. 4a). However, we did notice that despite this uniformity, regions of Im76-45 preferentially interact with different regions in Spy (Fig. 4b). For example, the N-terminal half of Im76-45 binds more consistently in the Spy cradle, whereas the C-terminal half predominantly binds to the outer edges of Spy’s concave surface. Not unexpectedly, we found that as Im76-45 progresses from the unfolded to the native state, its interactions with Spy shift accordingly. Whereas the least-folded Im76-45 pose in the ensemble forms the most hydrophobic contacts with Spy (Fig. 3), the two most-folded conformations form the fewest hydrophobic contacts (Fig. 3). This shift in contacts is likely due to hydrophobic residues of Im76-45 preferentially forming intra-molecular contacts upon folding (i.e., hydrophobic collapse), effectively removing themselves from the interaction sites. The diversity of conformations and binding sites observed here emphasizes the dynamic and heterogeneous nature of the chaperone-substrate ensemble. Although we do not yet have time resolution data of these various snapshots of Im76-45, this ensemble illustrates how a substrate samples its folding landscape while bound to a chaperone. Spy changes conformation upon substrate binding Comparing the structure of Spy in its substrate-bound and apo states revealed that the Spy dimer also undergoes significant conformational changes upon substrate binding (Fig. 5a and Supplementary Movie 2). Upon substrate binding, the Spy dimer twists 9° about its center relative to its apo form. This twist yields asymmetry and results in substantially different interaction patterns in the two Spy monomers (Fig. 4b). It is possible that this twist serves to increase heterogeneity in Spy by providing more binding poses. Additionally, we observed that the linker region (residues 47–57) of Spy, which participates in substrate interaction, becomes mostly disordered upon binding the substrate. This increased disorder might explain how Spy is able to recognize and bind different substrates and/or differing conformations of the same substrate. Importantly, we observed the same structural changes in Spy regardless of which of the four substrates was bound (Fig. 5b, Table 1). The RMSD between the well-folded sections of Spy in the four chaperone-substrate complexes was very small, less than 0.3 Å. Combined with competition experiments showing that the substrates compete in solution for Spy binding (Fig. 5c and Supplementary Fig. 8), we conclude that all the tested substrates share the same overall Spy binding site. DISCUSSION To shed light on how chaperones interact with their substrates, we developed a novel structural biology method (READ) and applied it to determine a conformational ensemble of the chaperone Spy bound to substrate. As a substrate, we used Im76-45, the chaperone-interacting portion of the protein-folding model protein Im7. In the chaperone-bound ensemble, Im76-45 samples unfolded, partially folded, and native-like states. The ensemble provides an unprecedented description of the conformations that a substrate assumes while exploring its chaperone-associated folding landscape. This substrate-chaperone ensemble helps accomplish the longstanding goal of obtaining a detailed view of how a chaperone aids protein folding. We recently showed that Im7 can fold while remaining continuously bound to Spy. The high-resolution ensemble obtained here now provides insight into exactly how this occurs. The structures of our ensemble agree well with lower-resolution crosslinking data, which indicate that chaperone-substrate interactions primarily occur on the concave surface of Spy. The ensemble suggests a model in which Spy provides an amphipathic surface that allows substrate proteins to assume different conformations while bound to the chaperone. This model is consistent with previous studies postulating that the flexible binding of chaperones allows for substrate protein folding. The amphipathic concave surface of Spy likely facilitates this flexible binding and may be a crucial feature for Spy and potentially other chaperones, allowing them to bind multiple conformations of many different substrates. In contrast to Spy’s binding hotspots, Im76-45 displays substantially less specificity in its binding sites. Nearly all Im76-45 residues come in contact with Spy. Unfolded substrate conformers interact with Spy through both hydrophobic and hydrophilic interactions, whereas the binding of native-like states is mainly hydrophilic. This trend suggests that complex formation between an ATP-independent chaperone and its unfolded substrate may initially involve hydrophobic interactions, effectively shielding the exposed aggregation-sensitive hydrophobic regions in the substrate. Once the substrate begins to fold within this protected environment, it progressively buries its own hydrophobic residues, and its interactions with the chaperone shift towards becoming more electrostatic. Notably, the most frequent contacts between Spy and Im76-45 are charge-charge interactions. The negatively charged Im7 residues Glu21, Asp32, and Asp35 reside on the surface of Im7 and form interactions with Spy’s positively charged cradle in both the unfolded and native-like states. Residues Asp32 and Asp35 are close to each other in the folded state of Im7. This proximity likely causes electrostatic repulsion that destabilizes Im7’s native state. Interaction with Spy’s positively-charged residues likely relieves the charge repulsion between Asp32 and Asp35, promoting their compaction into a helical conformation. As inter-molecular hydrophobic interactions between Spy and the substrate become progressively replaced by intra-molecular interactions within the substrate, the affinity between chaperone and substrates could decrease, eventually leading to release of the folded client protein. Recently, we employed a genetic selection system to improve the chaperone activity of Spy. This selection resulted in “Super Spy” variants that were more effective at both preventing aggregation and promoting protein folding. In conjunction with our bound Im76-45 ensemble, these mutants now allowed us to investigate structural features important to chaperone function. Previous analysis revealed that the Super Spy variants either bound Im7 tighter than WT Spy, increased chaperone flexibility as measured via H/D exchange, or both. Our ensemble revealed that two of the Super Spy mutations (H96L and Q100L) form part of the chaperone contact surface that binds to Im76-45 (Fig. 4a). Moreover, our co-structure suggests that the L32P substitution, which increases Spy’s flexibility, could operate by unhinging the N-terminal helix and effectively expanding the size of the disordered linker. This possibility is supported by the Spy:substrate structures, in which the linker region becomes more flexible compared to the apo state (Fig. 6a). This expansion would increase the structural plasticity for substrate binding. By sampling multiple conformations, this linker region may allow diverse substrate conformations to be accommodated. Other Super Spy mutations (F115I and F115L) caused increased flexibility but not tighter substrate binding. This residue does not directly contact Im76-45 in our READ-derived ensemble. Instead, when Spy is bound to substrate, F115 engages in close CH⋯π hydrogen bonds with Tyr104 (Fig. 6b). This interaction presumably reduces the mobility of the C-terminal helix. The F115I/L substitutions would replace these hydrogen bonds with hydrophobic interactions that have little angular dependence. As a result, the C-terminus, and possibly also the flexible linker, is likely to become more flexible and thus more accommodating of different conformations of substrates. Overall, comparison of our ensemble to the Super Spy variants provides specific examples to corroborate the importance of conformational flexibility in chaperone-substrate interactions. Despite extensive studies, exactly how complex chaperone machines help proteins fold remains controversial. Our study indicates that the chaperone Spy employs a simple surface binding approach that allows the substrate to explore various conformations and form transiently favorable interactions while being protected from aggregation. We speculate that many other chaperones could utilize a similar strategy. ATP and co-chaperone dependencies may have emerged later through evolution to better modulate and control chaperone action. In addition to insights into chaperone function, this work presents a new method for determining heterogeneous structural ensembles via a hybrid methodology of X-ray crystallography and computational modeling. Heterogeneous dynamic complexes or disordered regions of single proteins, once considered solely approachable by NMR spectroscopy, can now be visualized through X-ray crystallography. Consequently, this technique could enable structural characterization of many important dynamic and heterogeneous biomolecular systems. ONLINE METHODS For computational methods, including simulations of Spy-substrate interactions, binning the residual Im7 electron density, ensemble selection, validation tests, and contact map generation, please see Supplementary Note 1. Spy truncation mutants’ construction and in vitro and in vivo activity measurements To facilitate crystallization, we used Spy 29-124, a truncated Spy version that removes the unstructured N- and C-terminal tails (full length Spy is 138 amino acids). To determine if these alterations impact Spy’s chaperone activity in vitro, we performed in vitro chaperone activity assays and found that they had no significant effect; these deletions also had only a minor effect on Spy’s ability to stabilize Im7 in vivo (Supplementary Fig. 1). The in vitro activity of Spy 29-124 was assessed using the aldolase refolding assay as previously described. Briefly, in the denaturing step, 100 μM aldolase was denatured in buffer containing 6.6 M GdmCl, 40 mM HEPES pH 7.5, and 50 mM NaCl overnight at 22 °C (room temperature). In the refolding step, denatured aldolase was diluted to 3 μM in refolding buffer (40 mM HEPES, 150 mM NaCl, 5 mM DTT pH 7.5) in the presence of 6 μM WT Spy or Spy 29-124 (Spy:aldolase = 2:1). As a control, an identical experiment without Spy added was also performed. The refolding temperature was 37 °C with continuous shaking. The refolding status was monitored at different time points (1 min, 4 min, 10 min, and 20 min) and tested by diluting the refolding sample by 15-fold into the reaction buffer (0.15 mM NADH, 2 mM F1,6-DP, 1.8 U/ml GDH/TPI, 40 mM HEPES, and 150 mM NaCl pH 7.5) at 28 °C. The absorbance was monitored for 1.5 min at 340 nm. The percentage refolding was calculated and averaged over three repeats. To determine the in vivo activity of the Spy mutants, the quantity of the unstable Im7 variant L53A I54A expressed in the periplasm was compared during Spy variant co-expression as previously described. Plasmid Spy (pTrc-spy) was used as the template for the construction of the variant plasmids of Spy for in vivo chaperone activity measurement (Supplementary Table 3). To use the native signal sequence of spy for the periplasmic export of the Spy variants, an NheI site was first introduced between the signal sequence and the mature protein coding region of Spy. The vector was then digested with NheI and BamHI, purified, and ligated with the linear fragments corresponding to truncated sequences (21–130, 24–130, 27–130, 30–130, and 33–130) of Spy. Cells containing a strain that expressed the unstable Im7 mutant IL53A I54A (pCDFTrc-ssIm7L53A I54A) were transformed with plasmids that expressed either WT or one of the five truncated Spy mutants and grown to mid-log phase in LB medium at 37 °C. Im7 L53A I54A and Spy expression were induced with various concentrations of IPTG for 2 h to compare the in vivo chaperone activity of WT Spy and the truncated Spy mutants at similar expression levels. Periplasmic fractions were prepared as previously described and were separated on 16% Tricine gel (Life Technologies Inc.). The bands corresponding to Spy and the C-terminal His-tagged Im7 were either directly visualized on Coomassie stained gels or determined by western blot using anti-His antibody (Abcam ab1187; validation provided on manufacturer’s website). Protein expression and purification The gene for spy 29-124 was amplified from plasmid pET28sumo-spy with primer 1 (5′-CGC GGG ATC CTT CAA AGA CCT GAA CCT GAC CG-3′) and primer 2 (5′-CGC GCT CGA GTT ATG TCA GAC GCT TCT CAA AAT TAG C-3′), and was cloned into pET28sumo via BamHI and XhoI sites. The H96L variant was made by Phusion site-directed mutagenesis (New England Biolabs). WT and H96L Spy 29-124 were expressed and purified as described previously with the exception that Ni-HisTrap columns (GE Healthcare) were utilized instead of the Ni-NTA beads and mini-chromatography column. ULP1 cleavage occurred following elution from the Ni-HisTrap column overnight at 4 °C while dialyzing to 40 mM Tris, 300 mM NaCl, pH 8.0. After dialysis, Spy was passed over the HisTrap column to remove the cleaved SUMO tag (20 mM imidazole was left over from the dialysis). Cleavage of the SUMO tag leaves a single serine in position 28 of Spy. The flow-through was then concentrated and diluted 5 times with 20 mM Tris, pH 8 for further purification on a HiTrap Q column. Spy has an isoelectric point of 9.5 and therefore was collected in the flow-through. The flow-through containing Spy was concentrated and diluted 5-fold with 50 mM sodium phosphate at pH 6.5 and passed over a HiTrap SP column. Spy was then eluted with a gradient from 0 M to 1 M NaCl. Re-buffering to the final reaction buffer was accomplished by gel filtration, passing the pooled and concentrated fractions containing Spy over a HiLoad 75 column in 40 mM HEPES, 100 mM NaCl, pH 7.5. Fractions containing Spy were then concentrated, frozen in liquid nitrogen, and stored at −80 °C. WT Im7, Im7 L18A L19A L37A H40W, and Im7 L18A L19A L37A were purified by the same protocol as Spy, but without the SP column step. In addition to WT Im7 and these various Im7 mutants, co-crystallization experiments extensively utilized Im76-45, a minimal Spy-binding segment that encompasses the first two helices of Im7 and contains 46% of the total Im7 sequence. It displays partial helicity when free in solution (Supplementary Fig. 3). The 6-45 portion of Im7 (H2N-SISDYTEAEFVQLLKEIEKENVAATDDVLD VLLEHFVKIT-OH), 4-iodophenlyalanine variants, and a peptide corresponding to a portion of bovine alpha casein S1 148-177 (Ac-ELFRQFYQLDAYPSGAWYYVPLGTQYTDAP-amide) were obtained from New England Peptide at ≥ 95% purity. Anomalous signals for residues E12, E14, L19, and E21 substitutions were determined using a peptide containing Im7 6–26, which was also obtained from New England Peptide at ≥ 95% purity. Protein crystallization Co-crystals of WT Spy 29-124 and Spy H96L 29-124 in complex with Im7 variants and casein were grown by vapor diffusion. 25–130 mg/ml dimer Spy was incubated with various Im7 or casein substrates at concentrations ranging from equimolar to three-fold excess substrate in 22%–33% PEG 3000, 0.88–1.0 M imidazole pH 8.0, and 40–310 mM zinc acetate at 20 °C. Crystals were flash frozen in liquid nitrogen using 35% PEG 3000 as a cryo-protectant. It is worthwhile to note that the flash freezing could somewhat bias the conformations observed in the crystal structure. However, we chose to freeze the crystal to provide us with the maximum capability to identify and interpret the iodine anomalous signals. Assessing presence of substrate in crystals Crystals were washed by sequential transfer between three to six 2 μl drops of mother liquor, incubating in each wash solution for 2–10 s in an effort to remove all surface bound and precipitated substrate protein before being dissolved for visualization by SDS-PAGE. Before loading, samples were boiled for 10 min in reducing loading buffer, and then loaded onto 16% Tricine gels. Wash samples and dissolved crystal samples were analyzed by Lumetein staining (Biotium) and Flamingo staining (Bio-Rad) per manufacturer’s instructions, and imaged using a FluorChem M Imager (ProteinSimple). X-ray crystallography Data were collected at the LS-CAT beamlines at the Advanced Photon Source at 100 K. SeMet and native Spy:Im76-45 crystals were collected at 12.7 keV and 9.7 keV, respectively. Spy:Casein 148-177, and Spy H96L:WT Im7 crystals were collected also collected at 12.7 keV. Data integration and scaling were performed with iMosflm and AIMLESS, respectively. As molecular replacement attempts using the previously published apo Spy structures (PDB IDs: 3O39 and 3OEO) were unsuccessful, the Spy:Im76-45 complex was solved using Se-SAD phasing with SeMet-Spy, followed by density modification and initial model building by AutoSol in Phenix. The initial model was completed and refined using the native Spy:Im76-45 complex data. The rest of the structures were built using the native Spy:Im76-45 structure as a molecular replacement search model. Refinements, including TLS refinement, were performed using COOT and Phenix. All refined structures were validated using the Molprobity server, with Clashscores ranking better than the 90th percentile for all structures. Structural figures were rendered using PyMOL and UCSF Chimera, and movies generated using UCSF Chimera. Several partially occupied zinc atoms were observed in the crystal structure. Although some of these zinc atoms could also potentially modelled as water molecules, doing so resulted in an increase in the RFree. Additionally, a section of density near His A96 that is potentially partially occupied by a combination of water, Spy linker region, and possibly zinc, was modelled as containing water molecules. Spy H96L:Im76-45 was employed for iodine anomalous scattering experiments due to increased robustness and reproducibility of the crystals. The expected anomalous scatterers in the structures were S in methionine residues of Spy, Zn from the crystallization buffer, and I in the single pI-Phe residue of each synthetic Im76-45 peptide. Each I site is expected to be partially occupied as Im76-45 had diffuse density corresponding to multiple, partially occupied conformers; the Zn sites also may be partially occupied. To identify I, S, and Zn atomic positions using anomalous scattering, datasets were collected at 6.5 keV and 14.0 keV at 100 K using the ID-D beamline at LS-CAT. Anomalous difference maps for initial anomalous signal screening were calculated with phases from a molecular replacement search using the native Spy:Im76-45 (with no Im76-45 built in) complex as the search model. Anomalous difference maps calculated with the 14.0 keV data were used as controls to distinguish iodine from zinc atoms, as the iodine and zinc anomalous scattering factors are comparable at 14.0 keV, whereas at 6.5 keV, f″ is ~9-fold greater for iodine than for zinc. Anomalous differences were also collected and analyzed for a crystal of WT Spy 29-124:Im76-45 containing no iodine. The resulting anomalous difference map was inspected for peaks corresponding to sulfur, which were then excluded when selecting iodine peaks. Also, peaks that overlapped with Spy in the crystal lattice were excluded from analysis. As an initial screen for placing iodine atoms in the 6.5 keV anomalous difference maps, the median methionine sulfur signal was used as a cutoff for each individual map to control for varying data quality between crystals. Then, all anomalous atoms were refined in Phenix using anomalous group refinement. Refined B-factor of placed iodine ions was then used to estimate the positional fluctuation of the anomalous signals. This positional fluctuation was used as estimated error in the ensuing selection. A summary of all the anomalous signal heights (Supplementary Table 1) and anomalous difference maps (Supplemental Dataset 1) are displayed at varying contour levels for maximum clarity of iodine and methionine peak heights. Substrate binding to Spy The dissociation constant of Im76-45 was determined via a fluorescence-based competition experiment with Im7 L18A L19A L37A H40W, and its ability to compete with casein 148-177 for Spy binding was tested. Im7 L18A L19A L37A H40W was chosen for competition experiments due to its tight binding (Supplementary Fig. 8) and substantial fluorescence change upon binding. This mutant binds to Spy tighter than Im7 L18A L19A L37A. 10 μM Spy 29-124 dimer was mixed with 10 μM Im7 L18A L19A L37A H40W or casein 148-177 to form a 1:1 complex in a buffer containing 40 mM HEPES pH 7.5 and 100 mM NaCl at 22 °C. Complex formation was monitored with a QuantaMaster 400 (Photon Technology International) using the tryptophan fluorescence of Im7 L18A L19A L37A H40W. Naturally tryptophan-free Im76-45 was then titrated into the complex to compete with Im7 L18A L19A L37A H40W for Spy binding. The observed fluorescence intensity at 350 nm was plotted as a function of the logarithm of the Im76-45 or casein 148-177 concentration. The data were fit for a one-site-binding competition model (OriginLab 9.1):   where A1 and A2 are the maximum and minimum asymptotes, respectively, and x is the concentration of Im76-45. x0 is the apparent KD for Im76-45 based on its ability to compete with Im7 L18A L19A L37A H40W. Using the KD of Im7 L18A L19A L37A H40W binding to Spy 29-124, we then calculated the KD for Im76-45 binding to Spy 29-124 using the Cheng-Prusoff equation:   where L is the concentration of Im7 L18A L19A L37A H40W and KD is the dissociation constant for Im7 L18A L19A L37A H40W binding to Spy. Due to interaction between higher oligomer states of Im76-45 and casein 148-177 (Supplementary Fig. 8), the competition curve was unable to be fit for casein 148-177 competing with Im76-45. The stoichiometry of binding of casein 148-177 and Spy was determined by tryptophan fluorescence of the casein upon Spy 29-124 addition. Increasing concentrations of Spy 29-124 were titrated to 20 μM of casein 148-177 in 40 mM HEPES (pH 7.5), 100 mM NaCl, at 22 °C. Complex formation was monitored with a QuantaMaster 400 (Photon Technology International) using the tryptophan fluorescence of casein 148-177. The observed fluorescence intensity at 339 nm was plotted as a function of the Spy 29-124 dimer concentration and fit with a quadratic equation using Origin 9.1 (OriginLab). To determine the dissociation constant, increasing concentrations of Spy 29-124 were titrated to 0.25 μM of casein in 40 mM HEPES (pH 7.5), 100 mM NaCl, at 22 °C. Complex formation was monitored with a QuantaMaster 400 (Photon Technology International) using the tryptophan fluorescence of casein 148-177. The observed fluorescence intensity at 339 nm was corrected for dilution due to the titration and then plotted as a function of the Spy 29-124 dimer concentration. The data were fit using a square hyperbola function in Origin 9.1 (OriginLab):   where F is the recorded fluorescence signal, Fmax is the maximum fluorescence reached upon saturation of the complex, L is the concentration of free Spy in solution, KD is the dissociation constant, and C is a parameter for the offset. The calculated KD is an average of three independent repetitions. The measured dissociation constants for the different substrates ranged from 0.1 to 1 μM. Isothermal titration calorimetry (ITC) Spy 29-124 and Im7-L18A L19A L37A H40W were dialyzed overnight against 40 mM HEPES, 100 mM NaCl, pH 7.5. 165 μM Spy dimer was loaded into a syringe and titrated into a cell containing 15 μM Im7 L18A L19 AL37A H40W at 25 °C in an iTC200 (Malvern Instruments) with an injection interval of 120 s and an initial delay time of 60 s. The solution was stirred at 1000 rpm, and the reference power was set to 6 μcal s−1 in high feedback mode. Data analysis was conducted using a plugin for Origin 7 (OriginLab), the software provided by the manufacturer. Analytical ultracentrifugation Sedimentation velocity experiments for the Im76-45 and the bovine α-S1-casein peptide were performed using a Beckman Proteome Lab XL-I analytical ultracentrifuge (Beckman Coulter). Both peptides were first dialyzed against 40 mM HEPES, 100 mM NaCl, pH 7.5, then diluted to a concentration of 10 μM using the dialysis buffer. Samples were loaded into cells containing standard sector shaped 2-channel Epon centerpieces with 1.2 cm path-length (Beckman Coulter) and equilibrated to 22 °C for at least 1 h prior to sedimentation. All samples were spun at 48,000 rpm in a Beckman AN-50 Ti rotor, and the sedimentation of the protein was monitored continuously using interference optics, since the Im76-45 does not absorb strongly at 280 nm. Data analysis was conducted with SEDFIT (version 14.1), using the continuous c(s) distribution model. The confidence level for the maximum entropy (ME) regularization was set to 0.95. Buffer density and viscosity were calculated using SEDNTERP (http://sednterp.unh.edu/). Supplementary Material ACCESSION CODES Structures and datasets in this work have been deposited in the PDB under the IDs 5INA, 5IOG, 5IOE, and 5IOA. AUTHOR CONTRIBUTIONS Overall concept was conceived by S.H. and J.B. Experiments were designed by S.H., S.Q., J.B., R.T., H.B., and P.K. Experiments were performed by S.H., S.Q., P.K., R.M., and L.W. Analysis and computational modeling was designed by C.B., L.S., P.A., L.A., H.B., and S.H. Computational analysis was carried out by Q.X., S.H., L.S., L.A., P.A., P.K., and R.M. The manuscript was written primarily by S.H. and J.B., with assistance from L.S., L.A. and all other authors. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. Principles of protein-protein interactions: what are the preferred ways for proteins to interact? 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(a) 2mFo−DFc omit map of residual Im76-45 and flexible linker electron density contoured at 0.5 σ. This is the residual density that is used in the READ selection. (b) Composites of iodine positions detected from anomalous signals using pI-Phe substitutions, colored and numbered by sequence. Multiple iodine positions were detected for most residues. Agreement to the residual Im76-45 electron density (c) and anomalous iodine signals (d) for ensembles of varying size generated by randomly choosing from the MD pool (blue) and from the selection procedure (black). The agreement from back-calculating a subset of data excluded from the selection procedure is shown by the red curve (cross-validation). The cost function, χ2, decreases as the agreement to the experimental data increases and is defined in the Online Methods. Flowchart of the READ sample-and-select process. Spy:Im76-45 ensemble, arranged by RMSD to native state of Im76-45. Although the six-membered ensemble from the READ selection should be considered only as an ensemble, for clarity, the individual conformers are shown separately here. Spy is depicted as a gray surface and the Im76-45 conformer is shown as orange balls. Atoms that were either not directly selected in the READ procedure, or whose position could not be justified based on agreement with the residual electron density were removed, leading to non-contiguous sections. Dashed lines connect non-contiguous segments of the Im76-45 substrate. Residues of the Spy flexible linker region that fit the residual electron density are shown as larger gray spheres. Shown below each ensemble member is the RMSD of each conformer to the native state of Im76-45, as well as the percentage of contacts between Im76-45 and Spy that are hydrophobic. Contact maps of Spy:Im76-45 complex. (a) Spy:Im76-45 contact map projected onto the bound Spy dimer (above) and Im76-45 (below) structures. For clarity, Im76-45 is represented with a single conformation. The frequency plotted is calculated as the average contact frequency from Spy to every residue of Im76-45 and vice-versa. As the residues involved in contacts are more evenly distributed in Im76-45 compared to Spy, its contact map was amplified. (b) Detailed contact maps of Spy:Im76-45. Contacts to the two Spy monomers are depicted separately. Note that the flexible linker region of Spy (residues 47–57) is not represented in the 2D contact maps. Spy conformation changes upon substrate binding. (a) Overlay of apo Spy (PDB ID: 3O39, gray) and bound Spy (green). (b) Overlay of WT Spy bound to Im76-45 (green), H96L Spy bound to Im7 L18A L19 AL13A (blue), H96L Spy bound to WT Im7 (yellow), and WT Spy bound to casein (salmon). (c) Competition assay showing Im76-45 competes with Im7 L18A L19A L37A H40W for the same binding site on Spy (further substrate competition assays are shown in Supplementary Fig. 8). Error bars depict standard deviations of n=3 technical replicates. Flexibility of Spy linker region and effect of Super Spy mutants. (a) The Spy linker region adopts one dominant conformation in its apo state (PDB ID 3039, gray), but expands and adopts multiple conformations in bound states (green). (b) F115 and L32 tether Spy’s linker region to its cradle, decreasing Spy activity by limiting linker region flexibility. The Super Spy mutants F115L, F115I, and L32P are proposed to gain activity by increasing the flexibility or size of this linker region. L32, F115, and Y104 are rendered in purple to illustrate residues that are most affected by Super Spy mutations; CH⋯π hydrogen bonds are depicted by orange dashes. Crystallography Statistics \tSeMet Spy:Im76-45\tSpy:Im76-45\tSpy:Casein 148-177, substrate not modeled\tSpy H96L:Im7 L18A L19A L37A, substrate not modeled\tSpy H96L:WT Im7, substrate not modeled\t \tPDB ID\t\t5INA\t5IOG\t5IOE\t5IOA\t \tData collection\t \tSpace group\tP4122\tP4122\tP4122\tP4122\tP4122\t \tCell dimensions\t \t a, b, c (Å)\t42.9, 42.9, 259.3\t42.9, 42.9, 260.2\t43.0, 43.0, 258.2\t43.1, 43.1, 258.7\t43.1, 43.14, 260.2\t \t α, β, γ (°)\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t90, 90, 90\t \tResolution (Å)\t64.82–2.44(2.53–2.44)\t30.50–1.79(1.83–1.79)\t36.88–1.77 (1.80–1.77)\t30.48–1.87(1.91–1.87)\t33.21–1.87(1.91–1.87)\t \tRmerge (%)\t10.6(36)\t8.2(108)\t6.2(134)\t8.4(152)\t9.6(249)\t \tI/σ(I)\t15.1(6.8)\t7.0(1.1)\t15.3(1.6)\t13.8(1.8)\t13.2(1.3)\t \tCompleteness (%)\t100(100)\t94.0(90.1)\t99.9(99.5)\t100(100)\t96.8(93.1)\t \tRedundancy\t15.6(15.6)\t4.3(4.2)\t8.7(8.2)\t9.6(9.4)\t8.2(8.2)\t \tCC1/2\t\t0.998(0.689)\t0.999(0.745)\t0.999(0.676)\t0.998(0.606)\t \tRefinement\t \tResolution (Å)\t\t1.79\t1.77\t1.87\t1.87\t \tNo. of Reflections\t\t22583\t25052\t21505\t20838\t \tRwork/Rfree\t\t0.22/0.23\t0.21/0.24\t0.22/0.24\t0.21/0.25\t \tNo. of Atoms\t\t1765\t1669\t1715\t1653\t \t Protein\t\t1586\t1493\t1541\t1444\t \t Ligand/ion\t\t30\t56\t60\t30\t \t Water\t\t149\t120\t114\t179\t \tB-factors\t\t49.4\t48.5\t47.4\t39.2\t \t Protein\t\t49.0\t47.5\t46.3\t38.3\t \t Ligand/ion\t\t48.6\t65.9\t80.4\t62.9\t \t Water\t\t54.2\t51.9\t44.5\t42.1\t \tr.m.s. Deviations\t\t\t\t\t\t \t Bond lengths (Å)\t\t0.013\t0.013\t0.013\t0.014\t \t Bond angles (º)\t\t1.24\t1.30\t1.24\t1.39\t 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+[{"sourceid":"4968113","sourcedb":"","project":"","target":"","text":"Structural diversity in a human antibody germline library ABSTRACT To support antibody therapeutic development, the crystal structures of a set of 16 germline variants composed of 4 different kappa light chains paired with 4 different heavy chains have been determined. All four heavy chains of the antigen-binding fragments (Fabs) have the same complementarity-determining region (CDR) H3 that was reported in an earlier Fab structure. The structure analyses include comparisons of the overall structures, canonical structures of the CDRs and the VH:VL packing interactions. The CDR conformations for the most part are tightly clustered, especially for the ones with shorter lengths. The longer CDRs with tandem glycines or serines have more conformational diversity than the others. CDR H3, despite having the same amino acid sequence, exhibits the largest conformational diversity. About half of the structures have CDR H3 conformations similar to that of the parent; the others diverge significantly. One conclusion is that the CDR H3 conformations are influenced by both their amino acid sequence and their structural environment determined by the heavy and light chain pairing. The stem regions of 14 of the variant pairs are in the ‘kinked’ conformation, and only 2 are in the extended conformation. The packing of the VH and VL domains is consistent with our knowledge of antibody structure, and the tilt angles between these domains cover a range of 11 degrees. Two of 16 structures showed particularly large variations in the tilt angles when compared with the other pairings. The structures and their analyses provide a rich foundation for future antibody modeling and engineering efforts. Introduction At present, therapeutic antibodies are the largest class of biotherapeutic proteins that are in clinical trials. The use of monoclonal antibodies as therapeutics began in the early 1980s, and their composition has transitioned from murine antibodies to generally less immunogenic humanized and human antibodies. The technologies currently used to obtain human antibodies include transgenic mice containing human antibody repertoires, cloning directly from human B cells, and in vitro selection from antibody libraries using various display technologies. Once a candidate antibody is identified, protein engineering is usually required to produce a molecule with the right biophysical and functional properties. All engineering efforts are guided by our understanding of the atomic structures of antibodies. In such efforts, the crystal structure of the specific antibody may not be available, but modeling can be used to guide the engineering efforts. Today's antibody modeling approaches, which normally focus on the variable region, are being developed by the application of structural principles and insights that are evolving as our knowledge of antibody structures continues to expand. Our current structural knowledge of antibodies is based on a multitude of studies that used many techniques to gain insight into the functional and structural properties of this class of macromolecule. Five different antibody isotypes occur, IgG, IgD, IgE, IgA and IgM, and each isotype has a unique role in the adaptive immune system. IgG, IgD and IgE isotypes are composed of 2 heavy chains (HCs) and 2 light chains (LCs) linked through disulfide bonds, while IgA and IgM are double and quintuple versions of antibodies, respectively. Isotypes IgG, IgD and IgA each have 4 domains, one variable (V) and 3 constant (C) domains, while IgE and IgM each have the same 4 domains along with an additional C domain. These multimeric forms are linked with an additional J chain. The LCs that associate with the HCs are divided into 2 functionally indistinguishable classes, κ and λ. Both κ and λ polypeptide chains are composed of a single V domain and a single C domain. The heavy and light chains are composed of structural domains that have ∼110 amino acid residues. These domains have a common folding pattern often referred to as the “immunoglobulin fold,” formed by the packing together of 2 anti-parallel β-sheets. All immunoglobulin chains have an N-terminal V domain followed by 1 to 4 C domains, depending upon the chain type. In antibodies, the heavy and light chain V domains pack together forming the antigen combining site. This site, which interacts with the antigen (or target), is the focus of current antibody modeling efforts. This interaction site is composed of 6 complementarity-determining regions (CDRs) that were identified in early antibody amino acid sequence analyses to be hypervariable in nature, and thus are responsible for the sequence and structural diversity of our antibody repertoire. The sequence diversity of the CDR regions presents a substantial challenge to antibody modeling. However, an initial structural analysis of the combining sites of the small set of structures of immunoglobulin fragments available in the 1980s found that 5 of the 6 hypervariable loops or CDRs had canonical structures (a limited set of main-chain conformations). A CDR canonical structure is defined by its length and conserved residues located in the hypervariable loop and framework residues (V-region residues that are not part of the CDRs). Furthermore, studies of antibody sequences revealed that the total number of canonical structures are limited for each CDR, indicating possibly that antigen recognition may be affected by structural restrictions at the antigen-binding site. Later studies found that the CDR loop length is the primary determining factor of antigen-binding site topography because it is the primary factor for determining a canonical structure. Additional efforts have led to our current understanding that the LC CDRs L1, L2, and L3 have preferred sets of canonical structures based on length and amino acid sequence composition. This was also found to be the case for the H1 and H2 CDRs. Classification schemes for the canonical structures of these 5 CDRs have emerged and evolved as the number of depositions in the Protein Data Bank of Fab fragments of antibodies grow. Recently, a comprehensive CDR classification scheme was reported identifying 72 clusters of conformations observed in antibody structures. The knowledge and predictability of these CDR canonical structures have greatly advanced antibody modeling efforts. In contrast to CDRs L1, L2, L3, H1 and H2, no canonical structures have been observed for CDR H3, which is the most variable in length and amino acid sequence. Some clustering of conformations was observed for the shortest lengths; however, for the longer loops, only the portions nearest the framework (torso, stem or anchor region) were found to have defined conformations. In the torso region, 2 primary groups could be identified, which led to sequence-based rules that can predict with some degree of reliability the conformation of the stem region. The “kinked” or “bulged” conformation is the most prevalent, but an “extended” or “non-bulged” conformation is also, but less frequently, observed. The cataloging and development of the rules for predicting the conformation of the anchor region of CDR H3 continue to be refined, producing new insight into the CDR H3 conformations and new tools for antibody engineering. Current antibody modeling approaches take advantage of the most recent advances in homology modeling, the evolving understanding of the CDR canonical structures, the emerging rules for CDR H3 modeling and the growing body of antibody structural data available from the PDB. Recent antibody modeling assessments show continued improvement in the quality of the models being generated by a variety of modeling methods. Although antibody modeling is improving, the latest assessment revealed a number of challenges that need to be overcome to provide accurate 3-dimensional models of antibody V regions, including accuracies in the modeling of CDR H3. The need for improvement in this area was also highlighted in a recent study reporting an approach and results that may influence future antibody modeling efforts. One important finding of the antibody modeling assessments was that errors in the structural templates that are used as the basis for homology models can propagate into the final models, producing inaccuracies that may negatively influence the predictive nature of the V region model. To support antibody engineering and therapeutic development efforts, a phage library was designed and constructed based on a limited number of scaffolds built with frequently used human germ-line IGV and IGJ gene segments that encode antigen combining sites suitable for recognition of peptides and proteins. This Fab library is composed of 3 HC germlines, IGHV1-69 (H1-69), IGHV3-23 (H3-23) and IGHV5-51(H5-51), and 4 LC germlines (all κ), IGKV1-39 (L1-39), IGKV3-11 (L3-11), IGKV3-20 (L3-20) and IGKV4-1 (L4-1). Selection of these genes was based on the high frequency of their use and their cognate canonical structures that were found binding to peptides and proteins, as well as their ability to be expressed in bacteria and displayed on filamentous phage. The implementation of the library involves the diversification of the human germline genes to mimic that found in natural human libraries. The crystal structure determinations and structural analyses of all germline Fabs in the library described above along with the structures of a fourth HC germline, IGHV3-53 (H3-53), paired with the 4 LCs of the library have been carried out to support antibody therapeutic development. All 16 HCs of the Fabs have the same CDR H3 that was reported in an earlier Fab structure. This is the first systematic study of the same VH and VL structures in the context of different pairings. The structure analyses include comparisons of the overall structures, canonical structures of the L1, L2, L3, H1 and H2 CDRs, the structures of all CDR H3s, and the VH:VL packing interactions. The structures and their analyses provide a foundation for future antibody engineering and structure determination efforts. Results Crystal structures Crystal data, X-ray data, and refinement statistics. Fab\tH1-69:L1-39\tH1-69:L3-11\tH1-69:L3-20\tH1-69:L4-1\t \tPDB identifier\t5I15\t5I16\t5I17\t5I18\t \tCrystal Data\t \t \t \t \t \tCrystallization Solution\t \t \t \t \t \t Buffer, pH\t0.1 M MES- pH 6.5\t0.1 M MES pH 6.5\t0.1 M MES, pH 6.5\t0.1 M HEPES, pH 7.5\t \t Precipitant1\t5 M Na Formate\t25% PEG 3350\t2.0 M Amm Sulfate\t10% PEG 8000\t \t Additive1\t \t0.2 M Na Formate\t5% MPD\t8% EG\t \t Space Group\tP3121\tC2\tP422\tP4212\t \t Molecules/AU\t1\t2\t2\t1\t \t Unit Cell\t \t \t \t \t \t a(Å)\t129.2\t212.0\t152.5\t120.0\t \t b(Å)\t129.2\t55.1\t152.5\t120.0\t \t c(Å)\t91.8\t80.3\t123.4\t64.2\t \t β(°)\t90.0\t97.8\t90.0\t90.0\t \t γ(°)\t120.0\t90.0\t90.0\t90.0\t \t Vm (Å3/Da)\t4.67\t2.44\t3.77\t2.39\t \t Solvent Content (%)\t74\t50\t67\t48\t \tX-Ray Data2\t \t \t \t \t \t Resolution (Å)\t30-2.6 (2.7-2.6)\t30.0-1.9 (1.95-1.9)\t30.0-3.3 (3.4-3.3)\t30-1.9 (2.0-1.9)\t \t Measured Reflections\t136,745 (8,650)\t241,145 (16,580)\t237,504 (15,007)\t801,080 (19,309)\t \t Unique Reflections\t27,349 (1,730)\t71,932 (5,198)\t22,379 (1,590)\t35,965 (2,194)\t \t Completeness (%)\t99.3 (98.7)\t99.0 (97.3)\t99.5 (96.8)\t98.5 (82.8)\t \t Redundancy\t5.0 (5.0)\t3.4 (3.2)\t10.6 (9.4)\t22.3 (8.8)\t \t Rmerge\t0.048 (0.522)\t0.044 (0.245)\t0.086 (0.536)\t0.093 (0.231)\t \t \u003c I/σ \u003e\t21.2 (3.9)\t17.8 (4.7)\t25.5 (4.5)\t29.2 (8.1)\t \t B-factor (Å2)\t60.5\t33.2\t61.0\t19.6\t \tRefinement\t \t \t \t \t \t Resolution (Å)\t15-2.6\t15-1.9\t15-3.3\t15-1.9\t \t Number of Reflections\t26,238\t70,346\t21,197\t34,850\t \t Number of All Atoms\t3,224\t6,975\t6,398\t3,695\t \t Number of Waters\t2\t472\t0\t399\t \t R-factor (%)\t20.5\t19.2\t20.2\t16.7\t \t R-free (%)\t24.1\t22.2\t24.7\t21.3\t \tRMSD\t \t \t \t \t \t Bond Lengths (Å)\t0.006\t0.005\t0.005\t0.008\t \t Bond Angles (°)\t1.2\t1.1\t1.0\t1.1\t \t Mean B-factor (Å2)\t65.3\t34.4\t80.1\t20.0\t \tRamachandran Plot (%)\t \t \t \t \t \t Outliers\t0.0\t0.0\t0.9\t0.0\t \t Favored\t92.3\t96.9\t93.1\t96.9\t \t Abbreviations: Amm, ammonium;EG, ethylene glycol; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. (Continued) Crystal data, X-ray data, and refinement statistics. Fab\tH3-23:L1-39\tH3-23:L3-11\tH3-23:L3-20\tH3-23:L4-1\t \t PDB identifier\t5I19\t5I1A\t5I1C\t5I1D\t \tCrystal Data\t \t \t \t \t \tCrystallization Solution\t \t \t \t \t \t Buffer, pH\tNo Buffer\t0.1 M Na Acetate, pH 4.5\t0.1 M MES, pH 6.5\t0.1 M HEPES, pH 7.5\t \t Precipitant1\t20% PEG 3350\t2.0 M Amm Sulfate\t16% PEG 3350\t2.0 M Amm Sulfate\t \t Additive1\t0.2 M Li Citrate\t5% PEG 400\t0.2 M Amm Acetate\t2% PEG 400\t \t Space Group\tP41212\tP212121\tP6222\tP212121\t \t Molecules/AU\t1\t2\t1\t2\t \tUnit Cell\t \t \t \t \t \t a(Å)\t96.6\t60.9\t121.5\t62.7\t \t b(Å)\t96.6\t110.6\t121.5\t111.0\t \t c(Å)\t105.4\t158.9\t160.4\t160.0\t \t β(°)\t90\t90\t90\t90\t \t Vm (Å3/Da)\t2.60\t2.82\t3.60\t2.90\t \t Solvent Content (%)\t53\t56\t66\t57\t \tX-Ray Data2\t \t \t \t \t \t Resolution (Å)\t30-2.8 (2.9-2.8)\t30-2.0 (2.1-2.0)\t30-2.25 (2.3-2.25)\t30-2.0 (2.1-2.0)\t \t Measured Reflections\t177,681 (12,072)\t351,312 (8,634)\t887,349 (59,919)\t873,523 (49,118)\t \t Unique Reflections\t12,678 (899)\t58,989 (2,870)\t32,572 (2,300)\t75,540 (5,343)\t \t Completeness (%)\t99.5 (97.4)\t80.9 (54.2)\t96.9 (94.8)\t99.7 (96.9)\t \t Redundancy\t14.0 (13.4)\t6.0 (3.0)\t27.2 (26.1)\t11.6 (9.2)\t \t Rmerge\t0.091 (0.594)\t0.066 (0.204)\t0.086 (0.478)\t0.094 (0.488)\t \t \u003c I/σ \u003e\t31.2 (5.1)\t20.4 (4.6)\t37.0 (10.4)\t21.6 (5.0)\t \t B-factor (Å2)\t42.8\t27.1\t33.7\t29.4\t \tRefinement\t \t \t \t \t \t Resolution (Å)\t15-2.8\t15-2.0\t15-2.25\t15-2.0\t \t Number of Reflections\t11,972\t57,599\t31,411\t74,238\t \t Number of All Atoms\t3,234\t6,948\t3,472\t7,210\t \t Number of Waters\t0\t416\t222\t635\t \t R-factor (%)\t23.9\t20.5\t22.0\t21.6\t \t R-free (%)\t31.5\t25.5\t26.6\t25.1\t \tRMSD\t \t \t \t \t \t Bond Lengths (Å)\t0.009\t0.010\t0.005\t0.008\t \t Bond Angles (°)\t1.3\t1.3\t1.0\t1.1\t \t Mean B-factor (Å2)\t48.4\t36.7\t47.7\t46.4\t \tRamachandran Plot (%)\t \t \t \t \t \t Outliers\t0.0\t0.0\t0.0\t0.0\t \t Favored\t92.3\t96.8\t97.5\t97.6\t \t Abbreviations: Amm, ammonium; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. (Continued) Crystal data, X-ray data, and refinement statistics. Fab\tH3-53:L1-39\tH3-53:L3-11\tH3-53:L3-20\tH3-53:L4-1\t \tPDB indentifier\t5I1E\t5I1G\t5I1H\t5I1I\t \tCrystal Data\t \t \t \t \t \tCrystallization Solution\t \t \t \t \t \t Buffer, pH\tNo buffer\t0.1 M Na Acetate pH 4.5\t0.1 M Na Acetate pH 4.5\t0.1M MES, pH 6.5\t \t Precipitant1\t16% PEG 3350\t25% PEG 3350\t19% PEG 4000\t17% PEG 3350\t \t Additive1\t0.2 M Amm Sulfate 5% Dioxane\t0.2 M Li2SO4\t0.2 M Amm Sulfate\t0.2 M Na Formate, 5% MPD\t \t Space Group\tP6522\tP6522\tP6522\tP31\t \t Molecules/AU\t1\t1\t1\t1\t \tUnit Cell\t \t \t \t \t \t a(Å)\t89.4\t88.1\t89.4\t68.1\t \t b(Å)\t89.4\t88.1\t89.4\t68.1\t \t c(Å)\t212.4\t219.6\t211.7\t95.6\t \t β(°)\t90\t90\t90\t90\t \t γ(°)\t120\t120\t120\t120\t \t Vm (Å3/Da)\t2.57\t2.64\t2.57\t2.64\t \t Solvent Content (%)\t52\t53\t52\t53\t \tX-Ray Data2\t \t \t \t \t \t Resolution (Å)\t30-2.7 (2.8-2.7)\t30-2.3 (2.4-2.3)\t30-2.2 (2.3-2.0)\t30-2.5 (2.6-2.5)\t \t Measured Reflections\t297,367 (19,369)\t333,739 (8,008)\t381,125 (1,591)\t137,992 (9,883)\t \t Unique Reflections\t14,402 (1,003)\t21,683 (1,135)\t24,323 (964)\t16,727 (1,227)\t \t Completeness (%)\t99.6 (96.8)\t93.8 (68.4)\t95.3 (52.0)\t98.6 (98.1)\t \t Redundancy\t20.6 (19.3)\t15.4 (7.1)\t15.7 (1.7)\t8.2 (8.1)\t \t Rmerge\t0.095 (0.451)\t0.057 (0.324)\t0.062 (0.406)\t0.047 (0.445)\t \t \u003c I/σ \u003e\t38.3 (8.1)\t36.7 (5.5)\t36.2 (1.6)\t31.6 (5.6)\t \t B-factor (Å2)\t33.2\t37.3\t33.7\t54.8\t \tRefinement\t \t \t \t \t \t Resolution (Å)\t15-2.7\t15-2.3\t15-2.2\t15-2.5\t \t Number of Reflections\t13,583\t20,255\t24,962\t15,811\t \t Number of All Atoms\t3,335\t3,271\t3,298\t3,239\t \t Number of Waters\t88\t70\t71\t21\t \t R-factor (%)\t19.1\t29.8\t22.8\t25.0\t \t R-free (%)\t26.4\t38.3\t26.6\t33.7\t \tRMSD\t \t \t \t \t \t Bond Lengths (Å)\t0.008\t0.005\t0.005\t0.006\t \t Bond Angles (°)\t1.2\t1.0\t1.0\t1.1\t \t Mean B-factor (Å2)\t49.1\t46.3\t51.7\t88.9\t \tRamachandran Plot (%)\t \t \t \t \t \t Outliers\t0.2\t0.2\t0.2\t1.2\t \t Favored\t96.7\t97.1\t96.5\t90.9\t \t Abbreviations: Amm, ammonium; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. (Continued) Crystal data, X-ray data, and refinement statistics. Fab\tH5-51:L1-39\tH5-51:L3-11\tH5-51:L3-20\tH5-51:L4-1\t \t PDB identifier\t4KMT\t5I1J\t5I1K\t5I1L\t \tCrystal Data\t \t \t \t \t \tCrystallization Solution\t \t \t \t \t \t Buffer, pH\t0.1 M CHES, pH 9.5\t0.1 M Tris, pH 8.5\t0.1 M CHES, pH 9.5\t0.1 M Tris, pH 8.5\t \t Precipitant1\t1.8 M Amm Sulfate\t25% PEG 3350\t1.0 M Amm Sulfate\t24% PEG 3350\t \t Additive1\t5% dioxane\t0.2 M MgCl2\t \t0.2 M Amm Sulfate\t \t Space Group\tP212121\tP212121\tP212121\tP21\t \t Molecules/AU\t1\t1\t1\t2\t \tUnit Cell\t \t \t \t \t \t a(Å)\t63.7\t64.1\t63.8\t106.0\t \t b(Å)\t73.8\t73.8\t74.1\t38.0\t \t c(Å)\t103.1\t103.0\t103.0\t112.3\t \t β(°)\t90\t90\t90\t100.4\t \t Vm (Å3/Da)\t2.53\t2.56\t2.54\t2.28\t \t Solvent Content (%)\t51\t52\t51\t46\t \tX-Ray Data2\t \t \t \t \t \t Resolution (Å)\t30-2.1 (2.2-2.1)\t30-2.5 (2.6-2.5)\t30-1.65 (1.7-1.65)\t30-1.95 (2.0-1.95)\t \t Measured Reflections\t131,839 (6,655)\t120,521 (7,988)\t246,750 (4,142)\t320,324 (12,119)\t \t Unique Reflections\t27,026 (1,885)\t17,286 (1,236)\t53,058 (2,141)\t61,554 (3,243)\t \t Completeness (%)\t93.6 (89.8)\t99.7 (97.3)\t89.8 (49.8)\t94.4 (67.1)\t \t Redundancy\t4.9 (3.5)\t7.0 (6.5)\t4.7 (1.9)\t5.2 (3.7)\t \t Rmerge\t0.079 (0.278)\t0.080 (0.281)\t0.034 (0.131)\t0.060 (0.395)\t \t \u003c I/σ \u003e\t16.8 (5.7)\t21.1(6.9)\t27.5 (5.8)\t19.7 (3.1)\t \t B-factor (Å2)\t26.0\t27.0\t21.6\t31.4\t \tRefinement\t \t \t \t \t \t Resolution (Å)\t15-2.1\t15-2.5\t15-1.65\t15-1.95\t \t Number of Reflections\t25,857\t16,328\t51,882\t60,181\t \t Number of All Atoms\t3,676\t3,454\t3,814\t7,175\t \t Number of Waters\t302\t196\t527\t445\t \t R-factor (%)\t17.1\t17.7\t17.2\t19.4\t \t R-free (%)\t22.0\t25.8\t19.7\t25.8\t \tRMSD\t \t \t \t \t \t Bond Lengths (Å)\t0.006\t0.009\t0.005\t0.009\t \t Bond Angles (°)\t1.0\t1.3\t1.3\t1.3\t \t Mean B-factor (Å2)\t25.2\t38.2\t20.0\t19.5\t \tRamachandran Plot (%)\t \t \t \t \t \t Outliers\t0.0\t0.0\t0.0\t0.0\t \t Favored\t98.4\t97.9\t98.1\t98.0\t \t Abbreviations: Amm, ammonium; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. The crystal structures of a germline library composed of 16 Fabs generated by combining 4 HCs (H1-69, H3-23, H3-53 and H5-51) and 4 LCs (L1-39, L3-11, L3-20 and L4-1) have been determined. The Fab heavy and light chain sequences for the variants numbered according to Chothia are shown in Fig. S1. The four different HCs all have the same CDR H3 sequence, ARYDGIYGELDF. Crystallization of the 16 Fabs was previously reported. Three sets of the crystals were isomorphous with nearly identical unit cells (Table 1). These include (1) H3-23:L3-11 and H3-23:L4-1 in P212121, (2) H3-53:L1-39, H3-53:L3-11 and H3-53:L3-20 in P6522, and (3) H5-51:L1-39, H5-51:L3-11 and H5-51:L3-20 in P212121. Crystallization conditions for the 3 groups are also similar, but not identical (Table 1). Variations occur in the pH (buffer) and the additives, and, in group 3, PEG 3350 is the precipitant for one variants while ammonium sulfate is the precipitant for the other two. The similarity in the crystal forms is attributed in part to cross-seeding using the microseed matrix screening for groups 2 and 3. The crystal structures of the 16 Fabs have been determined at resolutions ranging from 3.3 Å to 1.65 Å (Table 1). The number of Fab molecules in the crystallographic asymmetric unit varies from 1 (for 12 Fabs) to 2 (for 4 Fabs). Overall the structures are fairly complete, and, as can be expected, the models for the higher resolution structures are more complete than those for the lower resolution structures (Table S1). Invariably, the HCs have more disorder than the LCs. For the LC, the disorder is observed at 2 of the C-terminal residues with few exceptions. Apart from the C-terminus, only a few surface residues in LC are disordered. The HCs feature the largest number of disordered residues, with the lower resolution structures having the most. The C-terminal residues including the 6xHis tags are disordered in all 16 structures. In addition to these, 2 primary disordered stretches of residues are observed in a number of structures (Table S1). One involves the loop connecting the first 2 β-strands of the constant domain (in all Fabs except H3-23:L1-39, H3-23:L3-11 and H3-53:L1-39). The other is located in CDR H3 (in H5-51:L3-11, H5-51:L3-20 and in one of 2 copies of H3-23:L4-1). CDR H1 and CDR H2 also show some degree of disorder, but to a lesser extent. CDR canonical structures Several CDR definitions have evolved over decades of antibody research. Depending on the focus of the study, the CDR boundaries differ slightly between various definitions. In this work, we use the CDR definition of North et al., which is similar to that of Martin with the following exceptions: 1) CDRs H1 and H3 begin immediately after the Cys; and 2) CDR L2 includes an additional residue at the N-terminal side, typically Tyr. CDR H1 The superposition of CDR H1 backbones for all HC:LC pairs with heavy chains: (A) H1-69, (B) H3-23, (C) H3-53 and (D) H5-51. Canonical structures. Pairs\tPDB\tCDR H1\tCDR H2\tCDR H3\t \tH1-69\t \tKASGGTFSSYAIS\tGIIPIFGTAN\tARYDGIYGELDF\t \tH1-69:L1-39\t5I15\tH1-13-4\tH2-10-1\tH3-12-1\t \tH1-69:L3-11\t5I16\tH1-13-1/H1-13-1\tH2-10-1/H2-10-1\tH3-12-1/H3-12-1\t \tH1-69:L3-20\t5I17\tH1-13-3/H1-13-6\tH2-10-1/NA\tH3-12-1/H3-12-1\t \tH1-69:L4-1\t5I18\tH1-13-10\tH2-10-1\tH3-12-1\t \tH3-23\t \tAASGFTFSSYAMS\tAISGSGGSTY\tAKYDGIYDGIYGELDF\t \tH3-23:L1-39\t5I19\tH1-13-1\tH2-10-2\tH3-12-1\t \tH3-23:L3-11\t5I1A\tH1-13-1/H1-13-1\tH2-10-2/H2-10-2\tH3-12-1/H3-12-1\t \tH3-23:L3-20\t5I1C\tH1-13-1\tH2-10-2\tH3-12-1\t \tH3-23:L4-1\t5I1D\tH1-13-1/H1-13-1\tH2-10-2/H2-10-2\tH3-12-1/NA\t \tH3-53\t \tAASGFTVSSNYMS\tVIYSGGSTY\tARYDGIYGELDF\t \tH3-53:L1-39\t5I1E\tH1-13-1\tH2-9-3\tH3-12-1\t \tH3-53:L3-11\t5I1G\tH1-13-1\tH2-9-3\tH3-12-1\t \tH3-53:L3-20\t5I1H\tH1-13-1\tH2-9-3\tH3-12-1\t \tH3-53:L4-1\t5I1I\tH1-13-1\tH2-9-3\tNA\t \tH5-51\t \tKGSGYSFTSYWIG\tIIYPGDSDTR\tARYDGIYGELDF\t \tH5-51:L1-39\t4KMT\tH1-13-1\tH2-10-1\tH3-12-1\t \tH5-51:L3-11\t5I1J\tH1-13-1\tH2-10-1\tNA\t \tH5-51:L3-20\t5I1K\tH1-13-1\tH2-10-1\tNA\t \tH5-51:L4-1\t5I1L\tH1-13-1/H1-13-1\tH2-10-1/H2-10-1\tH3-12-1/H3-12-1\t \t \t \tCDR L1\tCDR L2\tCDR L3\t \tL1-39\t \tRASQSISSYLN\tYAASSLQS\tQQSYSTPLT\t \tH1-69:L1-39\t5I15\tL1-11-1\tL2-8-1\tL3-9-cis7-1\t \tH3-23:L1-39\t5I19\tL1-11-1\tL2-8-1\tL3-9-cis7-1\t \tH3-53:L1-39\t5I1E\tL1-11-1\tL2-8-1\tL3-9-cis7-1\t \tH5-51:L1-39\t4KMT\tL1-11-1\tL2-8-1\tL3-9-cis7-1\t \tL3-11\t \tRASQSVSSYLA\tYDASNRAT\tQQRSNWPLT\t \tH1-69:L3-11\t5I16\tL1-11-1/L1-11-1\tL2-8-1/L2-8-1\tL3-9-cis7-1/L3-9-cis7-1\t \tH3-23:L3-11\t5I1A\tL1-11-1/L1-11-1\tL2-8-1/L2-8-1\tL3-9-cis7-1/L3-9-cis7-1\t \tH3-53:L3-11\t5I1G\tL1-11-1\tL2-8-1\tL3-9-cis7-1\t \tH5-51:L3-11\t5I1J\tL1-11-1\tL2-8-1\tL3-9-cis7-1\t \tL3-20\t \tRASQSVSSSYLA\tYGASSRAT\tQQYGSSPLT\t \tH1-69:L3-20\t5I17\tL1-12-2/L1-12-1\tL2-8-1/L2-8-1\tL3-9-cis7-1/L3-9-cis7-1\t \tH3-23:L3-20\t5I1C\tL1-12-2\tL2-8-1\tL3-9-cis7-1\t \tH3-53:L3-20\t5I1H\tL1-12-1\tL2-8-1\tL3-9-cis7-1\t \tH5-51:L3-20\t5I1K\tL1-12-1\tL2-8-1\tL3-9-cis7-1\t \tL4-1\t \tKSSQSVLYSSNNKNYLA\tYWASTRES\tQQYYSTPLT\t \tH1-69:L4-1\t5I18\tL1-17-1\tL2-8-1\tL3-9-cis7-1\t \tH3-23:L4-1\t5I1D\tL1-17-1/L1-17-1\tL2-8-1/L2-8-1\tL3-9-cis7-1/L3-9-cis7-1\t \tH3-53:L4-1\t5I1I\tL1-17-1\tL2-8-1\tL3-9-cis7-1\t \tH5-51:L4-1\t5I1L\tL1-17-1/L1-17-1\tL2-8-1/L2-8-1\tL3-9-cis7-1/L3-9-cis7-1\t \t CDRs are defined using the Dunbrack convention [12]. Assignments for 2 copies of the Fab in the asymmetric unit are given for 5 structures. No assignment (NA) for CDRs with missing residues. The four HCs feature CDR H1 of the same length, and their sequences are highly similar (Table 2). The CDR H1 backbone conformations for all variants for each of the HCs are shown in Fig. 1. Three of the HCs, H3-23, H3-53 and H5-51, have the same canonical structure, H1-13-1, and the backbone conformations are tightly clustered for each set of Fab structures as reflected in the rmsd values (Fig. 1B-D). Some deviation is observed for H3-53, mostly due to H3-53:L4-1, which exhibits a significant degree of disorder in CDR H1. The electron density for the backbone is weak and discontinuous, and completely missing for several side chains.    The CDR H1 structures with H1-69 shown in Fig. 1A are quite variable, both for the structures with different LCs and for the copies of the same Fab in the asymmetric unit, H1-69:L3-11 and H1-69:L3-20. In total, 6 independent Fab structures produce 5 different canonical structures, namely H1-13-1, H1-13-3, H1-13-4, H1-13-6 and H1-13-10. A major difference of H1-69 from the other germlines in the experimental data set is the presence of Gly instead of Phe or Tyr at position 27 (residue 5 of 13 in CDR H1). Glycine introduces the possibility of a higher degree of conformational flexibility that undoubtedly translates to the differences observed, and contributes to the elevated thermal parameters for the atoms in the amino acid residues in this region. CDR H2 The superposition of CDR H2 backbones for all HC:LC pairs with heavy chains: (A) H1-69, (B) H3-23, (C) H3-53 and (D) H5-51. The canonical structures of CDR H2 have fairly consistent conformations (Table 2, Fig. 2). Each of the 4 HCs adopts only one canonical structure regardless of the pairing LC. Germlines H1-69 and H5-51 have the same canonical structure assignment H2-10-1, H3-23 has H2-10-2, and H3-53 has H2-9-3. The conformations for all of these CDR H2s are tightly clustered (Fig. 2). In one case, in the second Fab of H1-69:L3-20, CDR H2 is partially disordered (Δ55-60).   Although three of the germlines have CDR H2 of the same length, 10 residues, they adopt 2 distinctively different conformations depending mostly on the residue at position 71 from the so-called CDR H4. Arg71 in H3-23 fills the space between CDRs H2 and H4, and defines the conformation of the tip of CDR H2 so that residue 54 points away from the antigen binding site. Germlines H1-69 and H5-51 are unique in the human repertoire in having an Ala at position 71 that leaves enough space for H-Pro52a to pack deeper against CDR H4 so that the following residues 53 and 54 point toward the putative antigen. Conformations of CDR H2 in H1-69 and H5-51, both of which have canonical structure H2-10-1, show little deviation within each set of 4 structures. However, there is a significant shift of the CDR as a rigid body when the 2 sets are superimposed. Most likely this is the result of interaction of CDR H2 with CDR H1, namely with the residue at position 33 (residue 11 of 13 in CDR H1). Germline H1-69 has Ala at position 33 whereas in H5-51 position 33 is occupied by a bulky Trp, which stacks against H-Tyr52 and drives CDR H2 away from the center. CDR L1 The superposition of CDR L1 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. The four LC CDRs L1 feature 3 different lengths (11, 12 and 17 residues) having a total of 4 different canonical structure assignments. Of these LCs, L1-39 and L3-11 have the same canonical structure, L1-11-1, and superimpose very well (Fig. 3A, B). For the remaining 2, L3-20 has 2 different assignments, L1-12-1 and L1-12-2, while L4-1 has a single assignment, L1-17-1.   L4-1 has the longest CDR L1, composed of 17 amino acid residues (Fig. 3D). Despite this, the conformations are tightly clustered (rmsd is 0.20 Å). The backbone conformations of the stem regions superimpose well. Some changes in conformation occur between residues 30a and 30f (residues 8 and 13 of 17 in CDR L1). This is the tip of the loop region, which appears to have similar conformations that fan out the structures because of the slight differences in torsion angles in the backbone near Tyr30a and Lys30f. L3-20 is the most variable in CDR L1 among the 4 germlines as indicated by an rmsd of 0.54 Å (Fig. 3C). Two structures, H3-53:L3-20 and H5-51:L3-20 are assigned to canonical structure L1-12-1 with virtually identical backbone conformations. The third structure, H3-23:L3-20, has CDR L1 as L1-12-2, which deviates from L1-12-1 at residues 29-32, i.e., at the site of insertion with respect to the 11-residue CDR. The fourth member of the set, H1-69:L3-20, was crystallized with 2 Fabs in the asymmetric unit. The conformation of CDR L1 in these 2 Fabs is slightly different, and both conformations fall somewhere between L1-12-1 and L1-12-2. This reflects the lack of accuracy in the structure due to low resolution of the X-ray data (3.3 Å). CDR L2 The superposition of CDR L2 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. All four LCs have CDR L2 of the same length and canonical structure, L2-8-1 (Table 2). The CDR L2 conformations for each of the LCs paired with the 4 HCs are clustered more tightly than any of the other CDRs (rmsd values are in the range 0.09-0.16 Å), and all 4 sets have virtually the same conformation despite the sequence diversity of the loop. No significant conformation outliers are observed (Fig. 4).   CDR L3 The superposition of CDR L3 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. As with CDR L2, all 4 LCs have CDR L3 of the same length and canonical structure, L3-9-cis7-1 (Table 2). The conformations of CDR L3 for L1-39, L3-11, and particularly for L320, are not as tightly clustered as those of L4-1 (Fig. 5). The slight conformational variability occurs in the region of amino acid residues 90-92, which is in contact with CDR H3.   CDR H3 conformational diversity As mentioned earlier, all 16 Fabs have the same CDR H3, for which the amino acid sequence is derived from the anti-CCL2 antibody CNTO 888. The loop and the 2 β-strands of the CDR H3 in this ‘parent’ structure are stabilized by H-bonds between the carbonyl oxygen and peptide nitrogen atoms in the 2 strands. An interesting feature of these CDR H3 structures is the presence of a water molecule that interacts with the peptide nitrogens and carbonyl oxygens near the bridging loop connecting the 2 β-strands. This water is present in both the bound (4DN4) and unbound (4DN3) forms of CNTO 888. The stem region of CDR H3 in the parental Fab is in a ‘kinked’ conformation, in which the indole nitrogen of Trp103 forms a hydrogen bond with the carbonyl oxygen of Leu100b. The carboxyl group of Asp101 forms a salt bridge with Arg94. These interactions are illustrated in Fig. S2. Ribbon representations of (A) the superposition of all CDR H3s of the structures with complete backbone traces. (B) The CDR H3s rotated 90° about the y axis of the page. The structure of each CDR H3 is represented with a different color. Despite having the same amino acid sequence in all variants, CDR H3 has the highest degree of structural diversity and disorder of all of the CDRs in the experimental set. Three of the 21 Fab structures (including multiple copies in the asymmetric unit), H5-51:L3-11, H551:L3-20 and H3-23:L4-1 (one of the 2 Fabs), have missing (disordered) residues at the apex of the CDR loop. Another four of the Fabs, H3-23:L1-39, H3-53:L1-39, H3-53:L3-11 and H3-53:L4-1 have missing side-chain atoms. The variations in CDR H3 conformation are illustrated in Fig. 6 for the 18 Fab structures that have ordered backbone atoms.   A comparison of representatives of the “kinked” and “extended” structures. (A) The “kinked” CDR H3 of H1-69:L3-11 with purple carbon atoms and yellow dashed lines connecting the H-bond pairs for Leu100b O and Trp103 NE1, Arg94 NE and Asp101 OD1, and Arg94 NH2 and Asp101 OD2. (B) The “extended” CDR H3 of H1-69:L3-20 with green carbon atoms and yellow dashed lines connecting the H-bond pairs for Asp101 OD1 and OD2 and Trp103 NE1. In 10 of the 18 Fab structures, H1-69:L1-39, H1-69:L3-11 (2 Fabs), H1-69:L4-1, H3-23:L3-11 (2 Fabs), H3-23:L3-20, H3-53:L3-11, H3-53:L3-20 and H5-51:L1-39, the CDRs have similar conformations to that found in 4DN3. The bases of these structures have the ‘kinked’ conformation with the H-bond between Trp103 and Leu100b. A representative CDR H3 structure for H1-69:L1-39 illustrating this is shown in Fig. 7A. The largest backbone conformational deviation for the set is at Tyr99, where the C=O is rotated by 90° relative to that observed in 4DN3. Also, it is worth noting that only one of these structures, H1-69:L4-1, has the conserved water molecule in CDR H3 observed in the 4DN3 and 4DN4 structures. In fact, it is the only Fab in the set that has a water molecule present at this site. The CDR H3 for this structure is shown in Fig. S3.   The remaining 8 Fabs can be grouped into 5 different conformational classes. Three of the Fabs, H3-23:L1-39, H3-23:L4-1 and H3-53:L1-39, have distinctive conformations. The stem regions in these 3 cases are in the ‘kinked’ conformation consistent with that observed for 4DN3. The five remaining Fabs, H5-51:L4-1 (2 copies), H1-69:L3-20 (2 copies) and H3-53:L4-1, have 3 different CDR H3 conformations (Fig. S4). The stem regions of CDR H3 for the H5-51:L4-1 Fabs are in the ‘kinked’ conformation while, surprisingly, those of the H1-69:L3-20 pair and H3-53:L4-1 are in the ‘extended’ conformation (Fig. 7B). VH:VL domain packing The VH and VL domains have a β-sandwich structure (also often referred as a Greek key motif) and each is composed of a 4-stranded and a 5-stranded antiparallel β-sheets. The two domains pack together such that the 5-stranded β-sheets, which have hydrophobic surfaces, interact with each other bringing the CDRs from both the VH and VL domains into close proximity. The domain packing of the variants was assessed by computing the domain interface interactions, the VH:VL tilt angles, the buried surface area and surface complementarity. The results of these analyses are shown in Tables 3, 4 and S2. VH:VL interface amino acid residue interactions The conserved VH:VL interactions as viewed along the VH/VL axis. The VH residues are in blue, the VL residues are in orange. The VH:VL interface is pseudosymmetric, and involves 2 stretches of the polypeptide chain from each domain, namely CDR3 and the framework region between CDRs 1 and 2. These stretches form antiparallel β-hairpins within the internal 5-stranded β-sheet. There are a few principal inter-domain interactions that are conserved not only in the experimental set of 16 Fabs, but in all human antibodies. They include: 1) a bidentate hydrogen bond between L-Gln38 and H-Gln39; 2) H-Leu45 in a hydrophobic pocket between L-Phe98, L-Tyr87 and L-Pro44; 3) L-Pro44 stacked against H-Trp103; and 4) L-Ala43 opposite the face of H-Tyr91 (Fig. 8). With the exception of L-Ala43, all other residues are conserved in human germlines. Position 43 may be alternatively occupied by Ser, Val or Pro (as in L4-1), but the hydrophobic interaction with H-Tyr91 is preserved. These core interactions provide enough stability to the VH:VL dimer so that additional VH-VL contacts can tolerate amino acid sequence variations in CDRs H3 and L3 that form part of the VH:VL interface.   In total, about 20 residues are involved in the VH:VL interactions on each side (Fig. S5). Half of them are in the framework regions and those residues (except residue 61 in HC, which is actually in CDR2 in Kabat's definition) are conserved in the set of 16 Fabs. The side chain conformations of these conserved residues are also highly similar. One notable exception is H-Trp47, which exhibits 2 conformations of the indole ring. In most of the structures, it has the χ2 angle of ∼80°, while the ring is flipped over (χ2 = −100°) in H5-51:L3:11 and H5-51:L3-20. Interestingly, these are the only 2 structures with residues missing in CDR H3 because of disorder, although both structures are determined at high resolution and the rest of the structure is well defined. Apparently, residues flanking CDR H3 in the 2 VH:VL pairings are inconsistent with any stable conformation of CDR H3, which translates into a less restricted conformational space for some of them, including H-Trp47. VH:VL tilt angles The relative orientation of VH and VL has been measured in a number of different ways. Presented here are the results of 2 different approaches for determining the orientation of one domain relative to the other. The first approach uses ABangles, the results of which are shown in Table S2. The four LCs all are classified as Type A because they have a proline at position 44, and the results for each orientation parameter are within the range of values of this type reported by Dunbar and co-workers. In fact, the parameter values for the set of 16 Fabs are in the middle of the distribution observed for 351 non-redundant antibody structures determined at 3.0 Å resolution or better. The only exception is HC1, which is shifted toward smaller angles with the mean value of 70.8° as compared to the distribution centered at 72° for the entire PDB. This probably reflects the invariance of CDR H3 in the current set as opposed to the CDR H3 diversity in the PDB. The second approach used for comparing tilt angles involved computing the difference in the tilt angles between all pairs of structures. For structures with 2 copies of the Fab in the asymmetric unit, only one structure was used. The differences between independent Fabs in the same structure are 4.9° for H1-69:L3-20, 1.6° for H1-69:L3-11, 1.4° for H3-23:L4-1, 3.3° for H3-23:L3-11, and 2.5° for H5-51:L4-1. With the exception of H1-69:L3-20, the angles are within the range of 2-3° as are observed in the identical structures in the PDB. In H1-69:L3-20, one of the Fabs is substantially disordered so that part of CDR H2 (the outer β-strand, residues 55-60) is completely missing. This kind of disorder may compromise the integrity of the VH domain and its interaction with the VL. Indeed, this Fab has the largest twist angle HC2 within the experimental set that exceeds the mean value by 2.5 standard deviations (Table S2). An illustration of the difference in tilt angle for 2 pairs of variants by the superposition of the VH domains of (A) H1-69:L3-20 on that of H5-51:L1-39 (the VL domain is off by a rigid-body roatation of 10.5°) and (B) H1-69:L4-1 on that of H5-51:L1-39 (the VL domain is off by a rigid-body roatation of 1.6°). Differences in VH:VL tilt angles.  \tH1-69:L1-39\tH1-69:L3-11\tH1-69:L3-20\tH1-69:L4-1\tH3-23:L1-39\tH3-23:L3-11\tH3-23:L3-20\tH3-23:L4-1\tH3-53:L1-39\tH3-53:L3-11\tH3-53:L3-20\tH3-53:L4-1\tH5-51:L1-39\tH5-51:L3-11\tH5-51:L3-20\tH5-51:L4-1\t \tH1-69:L1-39\t0\t2.1\t8.9\t1.1\t4.2\t3.0\t9.5\t1.5\t3.3\t3.6\t3.1\t1.6\t1.8\t2.9\t2.4\t5.2\t \tH1-69:L3-11\t \t0\t7.3\t2.9\t2.5\t2.0\t8.4\t1.3\t2.6\t2.9\t3.2\t1.8\t3.9\t4.6\t4.4\t5.0\t \tH1-69:L3-20\t \t \t0\t9.2\t5.0\t8.7\t7.4\t7.6\t8.9\t8.6\t9.4\t7.9\t10.5\t10.1\t11.0\t9.7\t \tH1-69:L4-1\t \t \t \t0\t4.6\t3.9\t10.1\t1.8\t4.4\t4.7\t4.1\t2.3\t1.6\t2.5\t2.3\t6.2\t \tH3-23:L1-39\t \t \t \t \t0\t4.0\t8.0\t2.8\t4.8\t4.8\t5.4\t3.6\t6.0\t6.2\t6.6\t6.6\t \tH3-23:L3-11\t \t \t \t \t \t0\t9.3\t3.0\t2.1\t2.9\t3.3\t3.3\t4.6\t5.8\t5.0\t5.2\t \tH3-23:L3-20\t \t \t \t \t \t \t0\t8.9\t7.9\t7.0\t7.6\t7.9\t10.5\t9.7\t10.7\t6.2\t \tH3-23:L4-1\t \t \t \t \t \t \t \t0\t3.6\t3.8\t3.7\t1.5\t3.2\t3.7\t3.8\t5.6\t \tH3-53:L1-39\t \t \t \t \t \t \t \t \t0\t1.0\t1.6\t2.9\t4.6\t5.3\t4.8\t3.1\t \tH3-53:L3-11\t \t \t \t \t \t \t \t \t \t0\t1.3\t2.9\t4.8\t5.2\t5.0\t2.3\t \tH3-53:L3-20\t \t \t \t \t \t \t \t \t \t \t0\t2.5\t3.8\t4.2\t3.9\t2.2\t \tH3-53:L4-1\t \t \t \t \t \t \t \t \t \t \t \t0\t2.9\t3.0\t3.3\t4.2\t \tH5-51:L1-39\t \t \t \t \t \t \t \t \t \t \t \t \t0\t1.9\t0.6\t5.8\t \tH5-51:L3-11\t \t \t \t \t \t \t \t \t \t \t \t \t \t0\t1.9\t5.7\t \tH5-51:L3-20\t \t \t \t \t \t \t \t \t \t \t \t \t \t \t0\t5.8\t \tH5-51:L4-1\t \t \t \t \t \t \t \t \t \t \t \t \t \t \t \t0\t \t The differences in the tilt angle are shown for all pairs of V regions in Table 3. They range from 0.6° to 11.0°. The smallest differences in the tilt angle are between the Fabs in isomorphous crystal forms. The largest deviations in the tilt angle, up to 11.0°, are found for 2 structures, H1-69:L3-20 and H3-23:L3-20, that stand out from the other Fabs. One of the 2 structures, H1-69:L3-20, has its CDR H3 in the ‘extended’ conformation; the other structure has it in the ‘kinked’ conformation. Two examples illustrating large (10.5°) and small (1.6°) differences in the tilt angles are shown in Fig. 9.    VH:VL buried surface area and complementarity VH:VL surface areas and surface complementarity. Chain Pairs\tPDB\tContact surfaceVH (Å2)\tContact surfaceVL (Å2)\tInterface(Å2)\tSurface complementarity\t \tH1-69:L1-39\t5I15\t727\t771\t749\t0.743\t \tH1-69:L3-11\t5I16\t802\t870\t836\t0.762\t \tH1-69:L3-20\t5I17\t713\t736\t725\t0.723\t \tH1-69:L4-1\t5I18\t729\t736\t733\t0.734\t \tH3-23:L1-39\t5I19\t795\t817\t806\t0.722\t \tH3-23:L3-11\t5I1A\t822\t834\t828\t0.725\t \tH3-23:L3-20\t5I1C\t670\t698\t684\t0.676\t \tH3-23:L4-1\t5I1D\t743\t770\t757\t0.708\t \tH3-53:L1-39\t5I1E\t698\t719\t709\t0.712\t \tH3-53:L3-11\t5I1G\t747\t758\t753\t0.690\t \tH3-53:L3-20\t5I1H\t743\t735\t739\t0.687\t \tH3-53:L4-1\t5I1I\t689\t693\t691\t0.711\t \tH5-51:L1-39\t4KMT\t761\t808\t785\t0.728\t \tH5-51:L3-11\t5I1J\t648\t714\t681\t0.717\t \tH5-51:L3-20\t5I1K\t622\t643\t633\t0.740\t \tH5-51:L4-1\t5I1L\t790\t792\t791\t0.704\t \t Some side chain atoms in CDR H3 are missing. Residues in CDR H3 are missing: YGE in H5-51:L3-11, GIY in H5-51:L3-20. The results of the PISA contact surface calculation and surface complementarity calculation are shown in Table 4. The interface areas are calculated as the average of the VH and VL contact surfaces. Six of the 16 structures have CDR H3 side chains or complete residues missing, and therefore their interfaces are much smaller than in the other 10 structures with complete CDRs (the results are provided for all Fabs for completeness). Among the complete structures, the interface areas range from 684 to 836 Å2. Interestingly, the 2 structures that have the largest tilt angle differences with the other variants, H3-23:L3-20 and H1-69:L3-20, have the smallest VH:VL interfaces, 684 and 725 Å2, respectively. H3-23:L3-20 is also unique in that it has the lowest value (0.676) of surface complementarity.   Stability of germline pairings Melting temperatures for the 16 Fabs.  \tL3-20\tL4-1\tL3-11\tL1-39\tHC average\t \tH1-69\t73.6\t74.8\t75.6\t80.3\t76.1\t \tH3-23\t74.8\t75.2\t4.8\t81.5\t76.6\t \tH3-53\t68.4\t68.0\t71.5\t73.9\t70.5\t \tH5-51\t68.4\t68.4\t71.9\t77.0\t71.4\t \tLC average\t71.3\t71.6\t73.5\t78.2\t \t \t Colors: blue (Tm \u003c 70°C), green (70°C \u003c Tm \u003c 73°C), yellow (73°C \u003c Tm \u003c 78°C), orange (Tm \u003e 78°C). Melting temperatures (Tm) were measured for all Fabs using differential scanning calorimetry (Table 5). It appears that for each given LC, the Fabs with germlines H1-69 and H3-23 are substantially more stable than those with germlines H3-53 and H5-51. In addition, L1-39 provides a much higher degree of stabilization than the other 3 LC germlines when combined with any of the HCs. As a result, the Tm for pairs H1-69:L1-39 and H3-23:L1-39 is 12-13° higher than for pairs H3-53:L3-20, H3-53:L4-1, H5-51:L3-20 and H5-51:L4-1.   These findings correlate well with the degree of conformational disorder observed in the crystal structures. Parts of CDR H3 main chain are completely disordered, and were not modeled in Fabs H5-51:L3-20 and H5-51:L3-11 that have the lowest Tms in the set. No electron density is observed for a number of side chains in CDRs H3 and L3 in all Fabs with germline H3-53, which indicates loose packing of the variable domains. All those molecules are relatively unstable, as is reflected in their low Tms. Discussion This is the first report of a systematic structural investigation of a phage germline library. The 16 Fab structures offer a unique look at all pairings of 4 different HCs (H1-69, H3-23, H3-53, and H5-51) and 4 different LCs (L1-39, L3-11, L3-20 and L4-1), all with the same CDR H3. The structural data set taken as a whole provides insight into how the backbone conformations of the CDRs of a specific heavy or light chain vary when it is paired with 4 different light or heavy chains, respectively. A large variability in the CDR conformations for the sets of HCs and LCs is observed. In some cases the CDR conformations for all members of a set are virtually identical, for others subtle changes occur in a few members of a set, and in some cases larger deviations are observed within a set. The five variants that crystallized with 2 copies of the Fab in the asymmetric unit serve somewhat as controls for the influence of crystal packing on the conformations of the CDRs. In four of the 5 structures the CDR conformations are consistent. In only one case, that of H1-69:L3-20 (the lowest resolution structure), do we see differences in the conformations of the 2 copies of CDRs H1 and L1. This variability is likely a result of 2 factors, crystal packing interactions and internal instability of the variable domain. For the CDRs with canonical structures, the largest changes in conformation occur for CDR H1 of H1-69 and H3-53. The other 2 HCs, H3-23 and H5-51, have canonical structures that are remarkably well conserved (Fig. 1). Of the 4 HCs, H1-69 has the greatest number of canonical structure assignments (Table 2). H1-69 is unique in having a pair of glycine residues at positions 26 and 27, which provide more conformational freedom in CDR H1. Besides IGHV1-69, only the germlines of the VH4 family possess double glycines in CDR H1, and it will be interesting to see if they are also conformationally unstable. Having all 16 VH:VL pairs with the same CDR H3 provides some insights into why molecular modeling efforts of CDR H3 have proven so difficult. As mentioned in the Results section, this data set is composed of 21 Fabs, since 5 of the 16 variants have 2 Fab copies in the asymmetric unit. For the 18 Fabs with complete backbone atoms for CDR H3, 10 have conformations similar to that of the parent, while the others have significantly different conformations (Fig. 6). Thus, it is likely that the CDR H3 conformation is dependent upon 2 dominating factors: 1) amino acid sequence; and 2) VH and VL context. More than half of the variants retain the conformation of the parent despite having differences in the VH:VL pairing. This subset includes 2 structures with 2 copies of the Fab in the asymmetric unit, all of which are nearly identical in conformation. This provides an internal control showing a consistency in the conformations. The remaining 8 structures exhibit “non-parental” conformations, indicating that the VH and VL context can also be a dominating factor influencing CDR H3. Importantly, there are 5 distinctive conformations in this subset. This subset also has 2 structures with 2 Fab copies in the asymmetric unit. Each pair has nearly identical conformations providing an internal check on the consistency of the conformations. Interestingly, as described earlier, these 2 pairs differ in the stem regions with the H1-69:L3-20 pair in the ‘extended’ conformation and H5-51:L4-1 pair in the ‘kinked’ conformation. The conformations are different from each other, as well as from the parent. The CDR H3 conformational analysis shows that, for each set of variants of one HC paired with the 4 different LCs, both “parental” and “non-parental” conformations are observed. The same variability is observed for the sets of variants composed of one LC paired with each of the 4 HCs. Thus, no patterns of conformational preference for a particular HC or LC emerge to shed any direct light on what drives the conformational differences. This finding supports the hypothesis of Weitzner et al. that the H3 conformation is controlled both by its sequence and its environment. In looking at a possible correlation between the tilt angle and the conformation of CDR H3, no clear trends are observed. Two variants, H1-69:L3-20 and H3-23:L3-20, have the largest differences in the tilt angles compared to other variants as seen in Table 3. The absolute VH:VL orientation parameters for the 2 Fabs (Table S2) show significant deviation in HL, LC1 and HC2 values (2-3 standard deviations from the mean). One of the variants, H3-23:L3-20, has the CDR H3 conformation similar to the parent, but the other, H1-69:L3-20, is different. As noted in the Results section, the 2 variants, H1-69:L3-20 and H3-23:L3-20, are outliers in terms of the tilt angle; at the same time, both have the smallest VH:VL interface. These smaller interfaces may perhaps translate to a significant deviation in how VH is oriented relative to VL than the other variants. These deviations from the other variants can also be seen to some extent in VH:VL orientation parameters in Table S2, as well as in the smaller number of residues involved in the VH:VL interfaces of these 2 variants (Fig. S5). These differences undoubtedly influence the conformation of the CDRs, in particular CDR H1 (Fig. 1A) and CDR L1 (Fig. 3C), especially with the tandem glycines and multiple serines present, respectively. Pairing of different germlines yields antibodies with various degrees of stability. As indicated by the melting temperatures, germlines H1-69 and H3-23 for HC and germline L1-39 for LC produce more stable Fabs compared to the other germlines in the experimental set. Structural determinants of the differential stability are not always easy to decipher. One possible explanation of the clear preference of LC germline L1-39 is that CDR L3 has smaller residues at positions 91 and 94, allowing for more room to accommodate CDR H3. Other germlines have bulky residues, Tyr, Arg and Trp, at these positions, whereas L1-39 has Ser and Thr. Various combinations of germline sequences for VL and VH impose certain constraints on CDR H3, which has to adapt to the environment. A more compact CDR L3 may be beneficial in this situation. At the other end of the stability range is LC germline L3-20, which yields antibodies with the lowest Tms. While pairings with H3-53 and H5-51 may be safely called a mismatch, those with H1-69 and H3-23 have Tms about 5-6° higher. Curiously, the 2 Fabs, H1-69:L3-20 and H3-23:L3-20, deviate markedly in their tilt angles from the rest of the panel. It is possible that by adopting extreme tilt angles the structure modulates CDR H3 and its environment, which apparently cannot be achieved solely by conformational rearrangement of the CDR. Note that most of the VH:VL interface residues are invariant; therefore, significant change of the tilt angle must come with a penalty in free energy. Yet, for the 2 antibodies, the total gain in stability merits the domain repacking. Overall, the stability of the Fab, as measured by Tm, is a result of the mutual adjustment of the HC and LC variable domains and adjustment of CDR H3 to the VH:VL cleft. The final conformation represents an energetic minimum; however, in most cases it is very shallow, so that a single mutation can cause a dramatic rearrangement of the structure. In summary, the analysis of this structural library of germline variants composed of all pairs of 4 HCs and 4LCs, all with the same CDR H3, offers some unique insights into antibody structure and how pairing and sequence may influence, or not, the canonical structures of the L1, L2, L3, H1 and H2 CDRs. Comparison of the CDR H3s reveals a large set of variants with conformations similar to the parent, while a second set has significant conformational variability, indicating that both the sequence and the structural context define the CDR H3 conformation. Quite unexpectedly, 2 of the variants, H1-69:L3-20 and H3-53:L4-1, have the ‘extended’ stem region differing from the other 14 that have a ‘kinked’ stem region. Why this is the case is unclear at present. These data reveal the difficulty of modeling CDR H3 accurately, as shown again in Antibody Modeling Assessment II. Furthermore, antibody CDRs, H3 in particular, may go through conformational changes upon binding their targets, making structural prediction for docking purposes an even more difficult task. Fortunately, for most applications of antibody modeling, such as engineering affinity and biophysical properties, an accurate CDR H3 structure is not always necessary. For those applications where accurate CDR structures are essential, such as docking, the results in this work demonstrate the importance of experimental structures. With the recent advances in expression and crystallization methods, Fab structures can be obtained rapidly. The set of 16 germline Fab structures offers a unique dataset to facilitate software development for antibody modeling. The results essentially support the underlying idea of canonical structures, indicating that most CDRs with germline sequences tend to adopt predefined conformations. From this point of view, a novel approach to design combinatorial antibody libraries would be to cover the range of CDR conformations that may not necessarily coincide with the germline usage in the human repertoire. This would insure more structural diversity, leading to a more diverse panel of antibodies that would bind to a broad spectrum of targets. Materials and methods Fab production, purification and crystallization The production, purification and crystallization of the Fabs reported in this article were described previously. Briefly, the 16 Fabs were produced by combining 4 different HC and 4 different LC germline constructs. The human HC germlines were IGHV1-69 (H1-69), IGHV3-23 (H3-23), IGHV3-53 (H3-53) and IGHV5-51 (H5-51) in the IMGT nomenclature. The human LC germlines were IGKV1-39 (L1-39), IGKV3-11 (L3-11), IGKV3-20 (L3-20) and IGKV4-1 (L4-1) corresponding to O12, L6, A27 and B3 in the V-BASE nomenclature. CDR H3 of the anti-CCL2 antibody CNTO 888 with the amino acid sequence ARYDGIYGELDF  was used in all Fab constructs. The J region genes were IGHJ1 for the HC and IGKJ1 for the LC for all Fabs. Human IgG1 and κ constant regions were used in all Fab constructs. A 6xHis tag was added to the C-terminus of the HC to facilitate purification. The Fabs were expressed in HEK 293E cells and purified by affinity and size-exclusion chromatography. For crystallization, the Fabs were dialyzed into 20 mM Tris buffer, pH 7.4, with 50 mM NaCl and concentrated to 12-18 mg/mL. Automated crystallization screening was carried out using the vapor diffusion method at 20°C with an Oryx4 (Douglas Instruments) or a Mosquito (TTP Labtech) crystallization robot in a sitting drop format using Corning 3550 plates. Initial screening was carried out with an in-house 192-well screen optimized for Fab crystallization and the Hampton 96-well Crystal Screen HT (Hampton Research). For the majority of the Fabs, the crystallization protocol employed microseed matrix screening using self-seeding or cross-seeding approaches. A summary of the final crystallization conditions for each of the Fabs is presented in Table 1. X-ray data collection For 13 of the Fab crystals, X-ray data collection was carried out at Janssen Research and Development, LLC using a Rigaku MicroMax™-007HF microfocus X-ray generator equipped with a Saturn 944 CCD detector and an X-stream™ 2000 cryocooling system (Rigaku), and for the remaining 3, X-ray data collection was carried out at the Advanced Photon Source (APS) synchrotron at Argonne National Laboratory using the IMCA 17-ID beamline with a Pilatus 6M detector. For X-ray data collection, the Fab crystals were soaked for a few seconds in a cryo-protectant solution containing the corresponding mother liquor supplemented with 17-25% glycerol (Table S1). The crystals for which data were collected in-house were flash cooled in the stream of nitrogen at 100 K. Crystals sent to the APS were flash cooled in liquid nitrogen prior to shipping them to the synchrotron. Diffraction data for all variants were processed with the program XDS. X-ray data statistics are given in Table 1. Structure determination A summary of the methods used in the structure solution and refinement of the 16 Fabs is presented in Table S1. Twelve of the structures were solved by molecular replacement with Phaser using different combinations of search models for the VH, VL and constant domains. Four of the structures, H3-53:L1-39, H3-53:L3-11, H5-51:L1-39 and H5-51:L3-11, were solved by direct replacement followed by rigid body refinement with REFMAC. All structures were refined using REFMAC. Model adjustments were carried out using the program Coot. The refinement statistics are given in Table 1. Other crystallographic calculations were performed with the CCP4 suite of programs. The structural figures were prepared using the PyMOL Molecular Graphics System, Version 1.0 (Schrödinger, LLC). Structural analysis The canonical structure assignments (Table 2) were made using PyIgClassify, an online canonical structure classification tool (http://dunbrack2.fccc.edu/pyigclassify/) that uses the rules set forth by Dunbrack and coworkers. The conformational variability within the CDRs was assessed by calculating the root-mean-square deviation (rmsd) from the average structure that was generated after superposition of all structures of the set using the main-chain atoms of the CDR in question. The rmsd was calculated for all main-chain atoms (N, CA, C, O) of the CDR. The contact surface areas of the VH and VL domains at the VH:VL inteface were computed with the CCP4 program PISA. The surface complementarity of the VH and VL domains was computed using the CCP4 program SC. VH:VL tilt angles The orientation of the VH domain with respect to the VL domain was assessed using 2 different approaches. The first approach calculates the 6 VH:VL orientation parameters that describe the VH:VL relationship according to Dunbar and co-workers using a script downloaded from the website (http://opig.stats.ox.ac.uk/webapps/abangle). The six parameters include 5 angles, HL, H1, H2, L1 and L2, and a distance, dc. These parameters are derived by first defining 2 planes, one for each domain, based on core residues in the domains. The distance between the planes, dc, is determined along a vector between the planes that is used to establish a consistent coordinate system. The torsion angle between the domains, HL, is much like the VH:VL packing angle defined by Abhinandan and Martin. The tilt of one domain relative to the other is defined by the HC1 and LC1 angles, and the twist of one domain relative to the other is defined by the HC2 and LC2 angles. The second approach calculates the difference in the tilt angle between pairs of Fvs, which reflects the relative orientation between the VH and VL domains. The difference with respect to the reference structure is calculated by sequential root-mean-square superposition of the VL and VH domains using β-sheet core Cα positions (Chothia numbering scheme): 3–13, 18–25, 33–38, 43–49, 61–67, 70–76, 85–90, 97–103 for VL; 3–7, 18–24, 34–40, 44–51, 56–59, 67–72, 77–82a, 87–94, 102–110 for VH. The κ angle in the spherical polar angular system (ω, ϕ, κ) of the latter transformation is the difference in the tilt angle. Differential scanning calorimetry DSC experiments were performed on a VP-capillary DSC system (MicroCal Inc., Northampton, MA) in which temperature differences between the reference and sample cell are continuously measured and calibrated to power units. Samples were heated from 10°C to 95°C at a heating rate of 60°C/hour. The pre-scan time was 15 minutes and the filtering period was 10 seconds. The concentration used in the DSC experiments was about 0.4 mg/mL in phosphate-buffered saline. Analysis of the resulting thermograms was performed using MicroCal Origin 7 software. Melting temperature of proteins was determined by deconvolution of the DSC scans using non-2 state model in the MicroCal Origin 7 software. Scans were deconvoluted using a non-2 state model with either 1-step transition or 2-step transition depending on the number of resolved peaks observed in a scan. Accession numbers Atomic coordinates and structure factors have been deposited in the Protein Data Bank with accession numbers 4KMT, 5I15, 5I16, 5I17, 5I18, 5I19, 5I1A, 5I1C, 5I1D, 5I1E, 5I1G, 5I1H, 5I1I, 5I1J, 5I1K and 5I1L. Supplementary Material Disclosure of potential conflicts of interest No potential conflicts of interest were disclosed. 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Structures and models De novo selection of high-affinity antibodies from synthetic fab libraries displayed on phage as pIX fusion proteins Analysis of heavy and light chain pairings indicates that receptor editing shapes the human antibody repertoire Structural basis for high selectivity of anti-CCL2 neutralizing antibody CNTO 888 Protein crystallization with microseed matrix screening: application to human germline antibody Fabs Framework residue-71 is a major determinant of the position and conformation of the 2nd hypervariable region in the VH domains of immunoglobulins Modeling the anti-CEA antibody combining site by homology and conformational search Major antigen-induced domain rearrangements in an antibody Energy-based analysis and prediction of the orientation between light- and heavy-chain antibody variable domains Analysis and prediction of VH/VL packing in antibodies ABangle: characterising the VH-VL orientation in antibodies Inference of macromolecular assemblies from crystalline state Shape complementarity at protein/protein interfaces Structural evidence for induced fit as a mechanism for antibody-antigen recognition Nomenclature of the human immunoglobulin genes  Two routes for production and purification of Fab fragments in biopharmaceutical discovery research: papain digestion of mAb and transient expression in mammalian cells An automated microseed matrix-screening method for protein crystallization Promoting crystallization of antibody-antigen complexes via microseed matrix screening XDS Phaser crystallographic software REFMAC5 for the refinement of macromolecular crystal structures Features and development of Coot Overview of the CCP4 suite and current 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diff --git a/BioC_XML/4746701_v0.xml b/annotated_BioC_XML/PMC4746701_ann.xml
similarity index 100%
rename from BioC_XML/4746701_v0.xml
rename to annotated_BioC_XML/PMC4746701_ann.xml
diff --git a/BioC_XML/4772114_v0.xml b/annotated_BioC_XML/PMC4772114_ann.xml
similarity index 100%
rename from BioC_XML/4772114_v0.xml
rename to annotated_BioC_XML/PMC4772114_ann.xml
diff --git a/BioC_XML/4773095_v0.xml b/annotated_BioC_XML/PMC4773095_ann.xml
similarity index 100%
rename from BioC_XML/4773095_v0.xml
rename to annotated_BioC_XML/PMC4773095_ann.xml
diff --git a/BioC_XML/4774019_v0.xml b/annotated_BioC_XML/PMC4774019_ann.xml
similarity index 100%
rename from BioC_XML/4774019_v0.xml
rename to annotated_BioC_XML/PMC4774019_ann.xml
diff --git a/BioC_XML/4781976_v0.xml b/annotated_BioC_XML/PMC4781976_ann.xml
similarity index 100%
rename from BioC_XML/4781976_v0.xml
rename to annotated_BioC_XML/PMC4781976_ann.xml
diff --git a/BioC_XML/4784909_v0.xml b/annotated_BioC_XML/PMC4784909_ann.xml
similarity index 100%
rename from BioC_XML/4784909_v0.xml
rename to annotated_BioC_XML/PMC4784909_ann.xml
diff --git a/BioC_XML/4786784_v0.xml b/annotated_BioC_XML/PMC4786784_ann.xml
similarity index 100%
rename from BioC_XML/4786784_v0.xml
rename to annotated_BioC_XML/PMC4786784_ann.xml
diff --git a/BioC_XML/4792962_v0.xml b/annotated_BioC_XML/PMC4792962_ann.xml
similarity index 100%
rename from BioC_XML/4792962_v0.xml
rename to annotated_BioC_XML/PMC4792962_ann.xml
diff --git a/BioC_XML/4795551_v0.xml b/annotated_BioC_XML/PMC4795551_ann.xml
similarity index 100%
rename from BioC_XML/4795551_v0.xml
rename to annotated_BioC_XML/PMC4795551_ann.xml
diff --git a/BioC_XML/4802042_v0.xml b/annotated_BioC_XML/PMC4802042_ann.xml
similarity index 100%
rename from BioC_XML/4802042_v0.xml
rename to annotated_BioC_XML/PMC4802042_ann.xml
diff --git a/BioC_XML/4802085_v0.xml b/annotated_BioC_XML/PMC4802085_ann.xml
similarity index 100%
rename from BioC_XML/4802085_v0.xml
rename to annotated_BioC_XML/PMC4802085_ann.xml
diff --git a/BioC_XML/4820378_v0.xml b/annotated_BioC_XML/PMC4820378_ann.xml
similarity index 100%
rename from BioC_XML/4820378_v0.xml
rename to annotated_BioC_XML/PMC4820378_ann.xml
diff --git a/BioC_XML/4822050_v0.xml b/annotated_BioC_XML/PMC4822050_ann.xml
similarity index 100%
rename from BioC_XML/4822050_v0.xml
rename to annotated_BioC_XML/PMC4822050_ann.xml
diff --git a/BioC_XML/4822561_v0.xml b/annotated_BioC_XML/PMC4822561_ann.xml
similarity index 100%
rename from BioC_XML/4822561_v0.xml
rename to annotated_BioC_XML/PMC4822561_ann.xml
diff --git a/BioC_XML/4831588_v0.xml b/annotated_BioC_XML/PMC4831588_ann.xml
similarity index 100%
rename from BioC_XML/4831588_v0.xml
rename to annotated_BioC_XML/PMC4831588_ann.xml
diff --git a/BioC_XML/4832331_v0.xml b/annotated_BioC_XML/PMC4832331_ann.xml
similarity index 100%
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diff --git a/BioC_XML/4848090_v0.xml b/annotated_BioC_XML/PMC4848090_ann.xml
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diff --git a/BioC_XML/4855620_v0.xml b/annotated_BioC_XML/PMC4855620_ann.xml
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diff --git a/BioC_XML/4857006_v0.xml b/annotated_BioC_XML/PMC4857006_ann.xml
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diff --git a/BioC_XML/4869123_v0.xml b/annotated_BioC_XML/PMC4869123_ann.xml
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diff --git a/BioC_XML/4888278_v0.xml b/annotated_BioC_XML/PMC4888278_ann.xml
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diff --git a/BioC_XML/4896748_v0.xml b/annotated_BioC_XML/PMC4896748_ann.xml
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diff --git a/BioC_XML/4918759_v0.xml b/annotated_BioC_XML/PMC4918759_ann.xml
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diff --git a/BioC_XML/4919469_v0.xml b/annotated_BioC_XML/PMC4919469_ann.xml
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