Insights into Ligand Binding and Catalysis of a Central Step in NAD+ Synthesis
2001; Elsevier BV; Volume: 276; Issue: 10 Linguagem: Inglês
10.1074/jbc.m008810200
ISSN1083-351X
AutoresV. Saridakis, Dinesh Christendat, Matthew S. Kimber, Akil Dharamsi, A.M. Edwards, E.F. Pai,
Tópico(s)Catalytic C–H Functionalization Methods
ResumoNicotinamide mononucleotide adenylyltransferase (NMNATase) catalyzes the linking of NMN+ or NaMN+ with ATP, which in all organisms is one of the common step in the synthesis of the ubiquitous coenzyme NAD+, via both de novo and salvage biosynthetic pathways. The structure of Methanobacterium thermoautotrophicum NMNATase determined using multiwavelength anomalous dispersion phasing revealed a nucleotide-binding fold common to nucleotidyltransferase proteins. An NAD+ molecule and a sulfate ion were bound in the active site allowing the identification of residues involved in product binding. In addition, the role of the conserved16HXGH19 active site motif in catalysis was probed by mutagenic, enzymatic and crystallographic techniques, including the characterization of an NMN+/SO 42–complex of mutant H19A NMNATase.1ej21hyb Nicotinamide mononucleotide adenylyltransferase (NMNATase) catalyzes the linking of NMN+ or NaMN+ with ATP, which in all organisms is one of the common step in the synthesis of the ubiquitous coenzyme NAD+, via both de novo and salvage biosynthetic pathways. The structure of Methanobacterium thermoautotrophicum NMNATase determined using multiwavelength anomalous dispersion phasing revealed a nucleotide-binding fold common to nucleotidyltransferase proteins. An NAD+ molecule and a sulfate ion were bound in the active site allowing the identification of residues involved in product binding. In addition, the role of the conserved16HXGH19 active site motif in catalysis was probed by mutagenic, enzymatic and crystallographic techniques, including the characterization of an NMN+/SO 42–complex of mutant H19A NMNATase.1ej21hyb nicotinic acid dinucleotide multiwavelength anomalous dispersion nicotinamide mononucleotide adenylyltransferase wild type glycerol-3-phosphate cytidyltransferase phosphopantetheine adenylyltransferase nicotinic acid mononucleotide root mean square deviation Nicotinamide mononucleotide adenylyltransferase (EC 2.7.7.1) catalyzes the synthesis of nicotinamide adenine dinucleotide (NAD+) or nicotinic acid dinucleotide (NaAD+)1 from nicotinamide mononucleotide (NMN+) or nicotinic acid mononucleotide (NaMN+), respectively, by transferring the adenylyl part of ATP and concomitantly releasing pyrophosphate (PPi) (Fig. 1 A). The reaction product, NAD+, plays a central role in many cellular processes; it functions as a coenzyme in reduction-oxidation reactions and as a substrate in DNA ligation and protein ADP-ribosylation reactions (1Magni G. Amici A. Emanuelli M. Raffaelli N. Ruggieri S. Adv. Enzymol. Relat. Areas Mol. Biol. 1999; 73: 135-182PubMed Google Scholar). There is also considerable medical interest in this enzyme as it is implicated in the metabolism of the antitumor drug tiazofurin (2Jayaram H.N. Pillwein K. Lui M.S. Faderan M.A. Weber G. Biochem. Pharmacol. 1986; 35: 587-593Crossref PubMed Scopus (30) Google Scholar). In vivo, tiazofurin is phosphorylated to tiazofurin monophosphate and then converted by NMNATase to the actual pharmacophore thiazole-4-carboxamide adenine dinucleotide, an analog of NAD+. Thiazole-4-carboxamide adenine dinucleotide is a potent inhibitor of inosine monophosphate dehydrogenase causing arrest of guanylate biosynthesis and thus inhibition of tumor cell proliferation. Consistent with this interpretation, low NMNATase activity is observed in cancer patients showing resistance to tiazofurin therapy (2Jayaram H.N. Pillwein K. Lui M.S. Faderan M.A. Weber G. Biochem. Pharmacol. 1986; 35: 587-593Crossref PubMed Scopus (30) Google Scholar). Two mechanisms for the NMNATase-catalyzed synthesis of NAD+have been postulated; the first one assumes a double displacement reaction that involves the formation of an adenylyl enzyme covalent intermediate upon release of pyrophosphate followed by transfer of the adenylyl group to NMN+ to form NAD+, whereas the second one describes a nucleophilic attack of the 5′-phosphate of NMN+ on the α-phosphate of ATP to form NAD+and releasing PPi. The latter mechanism is supported by17O NMR studies of NAD+ synthesis (3Lowe G. Tansley G. Eur. J. Biochem. 1983; 132: 117-120Crossref PubMed Scopus (19) Google Scholar), but a complete understanding of the catalytic chemistry awaited more detailed structural information. NMNATase proteins have been identified, purified, and characterized from archaea, bacteria, and eukarya. All of these proteins are oligomeric; trimeric, tetrameric, and hexameric forms have been observed (1Magni G. Amici A. Emanuelli M. Raffaelli N. Ruggieri S. Adv. Enzymol. Relat. Areas Mol. Biol. 1999; 73: 135-182PubMed Google Scholar). Several NMNATase genes have been sequenced from a variety of sources (Fig. 1 B). Although these gene products remain annotated in the GenBankTM data base as of unknown function, related sequences from Methanococcus jannaschii, Escherichia coli, Synechocystis sp., and Sulfolobus solfataricus have been overexpressed as recombinant proteins inE. coli and shown to exhibit NMNATase activity (4Raffaelli N. Pisani F.M. Lorenzi T. Emanuelli M. Amici A. Ruggieri S. Magni G. J. Bacteriol. 1997; 179: 7718-7723Crossref PubMed Google Scholar, 5Raffaelli N. Emanuelli M. Pisani F.M. Amici A. Lorenzi T. Ruggieri S. Magni G. Mol. Cell. Biochem. 1999; 193: 99-102Crossref PubMed Google Scholar, 6Raffaelli N. Lorenzi T. Amici A. Emanuelli M. Ruggieri S. Magni G. FEBS Lett. 1999; 444: 222-226Crossref PubMed Scopus (42) Google Scholar, 7Emmanuelli M. Carnevali F. Lorenzi M. Raffaelli N. Amici A. Ruggieri S. Magni G. FEBS Lett. 1999; 455: 13-17Crossref PubMed Scopus (51) Google Scholar, 8Raffaelli N. Lorenzi T. Mariani P.L. Emanuelli M. Amici A. Ruggieri S. Magni G. J. Bacteriol. 1999; 181: 5509-5511Crossref PubMed Google Scholar). Recently, the crystal structure of NMNATase from M. jannaschii in complex with ATP and Mg2+ was reported (9D'Angelo I. Raffaelli N. Dabusti V. Lorenzi T. Magni G. Rizzi M. Struct. Fold. Des. 2000; 8: 993-1004Abstract Full Text Full Text PDF Scopus (67) Google Scholar). Our results on the NAD+ and NMN+ complexes of the Methanobacterium thermoautotrophicum enzyme complement this result, especially when interpreting the catalytic mechanism of NMNATase. We describe the crystal structures of the NAD+ complex of NMNATase and of the NMN+complex of an active site mutant (H19A) of NMNATase at 1.9 and 2.5 Å resolution, respectively. These structural results, combined with mutagenesis and enzymatic experiments, define the spatial geometry of the ligand binding sites, identify residues with potential roles in substrate binding and catalysis, and suggest aspects of the product release mechanism. The NMNATase gene (GenBankTM accession number AE000803) was amplified by polymerase chain reaction using M. thermoautotrophicum genomic DNA and cloned into the pET15b (Novagen) expression vector at the NdeI and BglII sites. Recombinant NMNATase was overexpressed in E. coliBL21 Gold (DE3) cells (Stratagene) harboring a plasmid encoding rareE. coli tRNA genes. The cells were grown at 37 °C in Luria-Bertoni broth with carbenicillin (50 μg/ml) and kanamycin (50 μg/ml) to an A 600 nm of 0.7 and induced overnight with 0.5 mmisopropyl-β-d-thiogalactopyranoside at 24 °C. The bacteria were harvested by centrifugation and resuspended in binding buffer (50 mm Tris, 500 mm NaCl, 5% glycerol, and 5 mm imidazole) supplemented with 2 mmphenylmethylsulfonyl fluoride. Bacteria were lysed by several passages through a French pressure cell at 1.4 × 108 pascals, and DNA was sheared by sonication. Cell debris was removed through centrifugation for 30 min at 20,000 × g. ContaminatingE. coli proteins were removed by heating for 20 min at 55 °C followed by centrifugation at 5000 × g for 30 min. The supernatant was applied by gravity to a DE52 column (Whatman) immediately coupled to a Ni2+ column (Qiagen). The Ni2+ column was washed with 50 volumes of binding buffer containing 30 mm imidazole. The bound NMNATase was eluted from the Ni2+ column with 500 mm imidazole in binding buffer. The hexahistidine tag was cleaved by digesting for 16 h with thrombin (1 μg of thrombin per mg of recombinant protein) at 4 °C in binding buffer made 2.5 mm in CaCl2. NMNATase was then dialyzed against 500 mm NaCl in 10 mm HEPES (pH 7.5) and concentrated to 10 mg/ml using BioMax concentrators (Millipore). Mutant H19A NMNATase was purified as described above for WT NMNATase. For the preparation of selenomethionine (Se-Met)-enriched protein, NMNATase was expressed in a methionine auxotroph strain B834(DE3) of E. coli (Novagen) and purified under the same conditions as native NMNATase with the addition of 5 mmβ-mercaptoethanol in all buffers. Screening for crystallization conditions was performed using Hampton Research Crystal Screens I and II at room temperature in VDX plates with the hanging drop vapor diffusion method. 2 μl of protein solution (10 mg/ml) were mixed with 2 μl of the various reservoir solutions and equilibrated with 500 μl of this solution. Crystals in the form of hexagonal rods appeared after 24 h in crystallization set-ups containing ammonium sulfate or lithium sulfate as precipitant. Crystals selected for native and multiwavelength anomalous dispersion (MAD) data collection were grown in 1.5 m LiSO4 and 100 mm HEPES at pH 7.5 at 20 °C. Crystals of H19A NMNATase were grown in 1.6m LiSO4 and 100 mm HEPES at pH 7.5 at 20 °C. Making use of the anomalous scattering of selenium atoms, a three-wavelength MAD experiment was carried out at 100 K on beamline BM14D, APS, using a Q1 CCD detector (Area Detector Systems Corp.). The crystal was flash-frozen with crystallization buffer plus 30% glycerol as cryoprotectant. Diffraction data from native crystals of WT and H19A NMNATase were collected on beamline BM14C, APS, at 100 K using a Q4 CCD detector (ADSC). Both MAD and native data were processed and scaled with the DENZO/SCALEPACK suite of programs (10Otwinowski Z. Minor W. Methods Enzymol. 1997; 276: 307-326Crossref PubMed Scopus (38592) Google Scholar). The selenium sites were located using SOLVE (11Terwilliger T.C. Berendzen J. Acta Crystallogr. Sect. D Biol. Crystallogr. 1999; 55: 849-861Crossref PubMed Scopus (3219) Google Scholar) and refined using SHARP (12de la Fortelle E. Bricogne G. Methods Enzymol. 1997; 276: 472-494Crossref PubMed Scopus (1797) Google Scholar). The electron density map was improved using Density Modification from the CCP4 package (13Bailey S. Acta Crystallogr. Sect. D Biol. Crystallogr. 1994; 50: 760-763Crossref PubMed Scopus (42) Google Scholar). Model building was done with O (14Jones T.A. Zou J.Y. Cowan S.W. Kjeldgaard M. Acta Crystallogr. Sect. A. 1991; 47: 110-119Crossref PubMed Scopus (13014) Google Scholar), and CNS (crystallography and NMR system) (15Brunger A.T. Adams P.D. Clore G.M. DeLano W.L. Gros P. Grosse-Kunstleve R.W. Jiang J.S. Kuszewski J. Nilges M. Pannu N.S. Read R.J. Rice L.M. Simonson T. Warren G.L. Acta Crystallogr. Sect. D Biol. Crystallogr. 1998; 54: 905-921Crossref PubMed Scopus (16978) Google Scholar) was used for refinement. Water molecules were initially picked using CNS and then manually verified in O using the following criteria: a peak of at least 2.5 ς in an Fo − Fcmap, a peak of at least 1.0 ς in a 2Fo −Fc map, and reasonable intermolecular interactions. Crystallographic and refinement statistics are found in TablesI and II, respectively. The programs MOLSCRIPT (16Kraulis P.J. J. Appl. Crystallogr. 1991; 24: 946-950Crossref Google Scholar), RASTER 3D (17Merrit E.A. Murphy M.E.P. Acta Crystallogr. Sect. D Biol. Crystallogr. 1991; 50: 869-873Crossref Scopus (2858) Google Scholar), SPOCK (18Christopher, J. A (1998) SPOCK (Structural Properties Observation and Calculation Kit), The Center for Macromolecular Design, Texas A & M University, College Station, TX.Google Scholar), and LIGPLOT (19Wallace A.C. Laskowski R.A. Thornton J.M. Prot. Eng. 1995; 8: 127-134Crossref PubMed Scopus (4421) Google Scholar) were used in the production of the figures.Table ISummary of data collection statisticsX-ray dataNativePeakEdgeRemoteH19ASpace groupP6322P6322P6322P6322P6322Unit cell (Å3)89.0 × 89.0 × 109.989.2 × 89.2 × 110.389.2 × 89.2 × 110.389.2 × 89.2 × 110.389.7 × 89.7 × 109.7Resolution (Å)1.92.92.92.92.5Wavelength (λ)1.000000.979540.979300.953731.0000Se sites (no.)444Total observations (no.)49252612525112632512254860748Unique reflections (no.)245249311931193119479Intensity (I/ς〈I〉)37 (5)33 (10)30 (8)27 (6)30.2 (5.4)Completeness (%)99.4 (99.0)98.8 (99.5)98.3 (97.6)97.1 (97.3)99.3 (97.1)R sym 1-aRsym = Σ‖I − 〈I〉‖/ΣI, where I is the observed intensity and 〈I〉 is the average intensity from multiple observations of symmetry-related reflections.0.041 (0.351)0.079 (0.238)0.082 (0.249)0.076 (0.285)0.043 (0.273)Figure of merit 1-bFigure of merit = ‖ΣP(α)eiα‖/ΣP(α), where P(α) is the phase probability distribution and α is the phase angle. (%)40/75 1-cNumbers before and after slash show values before and after solvent flattening and histogram matching with Density Modification, respectively.Numbers in parentheses refer to the highest resolution shell, 1.97–1.90 Å for the native data, 3.01–2.9 Å for the MAD data, and 2.59–2.50 Å for the mutant data.1-a Rsym = Σ‖I − 〈I〉‖/ΣI, where I is the observed intensity and 〈I〉 is the average intensity from multiple observations of symmetry-related reflections.1-b Figure of merit = ‖ΣP(α)eiα‖/ΣP(α), where P(α) is the phase probability distribution and α is the phase angle.1-c Numbers before and after slash show values before and after solvent flattening and histogram matching with Density Modification, respectively. Open table in a new tab Table IISummary of refinement statisticsWTH19AR cryst aRcryst = Σ‖F obs −F calc‖/‖F obs‖.0.2120.236R free0.2420.294Protein atoms (no.)13401275Water molecules (no.)11910NAD atoms (no.)44NMN atoms (no.)22Sodium ions (no.)1Sulfate ions (no.)11r.m.s.d. bond lengths (Å)0.0160.008r.m.s.d. bond angles (°)1.81.3r.m.s.d. dihedrals (°)23.922.0Average main chainB-factor (Å2)33.742.7Average side chainB-factor (Å2)36.047.6Average ligandB-factor (Å2)38.359.1Rfree was calculated using randomly selected reflections (10%).a Rcryst = Σ‖F obs −F calc‖/‖F obs‖. Open table in a new tab Numbers in parentheses refer to the highest resolution shell, 1.97–1.90 Å for the native data, 3.01–2.9 Å for the MAD data, and 2.59–2.50 Å for the mutant data. Rfree was calculated using randomly selected reflections (10%). Gel filtration of NMNATase was performed with a Superdex 200 prep 16/60 (Amersham Pharmacia Biotech) column equilibrated with 10 mm HEPES and 500 mmNaCl using high pressure liquid chromatography (LKB-Wallac). Protein standards included aldolase, bovine serum albumin, ovalbumin, and cytochrome c. Chromatography was performed at 4 °C at a flow rate of 0.5 ml/min. Site-directed mutagenesis to change His-19 to Ala was carried out using QuikChange™ (Stratagene). The mutagenic primers were 5′-CACAGGGGCGCACTGCAGGTC and 5′-GACCTGCAGTGCGCCCCTGTG for H19A. DNA encoding WT NMNATase cloned into pET-15B was used as template for the polymerase chain reaction mutagenesis reaction. Briefly, 25 ng of template DNA was incubated with the appropriate mutagenic primers, dNTPs, andPfu DNA polymerase using the cycling parameters recommended in the supplier's manual. Following the temperature cycling step,DpnI was added to each amplification reaction and incubated at 37 °C for 6 h followed by transformation of the mutagenized plasmid into XL2 Blue cells. WT NMNATase activity was measured in a coupled assay according to Raffaelli et al. (6Raffaelli N. Lorenzi T. Amici A. Emanuelli M. Ruggieri S. Magni G. FEBS Lett. 1999; 444: 222-226Crossref PubMed Scopus (42) Google Scholar). Varying amounts of NMNATase (1–1000 ng) were incubated with 2 mmNMN+, 2 mm ATP, 10 mmMgCl2, and 50 mm HEPES (pH 7.5) at 65 °C for 20 min. The amount of NAD+ formed was measured spectrophotometrically at 340 nm using alcohol dehydrogenase to convert NAD+ to NADH. The enzymatic activity of H19A NMNATase was measured the same way but varying the amount of enzyme in each assay from 1 to 5000 μg. The structure of the NAD+complex of NMNATase has been determined by the MAD method using selenium as the anomalous scatterer. The resulting electron density is of high quality except for the loop consisting of residues 124–129, in which the main chain density is continuous but increased mobility has compromised the clarity of side chain densities. In addition, 12 C-terminal and 3 N-terminal amino acids are not visible in the electron density map. The final model contains 167 amino acids (residues 4–171), with Pro-14 in a cis-conformation; 119 water molecules; one molecule of NAD+; 1 sodium; and 1 sulfate ion (Fig. 2 A). Refinement at 1.9 Å resolution resulted in an R cryst of 0.212 and an R free of 0.242. According to PROCHECK (20Laskowski R.A. MacArthur M.W. Moss D.S. Thornton J.M. J. Appl. Crystallogr. 1993; 26: 283-291Crossref Google Scholar), 92% of the residues are in the most favored regions, and no residue is in the disallowed regions of the Ramachandran plot. The structure of the NMN+ complex of H19A NMNATase was determined by molecular replacement techniques. Its electron density map is of good quality except for, again, the loop including residues 124 and 129, for which no continuous electron density is observed at all. The final model of this complex contains 161 amino acids (residues 4–123 and 130–170), with Pro-14 still adopting acis-conformation; 10 water molecules; 1 molecule of NMN+; and 1 sulfate ion. Refinement at 2.5 Å resolution gave an R cryst of 0.236 and anR free of 0.294. 94.1% of the residues are in the most favored regions, and no residue is in the disallowed regions. Consistent with gel filtration studies, which for M. thermoautotrophicum NMNATase point to a hexamer as the functional unit in solution (data not shown), the 322 symmetry of NMNATase crystals combines the single monomers occupying each asymmetric unit into a hexameric arrangement (Fig.2 B). Each hexamer is created by two trimers rotated by 180° and layered on top of each other, with overall dimensions of approximately 80 × 80 × 60 Å. Each subunit consists of two domains (Fig. 2 A), the first of which is the dinucleotide-binding domain and comprises residues 4–130 with a topological arrangement of alternating β-strands and α-helices. The twisted β-sheet at the core of the subunit consists of five parallel β-strands with topology 3-2-1-4-5 (Fig. 2 A). The second domain (residues 131–170) is made up from three α-helices (helices 5–7) and is the major contributor to intratrimer subunit interactions. The intratrimer interactions are almost exclusively electrostatic and occur preferentially between helix 5 of subunit A and helix 7 of subunit B. Salt bridges are formed by Arg-110A and Glu-164B; Glu-114A with His-44B and Arg-165B; and Gln-109A with His-168B. These contacts are repeated between subunits B and C as well as subunits C and A (Fig. 2 B). In each case, 1457 Å2 of a total of 16,938 Å2 of molecular surface are buried upon oligomerization. In contrast to the intratrimer interactions, the dominant intertrimer contacts are nonpolar and hydrophobic. The side chains of β-strand 3 (Ile-75, Ile-76, and Val-78) of subunit A pack against their counterparts of β-strand 3 of subunit D with the same interactions repeated between subunits B and E, as well as C and F. In each case, 2796 Å2 of a total of 15,609 Å2 of molecular surface are buried (Fig. 2 B). The active site is located in a deep cleft facing a narrow channel running along the 3-fold symmetry axis of the hexamer. The site is open to solvent, possibly reflecting its readiness to release the product NAD+, which, together with a sulfate ion, could easily be identified in the electron density after the first round of refinement(Fig. 3A). As no NAD+ was added during enzyme purification and crystallization, thermophilic NMNATase must have trapped its product. NAD+ binds in an extended conformation with its adenine ring adopting an anti orientation and both the adenylyl and the nicotinamide ribose rings showing 3′-endo puckering (Fig.3 C). In contrast, the adenylyl ribose ring seems to be in the 2′-endo conformation in the M. jannaschii NMNATase structure (9D'Angelo I. Raffaelli N. Dabusti V. Lorenzi T. Magni G. Rizzi M. Struct. Fold. Des. 2000; 8: 993-1004Abstract Full Text Full Text PDF Scopus (67) Google Scholar). The exocyclic nitrogen of adenine is H-bonded to the main chain carbonyls of Phe-125 and Tyr-130, two aromatic amino acids the side chains of which interact closely. N1 binds to the main chain amide of Phe-125 and N7 to a water molecule. The adenylyl ribose forms an H-bond (3.1 Å long) between its 3′-hydroxyl and the backbone amide of Gly-104, in contrast to what has been found in NAD(P)+-dependent dehydrogenases, in which the adenylyl ribose ring is held in place by a conserved aspartate located at the C terminus of the second β-strand (21Lesk A.M. Curr. Opin. Struct. Biol. 1995; 5: 775-783Crossref PubMed Scopus (203) Google Scholar). In NMNATase, it is the nicotinamide ribose in which one finds H-bonds between its 2′-hydroxyl and the carboxyl side chain of Asp-80; the 3′-hydroxyl interacts with the side chain of Ser-39. The nicotinamide ring stacks with the aromatic ring of Trp-87 and its amide substituent links to the main chain of Ile-81, the oxygen to its amino and the nitrogen to its carbonyl atoms. In addition, the side chain of Asn-84 holds the amide NH2 of the substituent in place (Fig. 3, C andD). As the enzyme has to accept both NMN+ and NaMN+ as substrates, an amide side chain is ideal for this purpose. A single bond rotation will provide an H-bonding partner for nitrogen as well as oxygen atoms. A flip of the Ile-81/Glu-82 peptide bond could easily provide the proper interaction for the second oxygen atom in the carboxylate of NaMN+. Such a change in backbone conformation would not be difficult to achieve as this region of the chain runs along the surface of the protein molecule and seems unconstrained. The "NMN-phosphate" in the pyrophosphate linkage forms H-bonds to Asn-105. The oxygen atom bridging the two phosphorous atoms of the pyrophosphate group contacts the main chain amide of Arg-11. The "AMP-phosphate" binds to His-19, the main chain of Met-12, two water molecules and a rather high (3ς) spherical electron density that we interpret as a Na+ ion. This Na+ ion could contribute to balancing the negative charges of the pyrophosphate and of another electron-dense feature, the center of which is located 5.3 Å from the phosphorous atom of the AMP-phosphate; the shape and position of this feature indicate a sulfate ion, an excellent mimic of a phosphate group. Both sodium and sulfate ions could have been provided by the crystallization buffer (0.5 m NaCl, 1.5 m Li2SO4). The two waters mentioned above and the metal ion are located so they can bridge oxygen atoms from the AMP-phosphate and the SO 42– ion. The sulfate oxygens are bound to the guanidinium groups of Arg-11 and Arg-136 (Fig. 3,C and D), two residues absolutely conserved among the known archaeal and bacterial NMNATases (Fig. 1 B), and also to the guanidinium group of Arg-47 and the backbone of Thr-133. The interaction with Arg-47 seems to be accidental as the corresponding residue in the M. jannaschii structure is a glutamate (9D'Angelo I. Raffaelli N. Dabusti V. Lorenzi T. Magni G. Rizzi M. Struct. Fold. Des. 2000; 8: 993-1004Abstract Full Text Full Text PDF Scopus (67) Google Scholar). The location of the sulfate ion is consistent with it occupying the binding site of the γ-phosphate of the substrate ATP and is reminiscent of the sulfate ion bound at the γ-phosphate position in the active site of glutaminyl tRNA synthetase complexed with AMP (22Perona J.J. Rould M.A. Steitz T.A. Biochemistry. 1993; 32: 8758-8771Crossref PubMed Scopus (183) Google Scholar). This assignment gains further credibility by interpreting the figures portraying the binding of Mg2+-ATP to M. jannaschii NMNATase (9D'Angelo I. Raffaelli N. Dabusti V. Lorenzi T. Magni G. Rizzi M. Struct. Fold. Des. 2000; 8: 993-1004Abstract Full Text Full Text PDF Scopus (67) Google Scholar). Analysis of the structure of NMNATase using the program DALI (23Holm L. Sander C. J. Mol. Biol. 1993; 233: 123-138Crossref PubMed Scopus (3566) Google Scholar) identified several proteins all of them belonging to the nucleotidyltransferase superfamily of dinucleotide-binding fold containing α/β phosphodiesterases (24Izard T. Geerlof A. EMBO J. 1999; 18: 2021-2030Crossref PubMed Scopus (102) Google Scholar). Presently known members of this family are CTP:glycerol-3-phosphate cytidyltransferase (CTP:G3PCase) (25Weber C.H. Park Y.S. Sanker S. Kent C. Ludwig M.L. Struct. Fold Des. 1999; 7: 1113-1124Abstract Full Text Full Text PDF Scopus (92) Google Scholar), glutaminyl tRNA synthetase (26Rould M.A. Perona J.J. Soll D Steitz T.A. Science. 1992; 246: 1135-1142Crossref Scopus (803) Google Scholar), tyrosyl tRNA synthetase (27Brick P. Bhat T.N. Blow D.M. J. Mol. Biol. 1989; 208: 83-98Crossref PubMed Scopus (348) Google Scholar) and phosphopantetheine adenylyltransferase (PPATase) (24Izard T. Geerlof A. EMBO J. 1999; 18: 2021-2030Crossref PubMed Scopus (102) Google Scholar). Their overall structures are remarkably similar (r.m.s.d. of 116, 137, 124, and 143 Cα atoms, equal to 2.5, 3.5, 3.9, and 2.3 Å, respectively). PPATase not only closely resembles the overall fold of NMNATase but even forms a hexamer as the functional unit. Nevertheless, the arrangement of subunits relative to each other is quite different. All of these related proteins contain a nucleotide-binding domain, present the active site sequence motif (T/H)XGH, and catalyze a nucleotidyltransfer reaction that is similar to that of NMNATase. This reaction involves the attack of a nucleophilic group of one substrate at the α-phosphate of the nucleoside triphosphate (the second substrate), thereby forming a new phosphodiester bond and releasing a pyrophosphate molecule. Originally, identification of NAD+ in the active site cleft and the DALI results indicating nucleotidyltransferase function led us to propose that protein MT0150, annotated as "conserved" in theM. thermoautotrophicum genome sequence data base (28Smith D.R. Doucette-Stamm L.A. Deloughery C. Lee H. Dubois J. Aldredge T. Bashirzadeh R. Blakely D. Cook R. Gilbert K. Harrison D. Hoang L. Keagle P. Lumm W. Pothier B. Qiu D. Spadafora R. Vicaire R. Wang Y. Wierzbowski J. Gibson R. Jiwani N. Caruso A. Bush D. Safer H. Patwell D. Prabhakar S. McDougall S. Shimer G. Goyal A. Pietrokovski S. Church G. Daniels C. Mao J. Rice P. Nölling J. Reeve J.N. J. Bacteriol. 1997; 179: 7135-7155Crossref PubMed Scopus (1040) Google Scholar), was in fact an NMNATase. Preliminary enzymatic assays confirmed that the enzyme catalyzed the biosynthesis of NAD+ from NMN+ and ATP (Fig.4B). The validity of this assignment was established when a literature search revealed that the homologous enzyme from M. jannaschii had NMNATase activity (4Raffaelli N. Pisani F.M. Lorenzi T. Emanuelli M. Amici A. Ruggieri S. Magni G. J. Bacteriol. 1997; 179: 7718-7723Crossref PubMed Google Scholar, 6Raffaelli N. Lorenzi T. Amici A. Emanuelli M. Ruggieri S. Magni G. FEBS Lett. 1999; 444: 222-226Crossref PubMed Scopus (42) Google Scholar). In addition to the common nucleotide-binding fold, all of these enzymes contain an active site sequence motif, (T/H)XGH, that is absolutely conserved in all known archaeal and bacterial NMNATases (16HRGH19 in M. thermoautotrophicumNMNATase; Fig. 1 B). This active site sequence motif has also been identified in other nucleotide binding enzymes catalyzing adenylyltransferase reactions such as the ATP sulfurylase domain of human phosphoadenosine phosphosulfate synthase (425HNGH428), A. thaliana ATP sulfurylase, and E. coli flavin adenine dinucleotide synthetase (29HRGH32) (29Venkatachalam K.V. Fuda H. Koonin E.V. Strott C.A. J. Biol. Chem. 1999; 274 (Shao, Y. (1997) Science 277, 1453–1474): 2601-2604Abstract Full Text Full Text PDF PubMed Scopus (34) Google Scholar, 30Hatzfeld Y. Lee S. Lee M. Leustek T. Saito K. Gene. 2000; 248: 51-58Crossref PubMed Scopus (60) Google Scholar, 31Blattner F.R. Plunkett G. Bloch C.A. Perna N.T. Burland V. Riley M. Collado-Vides J. Glasner J.D. Rode C.K. Mayhew G.F. Gregor J. Davis N.W. Kirkpatrick H.A. Goeden M.A. Rose D.J. Mau B. Shao Y. Science. 1997; 277: 1453-1474Crossref PubMed Scopus (6050) Google Scholar). 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