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ATP binding by glutamyl-tRNA synthetase is switched to the productive mode by tRNA binding

2003; Springer Nature; Volume: 22; Issue: 3 Linguagem: Inglês

10.1093/emboj/cdg053

1460-2075

Shun‐ichi Sekine,

RNA Research and Splicing

Article3 February 2003free access ATP binding by glutamyl-tRNA synthetase is switched to the productive mode by tRNA binding Shun-ichi Sekine Shun-ichi Sekine Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Search for more papers by this author Osamu Nureki Osamu Nureki Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Department of Biophysics and Biochemistry, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033 Japan Search for more papers by this author Daniel Y. Dubois Daniel Y. Dubois Départements de Biochimie et Microbiologie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Stéphane Bernier Stéphane Bernier Chimie, Faculté des Sciences et de Génie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Robert Chênevert Robert Chênevert Chimie, Faculté des Sciences et de Génie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Jacques Lapointe Jacques Lapointe Départements de Biochimie et Microbiologie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Dmitry G. Vassylyev Corresponding Author Dmitry G. Vassylyev Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Search for more papers by this author Shigeyuki Yokoyama Corresponding Author Shigeyuki Yokoyama Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Department of Biophysics and Biochemistry, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033 Japan Genomic Sciences Center, RIKEN Yokohama Institute, 1-7-22 Suehiro-cho, Tsurumi, Yokohama, 230-0045 Japan Search for more papers by this author Shun-ichi Sekine Shun-ichi Sekine Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Search for more papers by this author Osamu Nureki Osamu Nureki Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Department of Biophysics and Biochemistry, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033 Japan Search for more papers by this author Daniel Y. Dubois Daniel Y. Dubois Départements de Biochimie et Microbiologie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Stéphane Bernier Stéphane Bernier Chimie, Faculté des Sciences et de Génie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Robert Chênevert Robert Chênevert Chimie, Faculté des Sciences et de Génie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Jacques Lapointe Jacques Lapointe Départements de Biochimie et Microbiologie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 Search for more papers by this author Dmitry G. Vassylyev Corresponding Author Dmitry G. Vassylyev Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Search for more papers by this author Shigeyuki Yokoyama Corresponding Author Shigeyuki Yokoyama Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan Department of Biophysics and Biochemistry, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033 Japan Genomic Sciences Center, RIKEN Yokohama Institute, 1-7-22 Suehiro-cho, Tsurumi, Yokohama, 230-0045 Japan Search for more papers by this author Author Information Shun-ichi Sekine1, Osamu Nureki1,2, Daniel Y. Dubois3, Stéphane Bernier4, Robert Chênevert4, Jacques Lapointe3, Dmitry G. Vassylyev 1 and Shigeyuki Yokoyama 1,2,5 1Cellular Signaling Laboratory and Structurome Group, RIKEN Harima Institute at SPring-8, 1-1-1 Kouto, Mikazuki-cho, Sayo, Hyogo, 679-5148 Japan 2Department of Biophysics and Biochemistry, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033 Japan 3Départements de Biochimie et Microbiologie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 4Chimie, Faculté des Sciences et de Génie, CREFSIP, Université Laval, Québec, Canada, G1K 7P4 5Genomic Sciences Center, RIKEN Yokohama Institute, 1-7-22 Suehiro-cho, Tsurumi, Yokohama, 230-0045 Japan *Corresponding authors. E-mail: [email protected] or E-mail: [email protected] The EMBO Journal (2003)22:676-688https://doi.org/10.1093/emboj/cdg053 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Aminoacyl-tRNA synthetases catalyze the formation of an aminoacyl-AMP from an amino acid and ATP, prior to the aminoacyl transfer to tRNA. A subset of aminoacyl-tRNA synthetases, including glutamyl-tRNA synthetase (GluRS), have a regulation mechanism to avoid aminoacyl-AMP formation in the absence of tRNA. In this study, we determined the crystal structure of the ‘non-productive’ complex of Thermus thermophilus GluRS, ATP and L-glutamate, together with those of the GluRS·ATP, GluRS·tRNA·ATP and GluRS·tRNA·GoA (a glutamyl-AMP analog) complexes. In the absence of tRNAGlu, ATP is accommodated in a ‘non-productive’ subsite within the ATP-binding site, so that the ATP α-phosphate and the glutamate α-carboxyl groups in GluRS· ATP·Glu are too far from each other (6.2 Å) to react. In contrast, the ATP-binding mode in GluRS·tRNA· ATP is dramatically different from those in GluRS·ATP·Glu and GluRS·ATP, but corresponds to the AMP moiety binding mode in GluRS·tRNA·GoA (the ‘productive’ subsite). Therefore, tRNA binding to GluRS switches the ATP-binding mode. The interactions of the three tRNAGlu regions with GluRS cause conformational changes around the ATP-binding site, and allow ATP to bind to the ‘productive’ subsite. Introduction The fidelity of genetic code translation is ensured by a set of aminoacyl-tRNA synthetases (aaRSs), each of which catalyzes the coupling of its specific tRNA(s) and amino acid with a high degree of accuracy. In general, aminoacylation is a two-step event. In the first step, an aaRS ‘activates’ the amino acid using ATP and Mg2+, yielding an enzyme-bound high energy intermediate, the aminoacyl-adenylate (aa-AMP). In the second step, the aminoacyl moiety is transferred from the aa-AMP to the 3′-terminal adenosine of tRNA. The aaRSs are divided into two classes, each consisting of 10 enzymes, on the basis of the two distinct ATP-binding cores (Eriani et al., 1990; Cusack, 1995). The class I ATP-binding site is formed with the conserved HIGH (His-Ile-Gly-His) and KMSKS (Lys-Met-Ser-Lys-Ser) sequence motifs, while the class II ATP-binding site is formed with three conserved motifs. Three of the class I aaRSs, the glutamyl-, glutaminyl- and arginyl-tRNA synthetases (GluRS, GlnRS and ArgRS, respectively), are known to share a special property: they catalyze the first step, amino acid activation, only in the presence of their cognate tRNA(s) (Ravel et al., 1965; Mitra and Mehler, 1966, 1967; Deutscher, 1967; Lee et al., 1967; Mehler and Mitra, 1967; Lapointe and Söll, 1972). As exceptions, several bacteria and archaea have the class I-type lysyl-tRNA synthetase (LysRS-I) instead of the class II-type LysRS (LysRS-II) (Ibba et al., 1997a, b). LysRS-I is structurally related to GluRS (Terada et al., 2002), and does not catalyze Lys-AMP formation in the absence of tRNA (Ibba et al., 1999). For these synthetases, the tRNA serves as the activator in the first step, and as the substrate in the second step of aminoacylation (Ravel et al., 1965; Mitra and Mehler, 1966, 1967; Lee et al., 1967; Mehler and Mitra, 1967; Kern and Lapointe, 1980). The activator function requires the integrity of the 3′ terminus of the tRNA, and chemical modification of this terminus abolishes the activity (Ravel et al., 1965; Mitra and Mehler, 1966; Lee et al., 1967; Kern and Lapointe, 1979). Glutamyl-AMP (Glu-AMP) is chemically unstable, and therefore it has been proposed that the tRNA-dependent glutamic acid activation is required to minimize ATP consumption, which would otherwise result from the spontaneous breakdown of the activated amino acid (Deutscher, 1967). On the basis of the crystal structure of the ternary complex of the Escherichia coli GlnRS, tRNAGln and ATP, it has been proposed that a characteristic structural element of GlnRS transfers the signal from the anticodon to the catalytic site (Rould et al., 1989, 1991). It was also reported that the 2′-hydroxyl group of the tRNA terminal adenosine is close to the bound ATP molecule (Perona et al., 1993). In the structure of another complex of GlnRS and tRNAGln with QSI (an analog of Gln-AMP), the 2′- and 3′-hydroxyl groups of the terminal adenosine form hydrogen bonds with the NH of the sulfamoyl group and the α-ammonium group, respectively, of QSI, indicating that the tRNA itself is involved in the Gln-AMP-binding site (Rath et al., 1998). In addition, the structures of the Thermus thermophilus GluRS (Nureki et al., 1995) and the GluRS·tRNAGlu complex (Sekine et al., 2001), the Saccharomyces cerevisiae ArgRS·L-arginine, ArgRS· tRNAArg and ArgRS·tRNAArg·L-arginine complexes (Cavarelli et al., 1998; Delagoutte et al., 2000), the T.thermophilus ArgRS (Shimada et al., 2001) and the Pyrococcus horikoshii LysRS-I and its complex with L-lysine (Terada et al., 2002) have been determined so far, but the mechanisms that prevent these four synthetases from catalyzing the amino acid activation in the absence of tRNA are not understood. In the present study, we determined four crystal structures of GluRS from T.thermophilus in the following complexes: GluRS·ATP·L-glutamate, GluRS·ATP, GluRS·tRNAGlu·ATP and GluRS· tRNAGlu·glutamol-AMP (GoA, a stable analog of Glu-AMP) (ERS/ATP/Glu, ERS/ATP, ERS/tRNA/ATP and ERS/tRNA/GoA, respectively; Table I). The high resolution structures of these T.thermophilus GluRS complexes showed that the ATP-binding site of GluRS has two distinct 'subsites’. In particular, the structure of the ternary GluRS·ATP·L-glutamate complex clearly shows that this complex is really ‘non-productive’: the ATP bound in this mode (the ‘non-productive’ subsite) in the absence of tRNAGlu is too far from the glutamate to react with it. In the presence of tRNAGlu, ATP binds to the ‘productive’ subsite, which is also used for binding the AMP moiety of the Glu-AMP analog. Interactions with the three regions of tRNAGlu cause conformational changes around the ATP-binding site of GluRS, thus switching the binding mode. Table 1. Crystallographic data and refinement statistics Complex 1 2 3 4 Code ERS/ATP/Glu ERS/ATP ERS/tRNA/ATP ERS/tRNA/GoA Macromolecules GluRS GluRS GluRS and tRNAGlu GluRS and tRNAGlu Ligands ATP and L-Glu ATP ATP Glutamol-AMP Data set SPring-8 BL station BL41XU BL45PX BL41XU BL41XU Space group P212121 P212121 C2221 C2221 Unit cell dimensions a = 81.92 b = 82.62 c = 83.11 Å a = 82.72 b = 84.06 c = 83.45 Å a = 110.69 b = 219.12 c = 135.22 Å a = 110.49 b = 219.87 c = 135.12 Å Resolution (Å) 40−1.8 50−1.9 50−2.4 50−2.1 Total reflections 208 385 175 753 247 929 426 837 Unique reflections 49026 46014 59840 92817 Completeness (%) 91.7 (92.8)a 98.3 (96.9) 91.2 (80.8) 96.4 (88.5) Rmergeb (%) 6.3 (44.4)c 5.9 (36.7) 7.2 (35.7) 7.4 (34.2) Refinement statistics Resolution (Å) 40−1.8 50−1.9 50−2.4 50−2.1 Reflections 48 560 45 764 58 889 90 631 Rcrystd (%) 19.9 20.9 21.3 21.9 Rfreed (%) 22.7 23.6 25.7 25.9 No. of protein atoms 3813 3813 7626 7626 No. of tRNA atoms – – 3194 3194 No. of water atoms 352 344 505 739 No. of ligand atoms 45 35 64 76 R.m.s.d. bonds (Å) 0.005 0.005 0.006 0.006 R.m.s.d. angles (°) 1.2 1.2 1.3 1.2 R.m.s.d. improper angles (°) 0.85 0.84 1.5 1.4 Ramachandran plot (most favorable region) (%) 94.8 95.5 93.7 93.7 a The completeness in the highest resolution shell is given in parentheses. b Rmerge = Σ|I − |/ΣI, where I is the observed intensity of reflections. c Rmerge in the highest resolution shell is given in parentheses. d Rcryst, free = Σ|Fobs − Fcalc|/ΣFobs, where the crystallographic R-factor is calculated including and excluding refinement reflections. In each refinement, free reflections consist of 5% of the total number of reflections. Results and discussion Structure determination The crystal structure of ERS/ATP/Glu (complex 1, Table I) was solved by molecular replacement using the free GluRS structure (Nureki et al., 1995) as the search model, and was refined against data to 1.8 Å resolution (Figure 1A). The crystallographic asymmetric unit contains one ERS/ATP/Glu. Under nearly the same conditions, the ERS/ATP crystals (complex 2, Table I) were obtained in the absence of glutamate. The structure was determined by molecular replacement, and refined to 1.9 Å resolution (Table I). The overall enzyme structures in ERS/ATP/Glu and ERS/ATP are almost the same; the root mean square deviation (r.m.s.d.) is 0.54 Å over all of the protein atoms. Figure 1.Thermus thermophilus GluRS crystal structures. (A) Ribbon representation of the ERS/ATP/Glu structure. Five domains, the Rossmann fold (1), connective peptide (or acceptor-binding) (2), stem-contact fold (3) and two anticodon-binding (4 and 5) domains, are colored khaki, light blue, pink, steel blue and deep blue, respectively. The HVGT and KISKR motifs of GluRS are highlighted in purple. The ATP and glutamate molecules in the GluRS catalytic pocket are shown in green. (B) Overall structure of ERS/tRNA/ATP. The ATP and tRNAGlu molecules in the complex are shown in orange and turquoise, respectively. These figures were produced using the MOLSCRIPT (Kraulis, 1991) and RASTER3D (Merritt and Murphy, 1994) programs. Download figure Download PowerPoint The ERS/tRNA/ATP (Figure 1B) and ERS/tRNA/GoA structures (complexes 3 and 4) were determined and refined to 2.4 and 2.1 Å resolution, respectively (Table I). Each structure contains two nearly identical complexes (a and b) in the crystallographic asymmetric unit. Herein, the discussion is based on the complex a structures, unless otherwise specified. The overall enzyme and tRNA structures in ERS/tRNA/ATP and ERS/tRNA/GoA are practically the same (r.m.s.d. = 0.41 Å over all of the protein and RNA atoms). Both of the complexes are superposable on the previous GluRS·tRNAGlu binary complex (ERS/tRNA) (Sekine et al., 2001) (r.m.s.d. ∼0.71 Å over all of the protein and RNA atoms for both comparisons). Herein, these GluRS structures are compared by superposition of the enzyme catalytic cores (domains 1 and 3). The catalytic domain structures in the ERS/ATP/Glu, ERS/ATP and ERS/tRNA/ATP structures (complexes 1–3) were fitted to those in the ERS/tRNA/GoA (complex 4) as a reference, by using the LSQKAB program (CCP4, 1994) (r.m.s.d. values are 0.49, 0.50 and 0.22 Å, respectively, for 156 Cα atoms). Flexible regions were excluded from the calculations. The differences thus found are described in the following sections. Conformational changes of GluRS upon tRNAGlu binding A comparison of the tRNA-free and tRNA-bound GluRS structures reveals the tRNA-dependent conformational changes within the domain linkers and loops (Figure 2A). For the specific interaction of the two anticodon-binding domains (4 and 5) with the tRNA anticodon loop (Sekine et al., 2001), these domains are reoriented, without changing their folds, by an ∼6° rotation of domain 4 relative to domain 3 (the SC fold domain), and by an ∼8° rotation of domain 5 relative to domain 4 (Figure 2A). In the present crystal protein structures, we could not see any direct structural linkage that may transmit the anticodon-binding signal to the catalytic site, in contrast to the structure of the E.coli GlnRS·tRNA·ATP complex (QRS/tRNA/ATP) (Rould et al., 1989, 1991). Figure 2.Conformational changes within GluRS upon tRNAGlu binding. (A) The ERS/tRNA/GoA backbone structure was superposed on that of ERS/ATP/Glu by the enzyme catalytic core (domains 1 and 3) (stereo view). The entire ERS/ATP/Glu structure is colored gray, while the ERS/tRNA/GoA structure is colored as in Figure 1B. The arrows indicate the tRNA-induced conformational changes within GluRS. Three tRNA regions involved in the enzyme active site rearrangement are highlighted in orange. (B) A stereo view showing the 3′-terminal region of the tRNAGlu in ERS/tRNA/GoA, and its interactions with GluRS. These interactions are the same as those observed in ERS/tRNA/ATP. Download figure Download PowerPoint The CP domain (domain 2) also changes its orientation upon tRNAGlu binding (Figure 2A). It rotates by ∼7° relative to domain 1, and thus binds the 3′ acceptor end of the tRNA molecule. The tRNAGlu C74 is trapped by a pocket formed on the CP domain, without stacking on the other bases in the 3′-terminal region (Figure 2B). The C74 base is recognized specifically by interactions mainly with the main chains of Glu107 and Ser181, while the C74 phosphate interacts with Arg147. Here, the 3′ end is bent back toward the enzyme active site, resulting in a hairpin conformation. In the GluRS-bound tRNAGlu, the G1·C72 base pair of the acceptor stem is intact (Figure 2B), in contrast to the disrupted U1·A72 base pair in the E.coli GlnRS-bound tRNAGln (Rould et al., 1989). Correspondingly, GlnRS possesses a unique insertion structure that interacts with A72, whereas GluRS does not (Nureki et al., 1995). The A73 base does not interact with any protein residues, but is involved in a stacking array formed by the A73, C75 and A76 bases and the Trp209 and Tyr187 rings (Figure 2B). The 2′-hydroxyl group of C75 interacts with the conserved Asp44 and Arg47 side chains on the Rossmann fold (domain 1). The phosphate group of the terminal adenosine (A76) interacts with Lys180 and Tyr187. The 5′-hydroxyl group of A76 interacts with the Thr43 side chain. In the previous structure of ERS/tRNA (Sekine et al., 2001), the electron density corresponding to C75 and A76 of the tRNA molecule was poor. Finally, tRNA-induced conformational changes are observed in the loop of residues 206–209 in the Rossmann fold domain and in the ‘KMSKS’ region in the SC fold domain (Figure 2A), which are described below. ATP and Glu binding in the absence of tRNA The 1.8 Å resolution structure of ERS/ATP/Glu (Table I; Figure 1A) was the first to reveal both the enzyme-bound ATP and amino acid substrates without a reaction (Figure 3A). This is in good agreement with the observations that the E.coli and T.thermophilus GluRSs are absolutely inactive in the absence of their cognate tRNAGlu (Lapointe and Söll, 1972; S.Sekine and S.Yokoyama, unpublished). The ATP-Mg2+ substrate interacts intensively with the region including the 243KISKR247 motif (the ‘KMSKS’ motif of the T.thermophilus GluRS) (Figure 3B). The KISKR loop has a different conformation from that within the substrate-free GluRS (Nureki et al., 1995) (not shown), which suggests an induced fit upon ATP binding. The adenine base is accommodated in a hydrophobic pocket formed by His15, Tyr20, Leu235, Leu236 and Ile244 (Figure 3B; Supplementary figure 1, available at The EMBO Journal Online). The N1 and N6 of the adenine make hydrogen bonds with the main chains of Leu236 and Ile244. The ATP phosphate groups hydrogen-bond extensively with Lys243, Ser245, Lys246 and Arg247. The Mg2+ ion has a unique octahedral coordination by the ATP α-, β- and γ-phosphate oxygen atoms and the three water molecules, which fix the ATP phosphate conformations (Figure 3A and B). The Glu208 and Lys243 side chains interact with the Mg2+ ion through water molecules. The 2′-hydroxyl group of the ATP ribose in the C2′-endo conformation interacts with the Glu208 and Trp209 side chains, while the back of the ATP interacts with the 15HVGT18 motif (the ‘HIGH’ motif of the T.thermophilus GluRS). Figure 3.The ATP and glutamate molecules in ERS/ATP/Glu. (A) A stereo view of the electron density, showing the ATP-Mg2+ and glutamate molecules in ERS/ATP/Glu. An annealed |Fo − Fc| omit electron density map was calculated using all of the data from 40 to 1.8 Å resolution and the complex model without the ATP-Mg2+ and glutamate. The refined models of the ATP-Mg2+ and glutamate are superimposed on the density countered at 3σ. The Mg2+ ion is shown by a yellow sphere. The average distance between the Mg2+ ion and the six liganded oxygen atoms is 2.02 Å. (B) ATP recognition in ERS/ATP/Glu (stereo view). The ATP recognition in this complex is the same as that in ERS/ATP. (C) Glutamate recognition in ERS/ATP/Glu (stereo view). Download figure Download PowerPoint On the other hand, the glutamate molecule lies along a β-strand, and its α-ammonium group hydrogen-bonds with the main chain of Ala7 and the side chains of Ser9 and Glu41 (Figures 3C and 4A). Four residues, Arg5, Tyr187, Asn191 and Arg205, constitute a pocket complementary to the γ-carboxyl group of glutamate, which determines the amino acid specificity. Remarkably, the α-carboxyl group of glutamate and the α-phosphate group of ATP are 6.2 Å apart (Figure 4A), and therefore are too far from each other to react. The glutamate α-carboxyl group instead interacts with the 3′-hydroxyl group of the ATP ribose. It should be noted that the position and the conformation of the ATP molecule in ERS/ATP/Glu are identical to those in ERS/ATP (Figures 4A and B, and 5A). Therefore, the non-productive arrangement of the two substrates in ERS/ATP/Glu (Figures 3A and 4A) is not due to the repulsion between the negatively charged carboxyl and phosphate groups. Thus, both the ATP and glutamate molecules can bind tightly to the GluRS substrate-binding site in the absence of tRNAGlu, but their arrangement is non-productive. This ‘dead-end’ ternary complex explains simply why the amino acid activation does not occur in the absence of tRNA. Figure 4.Substrate/ligand(s) binding in the GluRS complexes. (A–D) The GluRS catalytic site structures in the present complexes are shown in the same orientation. The HVGT and KISKR motifs are highlighted in purple. (A) The ERS/ATP/Glu structure. The ATP-Mg2+ and glutamate molecules are shown in green. (B) The ERS/ATP structure. The ATP-Mg2+ is colored light blue. (C) The ERS/tRNA/ATP structure. The ATP molecule is colored salmon, and the 3′-terminal adenosine (A76) of tRNAGlu is cyan. (D) The ERS/tRNA/GoA structure. The GoA (glutamol-AMP) molecule is colored yellow. (E) A stereo view showing the ATP recognition in ERS/tRNA/ATP. (F) A stereo view showing the GoA recognition in ERS/tRNA/GoA. Download figure Download PowerPoint Figure 5.Comparisons of the substrate/ligand positions among the GluRS complexes. (A) The ATP molecule (light blue) in ERS/ATP is compared with the ATP and glutamate (green) in ERS/ATP/Glu by superposition of the enzyme catalytic site structures. (B) The ATP (salmon) in ERS/tRNA/ATP is compared with the ATP and glutamate (green) in ERS/ATP/Glu by superposition. (C) The GoA (yellow) in ERS/tRNA/GoA is compared with the ATP and glutamate (green) in ERS/ATP/Glu by superposition. (D) The GoA (yellow) in ERS/tRNA/GoA is compared with the ATP (salmon) in ERS/tRNA/ATP by superposition. Download figure Download PowerPoint ATP binding in the presence of tRNA The structure of ERS/tRNA/ATP has been determined at 2.4 Å resolution (Table I, Figure 1B). A comparison of this complex with that of ERS/ATP/Glu (or ERS/ATP) reveals a remarkable difference in the modes of ATP binding (Figure 4C). In ERS/tRNA/ATP, the ATP molecule binds to the catalytic pocket in a rotated orientation in the same plane, by ∼37° around an axis near C2 of the adenine ring, relative to those in the ERS/ATP/Glu and ERS/ATP structures (Figures 4C and 5B). In ERS/tRNA/ATP, the adenine ring is accommodated in the same hydrophobic pocket, while the base is in a different orientation (Figure 4C). The ATP ribose is largely shifted, and is docked in the depths of the active site cleft. The 2′-hydroxyl group loses the hydrogen bond with Trp209, but gains a new hydrogen bond with the amide group of Ala206 (Figure 4E). It has been reported that 2′-deoxy ATP is a poor substrate for E.coli GluRS (Kern and Lapointe, 1979), which suggests that the ATP ribose binding to Ala206 is important for the reaction. The 3′-hydroxyl group of the ATP ribose interacts with a water molecule, which is bound to Arg5 and Ile204 (Supplementary figure 1C). The Ile21 side chain binds the sugar and base portions of the ATP by van der Waals contacts (Figure 4E). On the other hand, the interactions of the ATP phosphate groups with the KISKR residues (except Ile244) are missing. Instead, the ATP phosphates interact with the Thr11, Thr18 and Arg47 side chains. The tRNAGlu accommodates the 3′-terminal adenosine into the GluRS active site pocket by assuming a hairpin conformation in the CCA region (Figure 2B). It is remarkable that the 2′-hydroxyl group of the 3′-terminal adenosine of tRNAGlu (A76) hydrogen-bonds with the α- and γ-phosphates of the ATP (Figure 4E). Thus, in the presence of tRNAGlu, the ATP molecule is located deeper in the GluRS active site cleft (Figure 4C), and is fixed in a subsite distinct from that in the absence of tRNA (Figure 4A and B). The E.coli GluRS exhibits practically the same dissociation constants for ATP (Kd = 90 μM) in either the absence or presence of tRNAGlu (Kern and Lapointe, 1979), which suggests that the ATP-binding affinities of the two subsites are almost the same. Binding of the glutamyl-AMP analog GoA is a stable non-hydrolyzable analog of Glu-AMP (Desjardins et al., 1998) (Supplementary figure 2). The stability is achieved by minimal replacement of the labile anhydride function of Glu-AMP by a phosphate ester (the C=O moiety is reduced to a CH2). GoA belongs to the group of synthetic aminoalkyl adenylates, which are strong inhibitors of the corresponding aaRSs. It is a competitive inhibitor of the E.coli GluRS (Ki = 3 μM) (Desjardins et al., 1998). In the present study, we confirmed that GoA is also a competitive inhibitor of the T.thermophilus GluRS (Ki = 1.2 μM). Based on the 2.1 Å structure of the ERS/tRNA/GoA complex (Table I), the ATP binding observed in ERS/tRNA/ATP is concluded to be ‘productive’ binding. The ERS/tRNA/GoA structure represents the enzyme state after the first step and before the second step of aminoacylation. In this complex, the glutamol moiety of the analog is accommodated in the same pocket as the glutamate substrate in ERS/ATP/Glu (Figures 4D and F, and 5C). The A76 ribose directly interacts with the main chain part of the glutamol moiety (Figure 4F), consistent with the previous observation that the T.thermophilus GluRS can discriminate glutamate from non-cognate amino acids only in the presence of tRNAGlu (Hara-Yokoyama et al., 1986). On the other hand, the adenosine moiety of GoA is in a rotated orientation, as compared with that of the ATP in the ERS/ATP/Glu and ERS/ATP structures (Figures 4D and 5C). The adenosine binding is the same as that in ERS/tRNA/ATP and, remarkably, the phosphate group of GoA is superimposed on the ATP α-phosphate (Figure 5D). In fact, the distance between the glutamate α-carboxyl oxygen in ERS/ATP/Glu and the ATP α-phosphorus in ERS/tRNA/ATP is ∼2.9 Å upon superposition (Figure 5B). Thus, GluRS possesses two modes for ATP binding, the ‘non-productive’ and ‘productive’ binding modes, which can be switched in a tRNA-dependent manner. In the absence of tRNAGlu, the ATP-binding mode of GluRS is ‘non-productive’ (Figure 4A and B). GluRS, ATP and glutamate form a ‘dead-end’ complex (Figures 3A and 4A), which prevents Glu-AMP formation in the absence of tRNA, presumably to avoid the possible waste of the energy source on the enzyme, in the context of the instability of Glu-AMP (see above) and of the high intracellular glutamate concentration in many prokaryotes and eukaryotes (reviewed by Metzler, 1981; Csonka et al., 1989; Danbolt, 2001). When GluRS is in complex with tRNAGlu, ATP can bind to the ‘productive’ subsite (Figure 4C) to initiate the amino acid activation, probably via a penta-covalent transition state as suggested earlier for E.coli GlnRS (Perona et al., 1993), yielding the enzyme-bound Glu-AMP (Figure 4D). Active site rearrangement within GluRS upon tRNAGlu binding In order to examine how the choice is made between the two subsites for ATP binding, the ERS/ATP(/Glu) and ERS/tRNA/ATP structures were compared. The structural differences indicate that the interactions of GluRS with three regions of tRNAGlu are likely to account for the tRNA-dependent switching of the ATP-binding mode from ‘non-productive’ to ‘productive’. First, the D stem of tRNAGlu interacts with the SC fold (Figures 2A and 6A). The SC fold domain, specific to the class Ia and Ib aaRSs, is located between the Rossmann-fold and the anticodon-binding domains, and includes the KISKR (‘KMSKS’) loop (Sugiura et al., 2000) (Figure 1). In the ERS/tRNA/ATP structure, the backbone of nucleotide residues 10–13 in the D stem interacts with one of the β-strands (amino acid residues 301–305) and the KISKR loop of th