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CRYSTAL STRUCTURE OF BACULOVIRUS RNA TRIPHOSPHATASE (BVP) COMPLEXED WITH PHOSPHATE*

Anita Changela1, Alexandra Martins2, Stewart Shuman2, and Alfonso Mondragón1

From the 1Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205 Tech Drive, Evanston, Illinois 60208 and 2Molecular Biology Program, Sloan-

Kettering Institute, New York, New York 10021 Running Title: Crystal structure of baculovirus phosphatase

Address correspondence to: Alfonso Mondragón, Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, 2205 Tech Drive, Evanston, Illinois 60208, Tel: 847-491-7726, Fax: 847-467-6489, e-mail: a-mondragon@northwestern.edu

Baculovirus RNA 5’-triphosphatase (BVP) exemplifies a family of RNA-specific cysteine phosphatases that includes the RNA triphosphatase domains of metazoan and plant mRNA capping enzymes. Here we report the crystal structure of BVP in a phosphate-bound state at 1.5 Å resolution. BVP adopts the characteristic cysteine-phosphatase α/β fold and binds two phosphate ions in the active site region, one of which is proposed to mimic the phosphate of the product complex after hydrolysis of the covalent phosphoenzyme intermediate. The crystal structure highlights the role of backbone amides and side chains of the P-loop motif 118HCTHGxNRT126 in binding the cleavable phosphate and stabilizing the transition state. Comparison of the BVP structure to the apoenzyme of mammalian RNA triphosphatase reveals a concerted movement of the Arg125 side chain (to engage the phosphate directly) and closure of an associated surface loop over the phosphate in the active site. The structure highlights a direct catalytic role of Asn124, which is the signature P-loop residue of the RNA triphosphatase family and a likely determinant of the specificity of BVP for hydrolysis of phosphoanhydride linkages.

mRNA 5’ cap formation is initiated by hydrolysis of the γ phosphate of 5’-triphosphate-terminated pre-mRNA. The resulting 5’ diphosphate end is then capped by transfer of GMP from GTP to form an inverted terminal dinucleotide structure, G(5’)ppp(5’)N. The first reaction is catalyzed by RNA 5’-triphosphatase and the second by GTP:RNA guanylyltransferase (reviewed in

(1,2)). In metazoans and plants, the triphosphatase and guanylyltransferase activities reside in a single polypeptide composed of an N-terminal triphosphatase domain fused to a C-terminal guanylyltransferase domain. The metazoan and plant RNA triphosphatases belong to the cysteine phosphatase superfamily (3), which includes phosphoprotein phosphatases (4) and phosphoinositide phosphatases (5). Cysteine phosphatase-type RNA triphosphatases act via a two-step mechanism entailing attack by a cysteine thiolate nucleophile of the enzyme on the γ phosphate of triphosphate-terminated RNA to form a cysteinyl-phosphoenzyme intermediate, which is then hydrolyzed to liberate inorganic phosphate. The active site cysteine is located within a signature P-loop motif, HCTHGxNRT. We previously reported the crystal structure of the RNA triphosphatase domain of mouse capping enzyme Mce1 at 1.65 Å resolution (3). Mce1 adopts a globular fold consisting of a five-stranded parallel β-sheet flanked by α-helices on both sides. The active site cysteine is located at the bottom of a deep, positively charged pocket formed by essential amino acids that line the pocket walls and surface rim. Structural, biochemical, and mutational results showed that despite sharing a HCxxxxxR(S/T) P-loop motif, a phosphoenzyme intermediate, and a core α/βfold with other cysteine phosphatases, the mechanism of phosphoanhydride cleavage by Mce1 and its baculovirus homolog BVP differs from that used by phosphoprotein phosphatases to hydrolyze phosphomonoesters. The key distinction is the absence of a carboxylate general acid catalyst

JBC Papers in Press. Published on February 15, 2005 as Manuscript M500885200

Copyright 2005 by The American Society for Biochemistry and Molecular Biology, Inc.

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in the RNA-specific triphosphatases. Residues conserved uniquely among the RNA phosphatase subfamily are important for cap formation in vivo and have been proposed to play a role in substrate recognition. However, the structure of the Mce1 triphosphatase apoenzyme provided no direct information regarding the nature of the atomic contacts involved. Baculovirus phosphatase (BVP) is a 168-aa protein encoded by Autographa californicanucleopolyhedrovirus (6,7). BVP displays primary structure similarity to the N-terminal RNA triphosphatase domain of Mce1. BVP differs from the cellular capping enzyme in several respects. First, it is a monofunctional triphosphatase that is not covalently linked to a guanylyltransferase domain. Second, unlike the cellular capping enzyme, which hydrolyzes only the γ phosphate of a triphosphate-terminated substrate, BVP can also hydrolyze the β phosphate of either diphosphate-terminated RNA or free NDPs (6,7). Monofunctional RNA phosphatases are not confined to baculovirus; the monofunctional human phosphatase PIR1 is a homolog of BVP that also displays RNA triphosphatase and diphosphatase activities (8). Additionally, PIR1-like proteins are present in many metazoan proteomes, but their functions are not known (8). BVP activity also embraces the hydrolysis of inorganic tripolyphosphate and pyrophosphate (9). Despite its broad specificity for hydrolysis of phosphoanhydrides in vitro, BVP can act as an RNA triphosphatase in the cap synthetic pathway in vivo in yeast cells (9). The availability of biochemical and genetic readouts of BVP activity has prompted an extensive mutational analysis of BVP, whereby amino acids essential for triphosphatase activity in vitro and in vivowere identified initially by alanine-scanning (10), after which structure-activity relationships at the essential residues were determined via conservative substitutions (11). What is lacking is a structural explanation for the mutational effects based on atomic interactions with substrates, intermediates or products, as well as a structural framework for

the unique specificity of BVP for phosphoanhydrides and the broad specificity of BVP for triphosphate and diphosphate ends versus the more stringent specificity of the bifunctional capping enzymes for triphosphate ends. Here we report the crystal structure of BVP in a phosphate-bound state at 1.5 Å resolution, which provides new insights into these issues.

Materials and Methods Purification and crystallization of BVP. BVP was produced as an N-terminal His10-BVPfusion in E. coli BL21(DE3)pLysS cells and purified by Ni-agarose chromatography as described previously (11). For crystallization, His-tagged BVP was purified further by gel filtration chromatography, dialyzed into 50 mM Tris-HCl (pH 8.0), 0.5 M NaCl, 1 mM EDTA, and 1 mM DTT, and concentrated to 3-4 mg/ml. Attempts to remove the His-tag prior to crystallization resulted in protein precipitation and hence the crystallization trials were done using the His-tagged protein. Initial crystallization trials of His-tagged BVP at 22°C in hanging drops equilibrated against 2.0 M sodium/potassium phosphate (pH 7.0) and 0.1 M sodium acetate (pH 4.5) yielded showers of small needle-like crystals. In order to improve crystal quality, the affinity tag was cleaved off during crystallization by including 0.1 M guanidine-HCl and trypsin (1:5000 molar ratio of trypsin to BVP) in the original mother liquor prior to setting up the crystallization experiment. In this manner, strongly diffracting crystals of the untagged form of BVP appeared within 1 week and grew as large needles of ~0.06 x 0.06 x 0.4 mm3.Data collection and structure determination.Prior to data collection, crystals were transferred in a single step to crystallization solution supplemented with 25% glycerol for 1-2 minutes and then flash cooled in liquid nitrogen. All data were collected at 100° K using a MAR-CCD detector at beamline 5ID-B at the Advanced Photon Source. Data were integrated using XDS (12) and scaled with SCALA (13). The crystals belong to space group P21 with unit cell dimensions of a=40.9 Å, b=74.1 Å, c=105.2 Å, β=92.4°, and there

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are three molecules (molecules A, B, and C) in the asymmetric unit. The structure was solved by molecular replacement using the program AMoRe (13). The crystal structure of the RNA triphosphatase domain of the mouse mRNA capping enzyme (Mce1) (3), with non-conserved loop and C-terminal regions removed, was used as the search model. A clear solution for the positions of all three monomers was found with an R-factor of 47.5% and a correlation coefficient of 0.388. An initial model was built using an electron density map calculated to 2.5 Å resolution with improved phases obtained from prime-and-switch phasing in RESOLVE (14). Manual model building was carried out in O (15)and alternated with cycles of simulated annealing, positional, and individual temperature factor refinement in CNS (16). Two strong Fo-Fc difference density peaks for phosphate ions were found at the active site of each monomer. In molecule B, additional electron density adjacent to one of the phosphates (Site 2) was assigned as a second phosphate with low occupancy. It is not clear whether these two phosphates are part of a diphosphate molecule or not, and they were treated as two separate ions. Once most of the model had been built, the resolution was extended using data to 1.5 Å resolution obtained from a crystal grown in the presence of 5 mM ATP. No ATP was observed in the electron density maps. Subsequent refinement was performed with REFMAC5 (17). Molecules A and B are intact and contain residues 1-168 whereas in molecule C residues 1 and 34-41 are disordered. Additionally, in molecule B there is electron density for an N-terminal histidine residue encoded by the expression vector. Molecules A and B exhibit lower average temperature factors (average B-factor ~12-14 Å2) than molecule C (average B-factor=22 Å2). The final model has an Rfree=19.4% and R-factor=16.5%, includes 576 water molecules and 7 phosphate ions. 99.5% of all residues are found within the most favored or allowed regions in the Ramachandran plot with only 2 residues (Gln88 in the A and C molecules) in disallowed regions. Data collection and

refinement statistics are summarized in Table 1. Figures were generated using MOLSCRIPT (18), RASTER3D (19), GRASP (20), and SETOR and SETORPLOT (21). Coordinates and structure factors for BVP have been deposited in the RCSB Protein Data Bank under the accession code 1YN9. Purification of native BVP. In order to produce soluble, His-tag free BVP, the BVP gene was amplified by PCR from pET16b-BVP plasmid (6) using a sense primer designed to introduce a BamHI restriction site immediately upstream of the start codon. The PCR product was digested with BamHI and inserted into the BamHI site of the vector pET28-His-Smt3 to generate the plasmid pET-His-Smt3-BVP, which encodes BVP fused to an N-terminal tag consisting of a His6 leader peptide followed by the 98-aa S. cerevisiaeSmt3 protein and a single serine. (Smt3 is the yeast ortholog of the small ubiquitin-like modifier SUMO). The pET-His-Smt3-BVP plasmid was transformed into E. coliBL21(DE3)pLysS. A 1-liter bacterial culture amplified from a single transformant was grown at 37°C in Luria-Bertani medium containing 50 µg/ml kanamycin and 30 µg/ml chloramphenicol until A600 reached ~0.6. Expression was induced by adding IPTG to a final concentration of 0.1 mM. The culture was then incubated at 30°C for 4 h with continuous shaking. Cells were harvested by centrifugation, and the recombinant His6-Smt3-BVP protein was purified from the soluble bacterial extract by Ni-agarose chromatography as described previously (11). The 33-kDa His6-Smt3-BVP polypeptide was recovered in the 50 mM and 200 mM imidazole eluate fractions; the yield was approximately 18 mg. The enzyme preparation was dialyzed against buffer B (50 mM Tris-HCl, pH 8.0, 50 mM NaCl, 10% glycerol). The His6-Smt3 tag was removed by proteolytic cleavage with the S. cerevisiaeSmt3-specific protease Ulp1 (22). Briefly, a reaction mixture (560 µl) containing 50 mM Tris-HCl (pH 8.0), 1 mM DTT, 350 mM NaCl, 9% glycerol, 1.5 mg of His6-Smt3-BVP, and 1.5 µg of His6Ulp1-N 403 was incubated on ice for 30 min. To separate the cleaved His-

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Smt3 tag from native BVP, the digest was applied to a 0.2-ml column of nickel-NTA resin (Qiagen) that had been equilibrated with buffer C (50 mM Tris-HCl, pH 8.0, 300 mM NaCl, 10% glycerol). The 19 kDa native BVP protein was recovered in the flow-through and wash fractions, whereas His-Smt3 was retained on the resin and was eluted with 0.2 M imidazole. BVP concentrations were determined by SDS-PAGE analysis of serial dilutions of the BVP preparations in parallel with serial dilutions of a BSA standard. The gels were stained with Coomassie Blue, and the staining intensities of the BVP and BSA polypeptides were quantified using a digital imaging and analysis system from Alpha Innotech Corporation.Velocity sedimentation. Aliquots (100 µg) of the His10-BVP and native BVP preparations were applied to 5.0 ml 15–30% glycerol gradients containing 50 mM Tris-HCl (pH 8.0), 0.3 M NaCl, 1 mM EDTA, 1 mM DTT, 0.1% Triton X-100. The gradients were centrifuged in a SW50i rotor at 50,000 rpm for 18 h at 4°C. Protein standards catalase (75 µg), BSA (75 µg) and cytochrome c (75 µg) were sedimented in a parallel gradient. Fractions (21 drops each) were collected from the bottoms of the tubes. Aliquots (10 µl) of even numbered gradient fractions were analyzed by SDS–PAGE. Aliquots of the fractions were assayed for triphosphatase activity as specified in the figure legend.

Results

Structure determination of baculovirus phosphatase. Initial crystallization trials of His-tagged BVP yielded small crystals of the fusion protein that were unsuitable for diffraction studies. In order to improve crystal quality, attempts were made to remove the His10 tag from BVP in solution using various proteases, but all trials resulted in protein precipitation. However, strongly diffracting crystals could be grown by including small amounts of trypsin in the original mother liquor prior to setting up the crystallization experiment. SDS-PAGE analysis of crystals grown in the

presence of trypsin suggested that most or all of the affinity tag had been cleaved off during crystallization, which was later confirmed by the crystal structure. Due to solubility problems and as we could obtain excellent crystals using the above described procedure, no attempts to crystallize BVP using a tag-free version of the protein were done. The structure of BVP was determined by molecular replacement using the structure of the RNA triphosphatase domain of mammalian capping enzyme (3) as a search model. The final model containing three BVP molecules (A, B, and C) in the asymmetric unit was refined to 1.5 Å resolution with an Rfree=19.4% and R-factor=16.5% (Table 1). All three monomers are nearly identical in conformation, with root mean square deviation (rmsd) values ranging between 0.3 Å to 0.4 Å for all Cα atoms. Molecules A and B are complete (residues 1-168 are visualized; Fig. 1A) whereas in molecule C the N-terminal methionine and surface loop residues 34-41 are disordered. Within the asymmetric unit, molecules B and C are each related to molecule A by local two-fold symmetry. The occurrence of 3 protomers in the asymmetric unit of the crystal derived from BVP that was trypsinized in situ to remove the tag prompted us to compare the quaternary structures of His10-tagged BVP and native tag-free BVP. Zonal velocity sedimentation analysis of recombinant His10-tagged BVP revealed two distinct populations: a slow migrating species sedimenting between BSA (68 kDa) and cytochrome c (13 kDa) that we presume is a BVP monomer and a fast migrating species that sediments on the light side of catalase (248 kDa) that could represent an octameric or higher order oligomeric complex (Fig. 2A). Both components possessed ATP phosphohydrolase activity (Fig. 2A). To obtain tag-free BVP, we first produced BVP in bacteria as a His6-Smt3 fusion protein, purified the fusion protein by Ni-agarose chromatography, removed the His10-Smt3 tag by digesting the protein with the Smt3-specific protease Ulp1, and then isolated the “native” BVP protein free of His6-Smt3 by passing the digest over a Ni-agarose column and recovering native BVP in the

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flow-through fraction. Native BVP sedimented as a single catalytically active monomeric component, with no detectable oligomeric forms (Fig. 2B). We surmise that: (i) native BVP is monomeric; (ii) higher order oligomerization is induced by the His10-tag:and (iii) the arrangement of BVP molecules in the crystal structure does not reflect a biologically relevant oligomerization state. Although the structure presented here was derived from data obtained from a BVP crystal grown in the presence of ATP, no electron density was observed for an ATP molecule. Instead, strong density was observed in the Fo-Fc electron density map for two well-ordered phosphate ions in the active site region of each monomer (Fig. 3A), which were likely derived from the 2 M Na/K phosphate crystallization solution. Although the catalytic pockets in molecules A and B were blocked due to crystal packing interactions, the active site in molecule C was solvent-accessible (Fig. 4B-D). Extensive cocrystallization trials and crystal soaks in the presence of excess ATP and other substrate analogues failed to replace the bound phosphates with alternative ligands.

Overview of the BVP structure - a cysteine phosphatase fold. BVP is a single domain protein consisting of a central, twisted, five-stranded β-sheet surrounded by six α-helices, two on one side and four on the other (Fig. 1A). The conserved phosphate-binding P-loop motif connects the β5 strand and the α6 helix. The compact α/β domain structure adopted by BVP is nearly identical in topology to the RNA triphosphatase domain of Mce1 (Fig. 1B) and resembles the conserved catalytic core found in other cysteine phosphatases. Structural comparisons using the DALI server (23)indicate that BVP shares the most similarity to Mce1, the phosphoinositide phosphatase PTEN (24), and several dual-specificity protein phosphatases, including the kinase-associated phosphatase KAP (25), human Cdc14B (26), and VHR (27). The conformation of the P-loop residues 118HCTHGINRT126 and the overall active site environment are identical in all three BVP monomers. The Cys119 nucleophile extends

from the bottom of the substrate-binding pocket towards one of the bound phosphate ions (Site 1) (Fig. 3B). Other conserved P-loop residues and residues from neighboring surface loops lend an overall positive surface potential, providing an attractive docking site for either a diphosphate or triphosphate substrate. Positively charged, shallow grooves extending from the catalytic pocket indicate additional regions possibly involved in substrate binding (Fig. 4A). In other cysteine phosphatases, the depth of the active site pocket is a critical factor in defining substrate specificity (24,27,28). It was postulated that BVP would use a shallower pocket to act on diphosphate-terminated substrates that cannot be accommodated by the deep pocket in Mce1 (3). However, the depth of the active site pocket in BVP is ~8 Å, making it very similar to Mce1 and suggesting that other factors dictate substrate specificity in the RNA-specific cysteine phosphatase family (see below).

Two phosphate-binding sites. Onephosphate ion (Site 1) binds deep in the catalytic pocket and is coordinated by conserved P-loop residues and main-chain amide groups (Fig. 3B). The phosphate is directly above the catalytic Cys119 at a sulfur to phosphorous distance of ~3.8 Å, indicative of a non-covalent interaction. The phosphate in Site 1 exemplifies the binding mode expected for the terminal phosphate of an RNA substrate prior to formation of the covalent cysteinyl-phosphoenzyme intermediate or a product complex generated after hydrolysis of the phosphoenzyme. The invariant P-loop residue, Arg125, makes a bidentate contact with two of the phosphate oxygens at Site 1, consistent with the role proposed for this residue in substrate binding and transition-state stabilization (11). The key contribution of the bidentate contact is underscored by previous findings that mutation of Arg125 to alanine abolished BVP triphosphatase activity in vitro and in vivo and conservative substitution by lysine did not rescue function (11). P-loop residues His121 and Asn124 make additional hydrogen bonds to the Site 1 phosphate. The backbone amide

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and Nδ of His121 coordinate the same phosphate oxygen contacted by Arg125 NH2 (Fig. 3B). The phosphate contact to the His121 side chains is essential, insofar as alanine substitution abolished triphosphatase function in vitro and in vivo and conservative changes to glutamine or asparagine did not restore activity. (10) The hydrogen bond of Asn124 Nδ to the phosphate is also essential; mutations N124A, N124D, and N124Q abolished BVP triphosphatase activity in vivoand in vitro (10). The second phosphate ion (Site 2) is adjacent to the active site pocket in a nearby shallow cleft created primarily by residues from the loop connecting the β3 strand and α4helix (Fig. 1A and 4). This loop contains a 61LTNTSKYY68 motif that is conserved in other RNA-specific cysteine phosphatases. The Site 2 phosphate receives hydrogen bonds from Asn63 and Tyr67 of the β3-α4 loop and from P-loop residues Thr120 and Arg125 (Fig. 3B). The Site 2 phosphate is located ~7.7 Å from the Site 1 phosphate, a position inconsistent with that expected for the βphosphate of a triphosphate-terminated substrate or the α phosphate of a diphosphate-terminated substrate. Previous findings that alanine mutations of Tyr67 and Thr120 reduced triphosphatase activity in vitro by factors of about 5 and 10 (10,11), respectively, prompted us to query whether contact to the Site 2 phosphate might mimic those to the αphosphate of a triphosphate-terminated substrate, in which case the Y67A and T120A mutations might differentially affect triphosphatase activity, but not hydrolysis of a diphosphate substrate. Thus, we compared the ATPase and ADPase activities of wild-type BVP and the two alanine mutants (Fig. 5). As noted previously (9), wild-type BVP displayed higher specific activity in ADP hydrolysis than in ATP hydrolysis. The Y67A and T120A enzymes were 21% and 9% as active as wild-type with ATP and 14% and 15% as active as wild-type with ADP (Fig. 5). These results indicate that loss of the Tyr67 and Thr120 side chains does not differentially affect triphosphatase versus diphosphatase activity, i.e., neither side chain is likely to

make functionally significant contacts with the α phosphate of a 5’ triphosphate end. An alternative scenario, discussed below, is that the Site 2 phosphate mimics the position of the leaving 5’ phosphate moiety after cleavage of the phosphoanhydride linkage and formation of the cysteinyl phosphoenzyme intermediate.

Conformational differences between phosphate-bound BVP and unliganded Mce1. The BVP structure superimposes on the RNA triphosphatase domain of Mce1 with an rmsd of 1.3 Å for Cα atoms of 154 residues. The architecture of the core elements in both proteins is identical; most of the differences are localized to variable loop regions (Fig. 1B). A notable change in conformation elicited by phosphate binding involves the loop leading from the β4 strand into the α5helix (residues 82-91 in BVP). The β4-α5loop in BVP and Mce1 is structurally (though not functionally) analogous to the “WPD loop” or general acid loop in phosphoprotein phosphatases. In those enzymes, which specifically cleave phosphomonoesters, substrate binding triggers movement of the WPD loop toward the active site by as much as 8 Å, so that the loop closes over the substrate-binding pocket. This displacement brings a conserved aspartic acid of the loop into the substrate-binding site, where the Asp then acts as a proton donor to facilitate expulsion of the hydroxy amino acid leaving group. Comparison of the equivalent loop in the unliganded form of Mce1 and the phosphate-bound BVP reveals a similar displacement of the loop towards the active site by ~4 Å (Fig. 1B). Despite neighboring crystal-packing interactions in molecules A and B, the overall loop displacement is identical in all three monomers, suggesting that the conformational difference is not an artifact of crystal packing. Movement of the β4-α5 loop towards the active site in the BVP complex with phosphate effectively narrows the opening of the catalytic pocket and likely stabilizes the bound substrate. Among the RNA-specific members of the cysteine phosphatase family, there is no conserved residue on this loop that could serve as a general acid. Indeed, cleavage of

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phosphoanhydrides by the RNA-specific cysteine phosphatases does not depend on a general acid catalyst because of the low pKa of the leaving group (3,10). Nonetheless, the structural alignment of Mce1 and BVP reveals that the mobile loop contains a polar residue, such as Gln88 in BVP and His94 in Mce1, that might be involved in substrate recognition or binding (Fig. 6). Whereas movement of the loop brings Gln88 closer to the catalytic site in BVP, Gln88 is not optimally oriented towards the active site for substrate binding in any of the monomers. Mutational analysis showed that neither Gln88 in BVP nor the equivalent His94 in Mce1 are important for triphosphatase activity (3,11). We favor the idea that loop closure and opening play a topological role in coordinating substrate binding and product release, in a manner that does not rely on Gln88. The most significant difference between the Mce1 apoenzyme and the BVP phosphate complex is the switch in the position of the catalytic arginine side chain of the P-loop (Fig. 6). In the Mce1 apoenzyme, the arginine is held away from the phosphate-binding site by virtue of hydrogen bonds between its terminal guanidinium nitrogens and the backbone carbonyls of Leu67 in the β3-α4loop and Lys92 in the β4-α5 loop. This location of the arginine allows ingress of the phosphate to the exposed cysteine at the base of the active site pocket. In BVP, the arginine (Arg125) has rotated about the Cγ–Cδ bond allowing Nε and a terminal guanidinium nitrogen to coordinate the Site 1 phosphate. Additionally, Arg125 in BVP is involved in a hydrogen bond to the backbone carbonyl of Val85 in the β4-α5 loop; the equivalent position is Cys91 in Mce1, which is 1 residue upstream of the Lys92 residue with which the arginine interacts in the Mce1 apoenzyme. Thus, the contact between the P-loop arginine and the β4-α5 loop appears to be remodeled as the arginine and the β4-α5 loop move in concert toward the bound phosphate (Fig. 6). During this conformational switch, the backbone contact of the arginine to the leucine carbonyl oxygen of the β3-α4 loop is broken

and replaced by the hydrogen bond of the arginine to the Site 2 phosphate.

Discussion

The BVP structure likely exemplifies a product complex. We envision that the structure of BVP with a phosphate ion bound at Site 1 captures the state of the product complex subsequent to hydrolysis of the covalent phosphoenzyme intermediate. The Site 1 phosphate is oriented so that the cysteine sulfur is apical to the phosphate oxygen contacted by the Asn124 side chain (Fig. 3B). Said oxygen, in the product complex, would correspond to the nucleophilic water that attacks the phosphoenzyme in the second step of the BVP reaction mechanism. Thus, the structure suggests that at least one critical role of Asn124 would be to orient the water for its attack on the phosphoenzyme. Because mutation of Asn124 to alanine or glutamine abolishes the formation of the BVP phosphoenzyme intermediate (10), we can also invoke an essential role for this side chain in the first step of the reaction pathway, possibly in coordinating the bridging oxygen of the leaving group. Indeed, it is reasonable to think that the Site 1 phosphate resembles the terminal phosphate of the substrate prior to attack by the P-loop cysteine. The P-loop asparagine is invariant in all known RNA 5’ phosphatases that specifically cleave phosphoanhydride linkages. Moreover, the equivalent P-loop position is conspicuously not asparagine in the vast number of cysteine phosphatases that hydrolyze phosphomonoester substrates. For example, the equivalent residue is typically a glycine in protein tyrosine phosphatases (29). We suggest that the P-loop asparagine is the catalytic signature of the RNA 5’ phosphatase family. We envision that the asparagine plays a role loosely analogous to the catalytic aspartate found in other cysteine phosphatase subfamilies (which is missing in RNA 5’ phosphatases) insofar as the BVP asparagine orients the leaving and attacking groups during step 1 and 2; the mechanistic distinction is that BVP catalysis by the P-loop

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asparagine involves hydrogen bonding, not proton donation or removal by a general acid/base. The postulate that the BVP structure reflects a product complex can be extended to include the Site 2 phosphate, which could plausibly occupy the position of the terminal 5’ phosphate of the displaced leaving group, either the β phosphate of ADP formed from ATP or the α phosphate of AMP formed from ADP. As noted above, the Tyr67 and Thr120 side chains that contact the Site 2 phosphate are not responsible for discriminating between triphosphate and diphosphate substrates. Indeed, although alanine changes at these positions are deleterious, conservative mutational analysis indicates that any polar contacts made by Tyr67 or Thr120 to the Site 2 phosphate are not functionally important, i.e., Y67F and T120V mutants retain wild-type triphosphatase activity in vitro and in vivo(10,11). It is conceivable that Thr120 and Tyr67 help form a pocket for the leaving group phosphate, without needing to interact directly with that phosphate. In such a case, weakening polar contacts might even enhance dissociation of the step 1 reaction product, which may explain why T120V is more active than wild-type BVP in vitro (10).

Possible determinants of substrate specificity. The in silico exercise of modeling ATP or ADP into the catalytic pocket so that their terminal phosphates overlap Site 1 highlights candidate substrate-binding determinants. For example, Arg5 and Arg159 (Arg9 and Lys166 in Mce1) are conserved residues that line the rim of the substrate-binding pocket and could interact with the substrate (Fig. 6). Hydrogen bonds between Oδ of Asn124 and the amide groups of Glu158 and Arg159 facilitate optimal positioning of Arg159 for a possible role in substrate binding. Likewise, interactions between Nε of His128 and the carbonyl of Ala4 might orient Arg5 for substrate recognition. Changing BVP Arg159 to alanine reduced triphosphatase in vitro and inactivated BVP function in vivo. An R5A mutant displayed wild-type ATPase activity but reduced ability in phosphoenzyme

formation and a temperature-sensitive phenotype in vivo (11). We anticipated that the BVP structure would reveal why BVP is capable of hydrolyzing triphosphate and diphosphate substrates, whereas Mce1 is specific for triphosphate termini. Although the depth of the active site pocket is an important determinant of substrate specificity in other cysteine phosphatases (24,27,28), comparison of the BVP and Mce1 structures suggest that this may not be the case here. The pocket depth in Mce1 and BVP appear to be very similar. Aside from pocket depth analysis, it is difficult to make a direct comparison of the overall size and shape of the catalytic pockets in Mce1 and BVP, because of the β4-α5 loop movement in BVP. However, an overlay of residues in the catalytic region of BVP and Mce1 (Fig. 6), plus the comparison of primary structures of triphosphatase/diphosphatase enzymes like BVP and PIR1 with the triphosphate-specific cellular capping enzymes with fused triphosphatase and guanylyltransferase domains (Fig. 7), highlights subtle differences that might influence substrate specificity. For example, Glu158 in BVP is conserved as Glu in PIR1, but is a tyrosine in Mce1 (Tyr165) and other bifunctional cellular capping enzymes. The presence of a bulky aromatic group at the rim of the substrate-binding pocket in the capping enzymes might allow binding of a triphosphate group but discriminate against an ADP molecule. A glutamate at this position in BVP would pose little hindrance for binding of a triphosphate or diphosphate-terminated substrate. In Mce1, Tyr165 contacts Arg9 at the pocket rim, yet this interaction is not conserved in BVP. Glu158 in BVP is ~4 Å from Arg5 and instead engages in a hydrogen bond with Tyr9, a residue extending from the coil region leading into the β1 strand. Whereas Tyr9 is conserved in BVP and PIR1, the bifunctional capping enzymes all have a cysteine at the equivalent position. Definitive assessment of the basis for discrimination of triphosphate versus diphosphate ends will ultimately hinge on obtaining cocrystals of BVP with ADP and ATP and of Mce1 with a triphosphate substrate.

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A growing family of putative RNA-specific cysteine phosphatases. Mce1 and BVP are the best characterized members of a distinct branch of the cysteine phosphatase superfamily that has evolved to hydrolyze phosphoanhydride bonds. Although the tertiary structure is conserved between the RNA 5’-phosphatases and other cysteine phosphatases, there is little conservation of primary structure outside of the HCxxxxxR(S/T) P-loop segment that comprises the active site (3). As shown here, there is considerable structural conservation (both primary and tertiary) between Mce1 and BVP that can be extended by sequence alignments to the other members of the RNA phosphatase family, whether they are from cellular or viral sources, and whether they are bifunctional capping enzymes or monofunctional RNA phosphatases (Fig. 7). The present study highlights the P-loop Asn124 as the major structural signature of the active site of RNA 5’ phosphatases. In order to update the membership of this family, we searched the NCBI database for BVP-like proteins, and focused only on those that contained an Asn immediately preceding the P-loop arginine. BVP homologues are present in a subset of baculovirus proteomes, including the viruses of Choristoneura fumiferana, Orgyiapseudotsugata, Epiphyas postvittana, Bombyxmori, Antheraea pernyi, and Autographa californica. BVP is not essential for replication of the AcNPV baculovirus in cell culture (30), but its participation in the capping of viral mRNAs might be overshadowed by the fact that AcNPV and all other baculoviruses encode another RNA triphosphatase catalytic module, which is structurally and mechanistically unrelated to BVP, and is fused to a guanylyltransferase domain to form a bifunctional capping enzyme (named Lef4) (6,31,32). Lef4 is a subunit of the baculovirus RNA polymerase. Although the lef4 gene encoding the baculovirus capping enzyme/polymerase subunit is essential for virus replication, it may not be the case that the triphosphatase activity of the Lef4 subunit is essential if BVP provides a

backup mechanism for catalyzing the RNA triphosphatase step of the capping pathway. In this light, it is notable that we identified new BVP-like proteins encoded by two poxviruses – canarypox virus and Amsactamoorei entomopox virus (AmEPV) – that replicate in widely divergent vertebrate and invertebrate hosts: (Fig. 7). The 157-aa canarypox protein corresponds to a minimized cysteine phoshatase; the 403-aa AmEPV protein contains a unique C-terminal extension (without recognizable functional signatures) appended to an N-terminal cysteine phosphatase module. These two poxvirus phosphatases have the RNA phosphatase-type P-loop and also contain the signature residues that distinguish the monofunctional BVP/PIR1 subgroup from the subgroup of bifunctional cellular capping enzymes (Fig. 7). The sequence signatures suggest that the poxvirus proteins might resemble BVP and PIR1 in acting on triphosphate and diphosphate RNA ends. Identification of BVP-like putative RNA phosphatases encoded by a small subset of known poxvirus genera provides new insight to the diversification of the capping apparatus among eukaryal taxa. All poxviruses encode a polyfunctional capping enzyme that consists of an RNA triphosphatase module fused to a guanylyltransferase (33). As with the baculovirus Lef4 capping enzyme, the triphosphatase module of the omnipresent poxvirus capping enzyme is completely unrelated to the cysteine phosphatase-type RNA triphosphatases. Rather, the poxvirus and baculovirus RNA triphosphatases belong to a metal-dependent phosphohydrolase family that embraces the RNA triphosphatase components of the capping enzymes of fungi and protozoa (34-36). Metazoans and plant proteomes have no recognizable homologues of the metal-dependent RNA triphosphatases. It is proposed that the fungal/protozoal-type of capping apparatus is the ancestral state of this uniquely eukaryal mRNA processing pathway and that the metazoan/plant capping enzyme evolved in a stepwise process entailing: (i) elaboration of a new cysteine phoshatase paralog with specificity for phosphoanhydrides; (ii) fusion of the cysteine

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phosphatase RNA triphosphatase to a guanylyltransferase; and (iii) loss of the ancestral metal-dependent RNA triphosphatase from the metazoan and plant lineages (33). An evolutionary “transition-state” was inferred in which cysteine

phosphatase-type and metal-dependent RNA triphosphatases would coexist in the same proteome. It would appear that certain baculoviruses and poxviruses exemplify the proposed transition state.

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17. Murshudov, G., Vagin, A., and Dodson, E. (1997) Acta Crystallogr. D53, 240-255 18. Kraulis, P. J. (1991) J. Appl. Crystallogr. 24, 946-950 19. Merritt, E. A., and Murphy, M. E. P. (1994) Acta Crystallogr. D 50, 869-873 20. Nicholls, A., Sharp, K. A., and Honig, B. H. (1991) Proteins: Struct. Funct. and Gen. 11,

281-286 21. Evans, S. V. (1993) J. Mol. Graphics 11, 134-138 22. Mossessova, E., and Lima, C. D. (2000) Mol Cell 5, 865-876 23. Holm, L., and Sander, C. (1993) J. Mol. Biol. 233, 123-138 24. Lee, J. O., Yang, H., Georgescu, M. M., Di Cristofano, A., Maehama, T., Shi, Y., Dixon, J.

E., Pandolfi, P., and Pavletich, N. P. (1999) Cell 99, 323-334 25. Song, H., Hanlon, N., Brown, N. R., Noble, M. E., Johnson, L. N., and Barford, D. (2001)

Mol Cell 7, 615-626 26. Gray, C. H., Good, V. M., Tonks, N. K., and Barford, D. (2003) Embo J 22, 3524-3535 27. Yuvaniyama, J., Denu, J. M., Dixon, J. E., and Saper, M. A. (1996) Science 272, 1328-1331 28. Jia, Z., Barford, D., Flint, A. J., and Tonks, N. K. (1995) Science 268, 1754-1758 29. Andersen, J. N., Mortensen, O. H., Peters, G. H., Drake, P. G., Iversen, L. F., Olsen, O. H.,

Jansen, P. G., Andersen, H. S., Tonks, N. K., and Moller, N. P. (2001) Mol Cell Biol 21,7117-7136

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32. Jin, J., Dong, W., and Guarino, L. A. (1998) J Virol 72, 10011-10019 33. Shuman, S. (2002) Nat Rev Mol Cell Biol 3, 619-625 34. Martins, A., and Shuman, S. (2001) J Biol Chem 276, 45522-45529 35. Martins, A., and Shuman, S. (2003) Nucleic Acids Res 31, 1455-1463 36. Gong, C., and Shuman, S. (2003) Virology 309, 125-134 37. Diederichs, K., and Karplus, P. A. (1997) Nat. Struct. Biol. 4, 269-275

FOOTNOTES

*We thank Fan Zhang for help with the crystallization trials. This work was supported from grants from the NIH (GM51350 to A.M and GM42498 to S.S.). S.S. is an American Cancer Society Research Professor. AC present address: Department of Biochemistry, Duke University Medical Center, Durham, North Carolina 27710.

FIGURE LEGENDS

Fig. 1. Overall structure of BVP. (A) The overall α/β topology of the BVP structure is shown as a ribbon diagram with secondary structural elements numbered sequentially from the N-terminus to the C-terminus. The P-loop (residues 118-126) is colored in blue and highlighted by the catalytic Cys119, which is depicted in ball-and-stick. The two phosphate ions bound in the active site region are also shown in ball-and-stick. (B) A stereo view of the superposition of phosphate-bound BVP (red) and unliganded Mce1 (gray) illustrates the overall similarity between the two structures. The P-loop in BVP is colored blue while Cys119 and the phosphate ions in the BVP structure are shown as ball-and-stick. Residues 5-175 of Mce1 (PDB ID 1I9T) and BVP molecule A (residues 1-168) were superimposed using the program LSQKAB from the CCP4 suite (13). Differences between the two structures are mostly localized to variable loop regions, including a flexible loop that moves towards the phosphate-occupied active site in BVP.

Fig. 2. Glycerol gradient sedimentation of His10-BVP and native BVP. Velocity sedimentation of His10-BVP (A) or native BVP (B) was performed as described under Experimental Procedures. Aliquots of even numbered fractions were analyzed by SDS-PAGE. The Coomassie Blue-stained gels are shown in the top panels. Triphosphatase activity profiles are shown in the bottom panels.Reaction mixtures (10 µl) containing 50 mM Tris-HCl (pH 7.5), 5 mM DTT, and 100 µM [γ-32P]ATP, and 2 µl of the indicated gradient fractions enzyme were incubated for 15 min at 30°C, then quenched with 2.5 µl of 5 N formic acid. Aliquots were applied to polyethyleneimine cellulose TLC plates that were developed in 0.5 M LiCl, 1 M formic acid. The [γ-32P]ATP and 32Pi were quantified by scanning the TLC plate with a Fujix imaging apparatus. The positions of catalase, BSA, and cytochrome c standards sedimented in a parallel gradient are indicated by arrows in A, bottom panel.

Fig. 3. Interactions between BVP and bound phosphate ions. (A) A stereo view of the refined 2Fo-Fc electron density at 1.5 Å resolution for the active site environment of molecule A is shown contoured at 1.2 σ. The phosphate ions are colored green. The electron density shown here is representative of the well-ordered active site regions found in the other two monomers. Water molecules have been omitted for clarity. (B) A stereo view of the active site region in BVP illustrates hydrogen-bonding interactions between the bound phosphate ions and residues from the P-loop and neighboring loops. Several well-ordered water molecules in the active site vicinity are depicted as red spheres.

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Fig. 4. (A) Surface representation of the active site pocket with bound phosphate ions. A close-up view of the active site region in BVP is shown as a surface rendering colored by electrostatic potential. Blue and red regions correspond to positively and negatively charged areas, respectively. The phosphate at Site 1 binds deep in the positively charged catalytic pocket while the Site 2 phosphate ion sits in a neighboring hollow. The approximate locations of the catalytic cysteine and other charged residues that are ideally positioned to interact with a triphosphate moiety are indicated. (B, C, D) Comparison of crystal packing interactions at each BVP monomer active site. Molecules A, B, and C are colored red, green, and purple, respectively while neighboring symmetry related molecules and their residues are depicted in gray. The P-loop is highlighted in blue with catalytic residues Cys119 and Arg125 shown in ball-and-stick and the bound phosphates colored magenta.

Fig. 5. Effects of Y67A and T120A mutations on triphosphatase and diphosphatase activity. Reaction mixtures (50 µl) containing 50 mM Tris-HCl (pH 7.5), 5 mM DTT, 2 mM ATP (panel A) or ADP (panel B), and wild-type, Y67A, or T120A His10-BVP proteins as specified were incubated for 15 min at 30°C. The reactions were quenched by adding 1 ml of malachite green reagent (BIOMOL Research Laboratories, Plymouth Meeting, PA). Phosphate release was determined by measuring A620 and interpolating the data to a phosphate standard curve. Background levels of inorganic phosphate (~ 1 to 3% of the input levels of ATP or ADP substrate) determined from the A620 of control reaction mixtures lacking BVP were subtracted from the values measured for the reactions containing BVP.

Fig. 6. Comparison of BVP and Mce1 catalytic regions. A close-up stereo view of the overlaid P-loop regions in BVP (colored by atom) and Mce1 (magenta) is shown to highlight differences in the active site environment and the conformational changes that occur upon phosphate binding. BVP and equivalent Mce1 residues are labeled in black and magenta, respectively.

Fig. 7. Structural alignment of the RNA-specific cysteine phosphatases. A structure-based sequence alignment of RNA triphosphatases belonging to the cysteine phosphatase family reveals residues conserved throughout the family and highlights differences (denoted by |) between the monofunctional and bifunctional branches. The alignment includes amino acid sequences of the N-terminal RNA triphosphatase domains of bifunctional capping enzymes (CE) from mouse (Mus CE), C. elegans (Cel CE), D. melanogaster (Dme CE), X. laevis (Xla CE), Anophelesgambiae (Aga CE), two capping enzymes of A. thaliana (Ath CE and CE’) and a capping enzyme of the ISKNV iridovirus (ISKNV). Also included are the monofunctional RNA tri/diphosphatases human PIR1 (HsPIR1), baculovirus BVP, and homologs from canarypox virus (CPV) and Amsacta moorei entomopox virus (AmEPV). Gaps in the sequences are indicated by dashes. The cysteine phosphatase signature motif (the P-loop) is highlighted in the shaded box. Secondary structure elements of the mouse RNA triphosphatase (PDB ID code: 1I9S) and BVP are shown above and below the respective sequences, with α helices depicted as bars, β strands as arrows, and loops as solid lines. The residues defined by alanine-scanning as essential for BVP triphosphatase activity are indicated by below the sequence.

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Table 1. Data collection and refinement statistics Data collectionWavelength (Å) 0.9479 Cell dimensions (Å) (P21) a=40.9 Å, b=74.1 Å, c=105.2 Å, β=92.4°Resolution (Å) 1.50 Measured reflections 424,098 Unique reflections 97,263 Completeness (%)a 97.1 (90.4) Rsym (%)a,b 6.6 (25.0) Rmeas (%)a,c 7.3 (30.2) Redundancy 4.4 (2.8) I/σ(I) 6.8 (2.7) RefinementResolution (Å) 20.0-1.50 Number of reflections: working set/test set 92,343/4867 R-factord 16.5 Rfree

e 19.4 Number of protein atoms 4073 Number of water molecules 567 Number of ions 7 R.m.s.d. Bond lengths (Å) Bond angles (°)

0.011.2

Average B-factor (Å2): Main chain Side chain Solvent

15.117.430.3

aNumbers in parenthesis represent values in the highest resolution shell (1.50-1.54 Å). bRsym=Σ|I-<I>|/ΣI, where I=observed intensity, and <I>=average intensity obtained from multiple measurements. cRmeas as defined by (37). dR-factor=Σ||Fo|-|Fc||/Σ|Fo|, where |Fo|=observed structure factor amplitude and |Fc|=calculated structure factor amplitude. eRfree: R-factor based on 5% of the data excluded from refinement.

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A)

B)

Figure 1

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Figure 2

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A)

B)

Figure 3

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Figure 4

Site 2

Site 1

B C D

A

Monomer A Monomer B Monomer C

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0

5

10

15

20

25

30P

ire

leas

e(n

mol

)

0 5 10 15

BVP (g)

0

5

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25

30

0 5 10 15

BVP (g)

WT

Y67A

T120A

A. ATPase B. ADPase

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Figure 6

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P-loop

Mus CE KIPPRWLNCPRRGQPVAG-RFLPLKTMLGPRYDSQV-AEEN-RFHPSMLSNYLKSLKVKMSLLVDLTNTSRFYD-RNDIEKEGIKYIKLQCKGHGECPTTENTETFIRLC 110 Cel CE GLPDRWLHCPKTGTLINNL-FFPFKTPLCKMYDNQI-AERRYQFHPAEVFSHPHLHGKKIGLWIDLTNTDRYYF-REEVTEHECIYHKMKMAGRGVSPTQEDTDNFIKLV Dme CE PLPNRWLYCPRKSDTIIAERFLAFKTPLSNNFHDKM-PIEC-TFQPEMLFEYCKTLKVKLGLWVDLTNTKRFYD-RSAVEELGAKYIKLQCRGHGETPSPEQTHSFIEIV Xla CE KIPPRWLNCPRRGQPVAA-KFLPLKTMLGPKYDDQV-PEEN-RFHPSMLSNYLKSLKVKMGLLVDLTNTTRFYD-RNDIEKEGIKYIKLQCKGHGECPSQENTDTFLRLC Aga CE PIPHRWLHCPRKSDSIIADRFIAFKTPLKRDFQSQM-PVQC-SFAPSMLFDLMKRQKRRIGLWIDLTNTNRFYD-KNEIEDAGATYIKLKCRGHGETPSVEHVRSFIEIV Ath CE KIPQGWLDCPPSGNEIGF--LVPSKVPLNESYNNHV-PPGS-RYSFKQVIHNQRIAGRKLGLVIDLTNTTRYYS-TTDLKKEGIKHVKIACKGRDAVPDNVSVNAFVNEV Ath CE’ TIPQGWLDCPRFGQHIGL--IIPSKVPLSESYNDCV-PSGK-RYNFKQWLT-------KIGLVIDLTNTTRYYHPNTELRQNRIEYVKIRCSGRDSVPDNVSVNTFVHEV ISKNV CE LPPHAWVSCPAQGTEICG-FITPGKTFLDSRYDTYI-AAEA-HYRPPLPDMY--------SAVIDLTNTARYYN-GRAL---GACYHKIKCKGHNQCPSPRAVKAFIDTV HsPIR1 HIPERWKDYLPVGQRMPGTRFIAFKVPLQKSFEKKL-APEE-CFSPLDLFNKIREQNEELGLIIDLTYTQRYYKP--EDLPETVPYLKIFTVGH-QVPDDETIFKFKHAV AmEPV MLPYKWNNYFAHGTIIKCINTICFKLP----------CNGT-EWDICKLINTFPN----LKIVIDFRYSETCYNP-SDLNKLGIEYIKIPIKAQSL-PTDDKINKFFNII CPV KLPDKWLNYTPVGDIIKDTRFIAFKVPLNNKYDKAITDPIN-RFHLEDLINYLTDNGKQLGMIIDLSYSLRYYNP--KLLPSTIRHVKIMLKGRGEIPYIEDVLRFNSEV BVP MFPARWHNYLQCGQVIKDSNLICFKTPLRPELFAYVTSEED-VWTAEQIVKQNPS----IGAIIDLTNTSKYYDG-VHFLRAGLLYKKIQVPGQTL-PPESIVQEFIDTV 103

• | • ••

Mus CE ERFNERS--PPELIGVHCTHGFNRTGFLICAFLVEKMDW-SIEAAVATFAQARPPGIYKGDYLKELFRRYG 178 Cel CE QEFHKKY--PDRVVGVHCTHGFNRTGFLIAAYLFQVEEY-GLDAAIGEFAENRQKGIYKQDYIDDLFARYD Dme CE DNFINER--PFDVIAVHCTHGFNRTGFLIVCYLVERLDC-SVSAALAIFASARPPGIYKQDYINELYKRYE Xla CE DHFIDRN--PTELIGVHCTHGFNRTGFLICAFLVEKMDW-SIEAAVATFAQARPPGIYKADYLKELFRRYG Aga CE EEFIQEH--PLDVIGVHCTHGFNRTGFLIVSYMVERLDC-AVDAAVMAFAQARPPGIYKGDYLKELFARYG Ath CE NQFVLNLKHSKKYILVHCTHGHNRTGFMIVHYLMRSGPM-NVTQALKIFSDARPPGIYKPDYIDALYSFYH Ath CE’ TQFE-NHNLSEKYLLVHCTHGHNRTGFMIVHYLMRSRPMMSVTQALKIFSDARPPGIYKPDYIDALYRFYH ISKNV CE VAA-------PGLVYVHCTYGFNRTGYLICCYLVECRKM-SVHDAIRLFAEARPPGMYKADYIKTLCVKYN HsPIR1 NGFLKENKDNDKLIGVHCTHGLNRTGYLICRYLIDVEGV-RPDDAIELFNRCRGHCLERQNYIEDLQNG-- AmEPV DKYIEL----KYLIGIHCTHGINRTGYMVCKYLIYKFKI-PPYVAINIFEKNRGYYIEREIYINNLLYF* CPV NRFLQFNRDNNKLIGVHCTHGLNRTGYMICRYMIEVCGI-DPAAAIEMFSDARKHKIERPTYILDLMKRKH BVP KEFTEKC--PGMLVGVHCTHGINRTGYMVCRYLMHTLGI-APQEAIDRFEKARGHKIERQNYVQDLLI* 168

•••• •• • • |•

Figure 7

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Anita Changela, Alexandra Martins, Stewart Shuman and Alfonso Mondragonphosphate

Crystal structure of baculovirus RNA triphosphatase (BVP) complexed with

published online February 15, 2005J. Biol. Chem. 

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