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J. Biol. Chem., Vol. 279, Issue 6, 4394-4403, February 6, 2004
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From the
Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston, Massachusetts 02115 and the Department of Biochemistry, University of Wisconsin, Madison, Wisconsin 53706
Received for publication, August 5, 2003 , and in revised form, October 21, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-phosphate from RNA as well. Alanine substitutions in a putative nucleotide binding region of µ2 abrogated both functions but did not affect the purification profile of the protein or its known associations with microtubules and mORV µNS protein in vivo. In vitro microtubule binding by purified µ2 was also demonstrated and not affected by the mutations. Purified µ2 was further demonstrated to interact in vitro with the mORV RNA-dependent RNA polymerase, 
3, and the presence of 3 mildly stimulated the triphosphatase activities of µ2. These findings confirm that µ2 is an enzymatic component of the mORV core and may contribute several possible functions to viral mRNA synthesis. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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1 in the T = 1 shell, 2 in the pentameric surface turrets, and 
2 in the monomeric surface nodules (4). Not visualized in that structure are the two stoichiometrically minor proteins, 3 and µ2, which are located beneath the 1 shell near the icosahedral 5-fold axes (5). To obtain an atomic model for 3, which is the viral RNA-dependent RNA polymerase (6–8) and, thus, a functionally key element, a 2.5-Å crystal structure of recombinant 3 has been determined (9). In addition, the 3 structure has recently been fit into a 7.6-Å electron cryomicroscopy reconstruction of the mORV virion, revealing how 3 associates with the 1 shell (10). These studies leave µ2 as the only core protein for which no crystal structure is yet available (Table I). The roles of µ2 in core functions are also poorly defined.
Cores catalyze at least five different enzymatic reactions (Table I). In addition to the polymerase activity attributable to 3, there are the four reactions in RNA 5' capping: RNA 5'- triphosphatase (RTPase), guanylyltransferase, 7N-methyltransferase, and 2'O-methyltransferase (3, 11). The RTPase activity has been assigned to the shell protein 1 (12). The RNA guanylyl- and methyltransferases are contained in the turret protein 2 (4, 13–18), through which the nascent transcripts are cotranscriptionally exported (19, 20). Cores also catalyze removal of the -phosphate group from NTPs (21–24). This activity might represent the RTPase, as observed with some other such enzymes (25–29), but might alternatively reflect a distinct activity not encompassed by the five reactions above. In fact, published findings suggest there may be two different types of nonspecific nucleoside triphosphatases (NTPases) in cores (23, 24). One of these NTPases might represent a core-associated RNA helicase that is involved in melting the double-stranded RNA genome segments during transcription (23, 24, 30), and indeed evidence for NTP-dependent RNA/DNA helicase activity by shell protein 1 has been reported (30).
Two previous genetic studies have implicated µ2 in the enzymatic activities of cores. Yin et al. (31) show that the M1 genome segment, which encodes µ2, determines in vitro differences between cores of mORV strains type 1 Lang (T1L) and type 3 Dearing (T3D) in both the temperature optimum of transcription and the amounts of transcripts produced. Because 3 is the viral polymerase (6–9), these genetic findings suggest an auxiliary role for µ2 in 3 function. Noble and Nibert (24) show that M1/µ2 determines in vitro differences between T1L and T3D cores in both the response of the ATPase activities to temperature and the rate of GTP hydrolysis at certain conditions. The latter genetic associations are consistent with µ2 functioning as an NTPase or playing a regulatory role in the NTPase function of another core protein, most likely 1 (23, 24, 30). Both µ2 and 1 in vitro have also been shown to bind RNA (32–34). Thus, questions remain about the relative roles of µ2 and 1 in the NTPase activities of cores and the specific roles each plays in viral mRNA synthesis.
Recent studies have used immunofluorescence microscopy to identify in vivo associations between µ2 and other proteins. Initially, µ2 was shown to associate with and stabilize microtubules in both infected and transfected cells (35). Subsequently, µ2 was shown to associate also with the major mORV nonstructural protein µNS (36). These findings suggest important roles for µ2 and µNS in building the cytoplasmic "factories" in which viral genome replication and assembly are thought to occur (35–38).
To learn more about the activities of µ2, we devised a purification protocol and undertook functional studies of the purified protein. Results in this report provide biochemical and genetic evidence that µ2 is itself a divalent cation-dependent NTPase and RTPase. Alanine substitutions in a putative nucleotide binding region of µ2 abrogated both functions but did not substantially affect the purification profile of the protein or its known associations with microtubules and µNS protein in vivo. In vitro microtubule binding by purified µ2 was also demonstrated and not affected by the mutations. Purified µ2 was further demonstrated to interact in vitro with the viral polymerase, 3, and the presence of 3 mildly stimulated the triphosphatase activities of µ2. These findings suggest that µ2 is an enzymatic component of the mORV core and may contribute several possible functions to viral mRNA synthesis.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Generation of a Recombinant Baculovirus for Expression of Mutant K415A/K419A µ2—To obtain an M1 gene encoding alanine substitutions for lysines 415 and 419 in µ2, we first removed the BamHI site from the multiple-cloning region of pBS-M1(T1L) by cutting with BamHI, filling the overhangs with Klenow polymerase, and religating. The resulting plasmid pBS'-M1(T1L) was used as template in an inverse polymerase chain reaction with forward primer 5'-GGGGATCCTTTGCATCAACAATTATGAGAGTTC-3' and reverse primer 5'- GGGGATCCCGCAGGCAGCACAGCGCC-3' (bold italics indicate nucleotide changes for Ala-419 and Ala-415, respectively). In each primer one silent nucleotide change was also introduced to generate a BamHI site (underlined) without affecting the µ2 amino acid sequence. The amplification conditions were the same as those described in Kim et al. (39). The resulting plasmid was subjected to nucleotide sequencing to confirm that between the BsmI and SphI sites in M1, and the BsmI-SphI fragment was then swapped for the same region of pBS-M1(T1L). The XbaI-HindIII fragment from the resulting plasmid pBS-M1(T1L)K415A/K419A was swapped for the same region of pFB-M1(T1L). The resulting plasmid pFB-M1(T1L)K415A/K419A was used to generate a recombinant baculovirus as indicated in the preceding section.
Cloning M1 Genes into the pCI-neo Vector—Generation of plasmid pCI-M1(T1L), encoding wt T1L µ2 protein, was previously described (35). The plasmid pCI-M1(T1L)K415A/K419A was generated in the same manner in that the M1 gene from pBS-M1(T1L)K415A/K419A was excised by digestion with SpeI and XhoI and ligated into the pCI-neo vector (Promega) that had been digested with NheI and SalI.
Purification of wt and K415A/K419A µ2—Sf21 cells were prepared in a 1-liter spinner at a concentration of 1 x 106 cells/ml, into which the recombinant baculovirus to express either wt or K415A/K419A µ2 was inoculated at 5–10 plaque-forming units/cell. At 60 h post-infection, the cells were harvested and washed 3 times with cold phosphate-buffered saline (137 mM NaCl, 3 mM KCl, 8 mM Na2HPO4, 1 mM KH2PO4, pH 7.5). Nuclear extracts were then prepared from these cells as follows. The cell pellet was washed twice with 50 ml of ice-cold Nonidet P-40 buffer and then twice with 50 ml of ice-cold CaCl2 buffer. To make the Nonidet P-40 and CaCl2 buffers 100 ml of Chelsky buffer (10 mM Tris, pH 7.0, 10 mM NaCl, 3 mM MgCl2, 30 mM sucrose) was supplemented with Nonidet P-40 to 0.5% or CaCl2 to 10 mM. The nuclear pellet was resuspended in 50 ml of lysis buffer (20 mM Tris, pH 7.9, 100 mM KCl, 0.2 mM EDTA, 20% glycerol, 1x protease inhibitor mixture (Roche Diagnostics)). Nuclei were lysed on ice with a syringe, and the nucleoplasm was separated from the postnuclear pellet by centrifugation at 15,000 rpm for 30 min in an RC-5B centrifuge with an SS34 rotor (DuPont Sorvall). The nucleoplasm was dialyzed overnight at 4 °C against A0 buffer (20 mM Tris acetate, pH 8.5, 50 mM KCl, 5 mM MgCl2, 10% glycerol, 2 mM 
-mercaptoethanol).
All subsequent column work was performed at room temperature with an Akta-FPLC system (Amersham Biosciences). At each step the column fractions containing µ2 were identified by SDS-polyacrylamide gel electrophoresis and immunoblotting with µ2-specific polyclonal antiserum (36) and an alkaline phosphatase-based color detection method (39). The dialyzed nucleoplasm was applied to a 5-ml HiTrap DEAE-Sepharose column (Amersham Biosciences) preequilibrated with A0 buffer. µ2 was eluted from this column at 
100 mM NaCl in a 0–1 M NaCl gradient in A0 buffer as identified. The fractions containing µ2 were pooled, mixed 1:1 with A0 buffer, and applied to a 1-ml HiTrap Blue (Cibacron Blue)-agarose column (Amersham Biosciences) preequilibrated with A50 buffer (A0 buffer supplemented with 50 mM NaCl). µ2 eluted from this column at 0.8–1.2 M NaCl in a 0–2 M NaCl gradient in A0 buffer. The fractions containing µ2 were pooled, dialyzed overnight at 4 °C against A0 buffer, and applied to a 1-ml HiTrap heparin-Sepharose column (Amersham Biosciences) preequilibrated with A50 buffer. µ2 eluted from this column at 200 mM NaCl in a 0–1 M NaCl gradient in A0 buffer. The fractions containing µ2 were pooled, dialyzed against A100 buffer (A0 buffer supplemented with 100 mM NaCl), and used for all subsequent work as purified µ2. Protein concentration was estimated from absorbance at 280 nm. Yields from this procedure were routinely in the range of 300– 400 µg of µ2 per 4 x 108 cells.
Purification of wt 3 Protein—The T3D 3 protein was expressed and purified as previously described (9). Immediately before use in each experiment, 3 (or bovine serum albumin used in parallel samples) was dialyzed against A100 buffer for 2 h at 4 °C using a Slide-A-Lyzer 10K cassette (Pierce).
NTPase Assays—Reactions for analysis in the colorimetric NTPase assay were performed in A100 buffer lacking -mercaptoethanol. The reactions also contained 2 mM ATP and the indicated amount(s) of protein(s) in a total reaction volume of 60 µl. For some experiments ingredients were altered as specifically described in the figure legends. After incubation at room temperature for an appropriate time (standard, 45 min), the reaction was stopped with 60 µl of 10% trichloroacetic acid. 100 µl of the stopped reaction was then mixed with 100 µl of colorimetric reagent prepared by mixing 6 N sulfuric acid, 0.8% ammonium molybdate, and 10% ascorbic acid in a 1:3:1 ratio. This mixture was incubated at 37 °C for 30 min, and absorbance at 655 nm (A655) was measured with a microplate colorimeter (Molecular Devices). Standard curves generated with a dilution series of KH2PO4 were used to convert the A655 values to amounts of inorganic phosphate (Pi) released (23).
For the radiographic NTPase assay reaction conditions were the same as for the colorimetric assay except that the reaction volume was only 20 µl, and 5 µCi of [
-32P]ATP (3000 Ci/mmol) (PerkinElmer Life Sciences) was used instead of 2 mM ATP. After incubating at room temperature for 30 min, the reaction was stopped with 10 mM EDTA. 1 µl of the stopped reaction was spotted onto a polyethyleneimine-cellulose thin-layer chromatography (TLC) plate (EM Science). After drying the TLC plate was developed in 1.2 M LiCl solvent, and the separated reaction products were visualized by either phosphorimaging (Molecular Dynamics) or exposure to x-ray film (Fuji or Kodak) in the presence of an intensifying screen.
RTPase Assay—MAXIscript in vitro transcription kit (Ambion) was used to generate -labeled 45-nucleotide RNA substrates following the manufacturer's directions. Each 20-µl reaction mixture contained 30 units of T7 RNA polymerase (from the kit), 1x transcription buffer (from the kit), 1 mM ATP, 1 mM CTP, 1 mM UTP, 0.03 mM GTP, 50 µCi of either [-32P]GTP (6000 Ci/mmol) or [-32P]GTP (3000 Ci/mmol) (PerkinElmer), 1 µg of pGEM-4Z (Promega) (linearized with SmaI and purified from an agarose gel), and 1 unit/µl RNasin (Promega). After a 2-h incubation at 37 °C the reaction was adjusted to 50 µl with H2O. Template DNA was degraded by treatment for 30 min at 37 °C with 2 units of DNase I (from the kit). Free nucleotides were removed with a nucleotide removal kit (Qiagen) and then a Sephadex G-25 column (Amersham Biosciences). The quality and purity of the purified 45-mer [-32P]- or [-32P]RNA (i.e. appropriate RNA size and essential absence of [-32P]- or [-32P]RNA and small abortive transcripts) were verified on 10% polyacrylamide gels containing 7 M urea (data not shown). Efficient removal of GTP and small abortive transcripts from the [-32P]RNA was also verified by TLC, in which [-32P]GTP was found to migrate slightly faster than the upper spot of the radiolabeled RNA (data not shown). RNA concentration was estimated by absorbance at 260 nm.
For the RTPase assay reaction conditions were the same as for the radiographic NTPase assay except that specified amounts of 32P-labeled RNA were added instead of ATP. After incubating for the appropriate time at room temperature (standard, 1 h), the reaction was stopped with 10 mM EDTA. For Fig. 6, RNAs were resolved on a 10% polyacrylamide gel containing 7 M urea and then visualized by exposure to x-ray film in the presence of an intensifying screen. Otherwise, reaction products were separated by TLC as described for the radiographic NTPase assay and visualized by phosphorimaging. For quantitation from TLC, the intensity of the 32Pi spot obtained in each µ2- and/or 3-containing sample was expressed as a fraction of that obtained by treatment of the RNA with calf intestinal phosphatase (New England Biolabs). The upper spot attributable to substrate RNA in the TLC plates (see Fig. 7, A and C, for results with [-32P]RNA substrate) was unexpected, suggesting that despite the purification steps and tests of purity described in the preceding section, the samples might be contaminated with GTP. However, the [-32P]RNA substrate was identically resolved into two spots by TLC, the upper of which was not abolished by prior treatment of the sample with calf intestinal phosphatase (data not shown), demonstrating that the upper spot is indeed attributable to RNA, not GTP.
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- and -tubulin by incubation at 37 °C for 20 min. After 40 min at room temperature, the mixtures were overlaid on a 100-µl sucrose cushion and spun at 100,000 x g for 40 min at room temperature in a TL-100 tabletop ultracentrifuge with a TLA-100 rotor (Beckman). The supernatant was removed, and the pellet was resuspended in 50 µl of phosphate-buffered saline. 10 µl of the resuspended pellet was then separated by SDS-polyacrylamide gel electrophoresis and analyzed by immunoblotting with µ2-specific polyclonal antiserum (36) and a horseradish peroxidase-based enhanced chemiluminescence detection method (Pierce). Immunofluorescence Microscopy—Transfection was performed using 2 µg of plasmid DNA and 6 µl of LipofectAMINE reagent (Invitrogen) as previously described (35). At 18 h post-transfection cells were fixed with 100% methanol and stained with either Oregon Green-conjugated µ2-specific polyclonal antibodies, Texas Red-conjugated µNS-specific polyclonal antibodies, or both, as previously described (36). Fluorescence was visualized with a TE-300 inverted light microscope (Nikon), and the collected images were processed and optimized with Photoshop software (Adobe Systems).
Immunoprecipitation—Each of the indicated amounts of purified wt or K415A/K419A µ2 was mixed with 300 ng of purified 3 in binding buffer (20 mM Tris, pH 8.5, 50 mM NaCl, 50 mM KCl, 5 mM MgCl2, 10% glycerol, 0.01% Triton X-100, 2 mM -mercaptoethanol, 1x protease inhibitor mixture). The mixtures were incubated overnight with rocking at 4 °C. µ2-specific polyclonal antibodies (purified from antiserum using a protein A column (36)) were added to a final concentration of 33 µg/ml, and the mixtures were incubated for an additional 2 h at 4 °C. Each mixture was supplemented with 10 µl of protein A Dynabeads (Dynal Biotech) preequilibrated with binding buffer and then incubated for 30 min at room temperature. Bead-bound antigen-antibody complexes were isolated with the use of a magnetic stand (Dynal Biotech) and washed twice with binding buffer and twice with washing buffer (binding buffer supplemented with NaCl to 200 mM and Triton X-100 to 0.05%). The washed beads were resuspended in 50 µl of 1x gel sample buffer (125 mM Tris, pH 8.0, 1% SDS, 2% -mercaptoethanol, 10% sucrose, 0.01% bromphenol blue), boiled, separated by SDS-polyacrylamide gel electrophoresis, and analyzed by immunoblotting with either µ2- or 3-specific polyclonal antiserum (36, 40) and an alkaline phosphatase-based color detection method (39).
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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80,000) was strongly enriched. This band comigrated with the µ2 protein from mORV cores during electrophoresis (Fig. 1B, left) and was also recognized by the µ2 antiserum in immunoblots (Fig. 1B, right). Another band sometimes seen migrating above µ2 (Fig. 1, B, left, and C) was a contaminant found in different amounts in different preparations; it was not recognized by the µ2 antiserum. A minor band migrating below µ2 (Mr 50,000– 60,000) but recognized by the µ2 antiserum was seen in samples of both cores and purified protein (Fig. 1B), suggesting it is a common degradation product of µ2. Preliminary evidence from dynamic light scattering and gel filtration chromatography suggested the purified form of µ2 is most likely a dimer.2 We also found that µ2 expressed from this same baculovirus vector was competent to be packaged into core-like particles when coexpressed with other core proteins.3
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NTP Hydrolysis by Purified µ2 Protein—To determine whether purified µ2 has ATPase activity we examined aliquots of the heparin-column fractions with a colorimetric assay that detects the Pi released by ATP hydrolysis (23, 24). Fig. 1, C and D, show results of a representative experiment. The assay revealed that fraction numbers 11–13 in this experiment had a level of ATPase activity substantially above the background level of other fractions (Fig. 1D). These same fractions were shown to contain the highest amounts of µ2 protein by gel analysis (Fig. 1C). Moreover, the level of ATPase activity of each fraction was proportional to the amount of µ2 the fraction contained (fraction 12 > 13 > 11), strongly suggesting that µ2 is required for the activity. Other colorimetric assays with the peak fractions provided evidence that µ2 is able to hydrolyze GTP, CTP, and UTP as less preferred substrates (data not shown), suggesting that µ2 is a nonspecific NTPase.
Mutant µ2 Protein (K415A/K419A) with Alanine Substitutions in a Putative Nucleotide Binding Motif—After the preceding results we remained concerned that the NTPase activity might be attributable to a contaminant and not µ2 in the purified preparation. To prove that NTP hydrolysis was mediated by µ2 we sought to create a µ2 mutant that lacked this activity. We have previously noted a conserved region of the µ2 primary sequence with some similarities to nucleotide binding A and B motifs (24) (Fig. 2A). Comparison with recently reported sequences for the homologous VP5 proteins of Aquareovirus (41) showed these motifs are conserved despite an overall µ2-VP5 homology of only 24%4 (Fig. 2A). Moreover, the putative A motif of Ortho- and Aquareovirus is very similar to that of Alphavirus Nsp2, a known NTPase and RTPase (28, 42) (Fig. 2A). We, therefore, introduced two alanine-for-lysine substitutions at positions 415 and 419 in the putative A motif (Fig. 2A) in an effort to eliminate the apparent NTPase activity of µ2.
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The µ2 mutant K415A/K419A was expressed using a baculovirus vector and purified through the same procedures as wt µ2. Gels showed no mobility difference between the purified K415A/K419A and wt proteins (Fig. 2B; data not shown). Over the series of columns in the purification, the mutant behaved almost identically to wt protein, except that K415A/K419A µ2 was eluted at slightly lower salt concentration from the Cibacron Blue column (data not shown). The similar purification patterns suggest the mutant is folded and structurally similar to wt protein (see Fig. 3 and "Microtubule Binding and µNS Association by wt and K415A/K419A µ2" below for more evidence). The colorimetric assay revealed that the ATPase activity of purified wt µ2 increased in concert with protein concentration, whereas the activity of K415A/K419A µ2 remained at background level across the range of concentrations (Fig. 2C). These results indicate that µ2 is itself an ATPase and suggest that its putative nucleotide binding A motif is involved in this activity. Based on the comparison with Alphavirus Nsp2 (Fig. 2A), we consider it likely that the conserved Lys-419 residue in mORV µ2 is especially important. It is also interesting that the proline residue conserved in Ortho/Aquareovirus µ2/VP5 and Alphavirus Nsp2 is one of two residues mutated in the temperature-sensitive µ2 protein of mORV mutant tsH11.2 (43).
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Specificity of wt µ2 for Release of -Phosphate from NTPs— The µ2- and 1-influenced NTPase activities in mORV cores are specific for triphosphorylated nucleotides and release only the -phosphate (21–24, 30). We, therefore, tested for such specificity with the purified µ2 proteins. [-32P]ATP was used as substrate in a radiographic assay in which the reaction products were analyzed by TLC and fluorography. In this assay wt µ2 generated ADP as the only radiolabeled product (Fig. 4A, lane 2), indicating that µ2 specifically attacks the - phosphodiester linkage of ATP, generating ADP (labeled) and Pi (not labeled) as products. As expected from previous results, K415A/K419A µ2 showed no activity in this assay (Fig. 4A, lane 3). The addition of EDTA to the reaction containing wt µ2 completely abolished the ATPase activity (Fig. 4A, lane 4), indicating that the activity is dependent on divalent cations, as are many other NTPases (25–29) including those in mORV cores (23) (see below for additional evidence). Specific cleavage of the - phosphodiester linkage was further indicated by experiments in which the colorimetric assay was used to examine the capacity of wt µ2 to hydrolyze GTP, GDP, or GMP and which showed that only GTP could serve for Pi release (Fig. 4B).
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5.7 and >9.0 (Fig. 4D). At higher temperatures (45 or 55 °C) little activity was seen, and the activity at 25 °C was higher than that at 35 °C (Fig. 4D). Experiments to estimate the Vmax, apparent Km, and Kcat values of wt µ2 in hydrolysis of each NTP were performed with increasing concentrations of ATP, GTP, CTP, or UTP (Fig. 4E). The results confirmed ATP as the preferred substrate for hydrolysis and suggested this preference is based in modest increases in both substrate binding affinity and catalytic efficiency relative to GTP or CTP (Table II). UTP was a relatively poorer substrate for hydrolysis (Table II). These estimated values for the kinetic properties of µ2 in NTP hydrolysis are similar to published findings for several other viral NTPases (see "Discussion").
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3 Interaction with wt and K415A/K419A µ2—The two stoichiometrically minor core proteins, µ2 and 3, have been hypothesized to interact within cores (46–48). Direct evidence has remained lacking, but evidence from electron cryomicroscopy has suggested that the two proteins are at least juxtaposed in the core interior (5). In addition, the genetic evidence that µ2 can influence core transcriptase behaviors has suggested a µ2-3 interaction (31). A protocol is available for large scale purification of 3 (8, 9) and was used in this study to obtain purified 3 for investigating its possible interaction with purified µ2 in vitro. In the absence of µ2, 3 was not immunoprecipitated by µ2-specific polyclonal antibodies (Fig. 5A, lanes 1 and 5). In the presence of increasing concentrations of either wt or K415A/K419A µ2, however, the amount of 3 precipitated by the µ2 antibodies was progressively increased (Fig. 5A, lanes 2–4 and 6– 8). These results indicate that µ2 can indeed interact with 3 in vitro. They also indicate that the K415A/K419A mutations do not affect this interaction, providing further evidence for the largely intact conformation of this mutant µ2.
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3 Stimulation of ATPase Activity of wt µ2—Having demonstrated µ2-3 interaction, we hypothesized that 3 binding to µ2 may stimulate the enzymatic activities of the latter. In the colorimetric assay for ATPase activity, neither 3 alone nor mixtures of K415A/K419A µ2 with increasing concentrations of 3 showed activity (Fig. 5B). These findings suggested that 3 does not have intrinsic ATPase activity (that is, in the presence of ATP but no other nucleoside triphosphates or template RNA) and also that K415A/K419A µ2 could not be stimulated to demonstrate this activity by its interaction with 3. On the other hand, mixtures of wt µ2 with increasing concentrations of 3 exhibited increasing levels of ATPase activity (Fig. 5, B and C), suggesting that µ2-3 interaction mildly stimulated this activity of wt µ2. Increasing concentrations of bovine serum albumin had no effect on the ATPase activity of wt µ2 (Fig. 5C), providing evidence that the stimulatory effect was specific to 3. An experiment to estimate the effects of 3 on the kinetic properties of µ2 in ATP hydrolysis (data not shown) suggested that increasing amounts of 3 progressively increased the Kcat of the reaction but had little effect on its apparent Km (Table II). This last finding suggests that 3 acted to increase the catalytic efficiency of µ2 and not its substrate binding affinity.
RTPase Activity of wt µ2—The capacity of wt µ2 to release the -phosphate from an NTP led us to hypothesize that µ2 may also release the 5' -phosphate from an RNA molecule. mORV cores mediate such an RTPase activity, which yields a diphosphorylated RNA 5' end (11, 49) for linkage to GMP by the 2-associated guanylyltransferase (4, 11, 14, 15, 17, 18), as part of the mRNA capping process. The RTPase activity has been previously attributed to the 1 shell protein (12). We nevertheless considered it possible that this activity may instead be represented by the NTPase activity of µ2, and we, therefore, tested purified µ2 for its capacity to release the -phosphate from RNA. An in vitro transcription reaction driven by T7 RNA polymerase was performed in the presence of [-32P]GTP to generate RNA transcripts 45 nucleotides in length in which only the 5'-terminal -phosphate was radiolabeled. Alternatively, the reaction was performed in the presence of [-32P]GTP to generate 45-mer transcripts with the radiolabel in internal positions. The quality of these substrate RNAs and their essential lack of contamination with [-32P]- or [-32P]RNA were confirmed by electrophoresis in denaturing polyacrylamide gels (data not shown). After incubation in the presence of purified wt µ2 followed by denaturing gel analysis, loss of radiolabel from the 45-mer RNA was observed with the [-32P]GTP-labeled substrate but not the [-32P]GTP-labeled substrate (Fig. 6). These results indicate that µ2 indeed displays RTPase activity and not a nonspecific nuclease activity that would have degraded both substrates.
The RTPase activity of µ2 was further analyzed by TLC. This assay revealed that wt µ2 in the presence of increasing concentrations of [-32P]GTP-labeled substrate RNA released increasing amounts of 32P-labeled Pi (Fig. 7A, lanes 3–7). The addition of EDTA abolished the -phosphate release from RNA by wt µ2 (Fig. 7A, lane 8), indicating that the RTPase activity is dependent on divalent cations, as is the NTPase activity. K415A/K419A µ2 showed no activity in the RTPase assay (Fig. 7A, lane 9). This last result suggests that the RTPase activity involves the same putative nucleotide binding region of µ2 as the NTPase, since the K415A/K419A mutations abolished both activities. Consistent with the release of -phosphate by wt µ2, the amount of uncleaved RNA substrate was decreased relative to that in the RNA-alone, EDTA, or K415A/K419A µ2 lane (Fig. 7A, compare lane 7 with lanes 1, 8,or 9). The decreased amount of uncleaved [-32P]RNA substrate, after incubation with wt µ2, was confirmed by electrophoresis in a denaturing 10% polyacrylamide gel, as was the failure of wt µ2 to degrade [-32P]RNA (data not shown). An experiment to estimate the kinetic properties of µ2 in the RTPase reaction (Fig. 7B) suggested that its Vmax, apparent Km, and Kcat values were all substantially lower than those for NTP hydrolysis (Table II). This is consistent with published findings for other viral proteins having both NTPase and RTPase activities (see "Discussion").
3 Stimulation of RTPase Activity of wt µ2—Preceding evidence for the capacity of 3 to stimulate the ATPase activity of wt µ2 (Fig. 5, B and C) led us to expect a similar effect on RTPase activity. Indeed, when [-32P]RNA was used as substrate neither 3 alone (Fig. 7C, lane 5) nor mixtures of K415A/K419A µ2 with 3 (data not shown) exhibited RTPase activity, but mixtures of wt µ2 with increasing concentrations of 3 exhibited mildly increasing levels of RTPase activity (Fig. 7C, lanes 3 and 4; Fig. 7D). These findings suggest that 3 does not have intrinsic RTPase activity, that K415A/K419A µ2 cannot be stimulated to demonstrate RTPase activity by its interaction with 3, and that the presence of 3 mildly stimulates the RTPase activity of wt µ2.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-phosphate from either an NTP or the 5' end of an RNA. When a purified protein is shown to exhibit a new enzymatic activity, it is always possible that the activity is attributable to a contaminant in the preparation and not the protein of interest. To rule out that possibility we introduced alanine substitutions in µ2 that we expected to reduce its NTPase and/or RTPase activities because the substitutions were located in a putative nucleotide binding region of µ2. These substitutions did not substantially affect the purification profile of the protein but did abolish its capacity for -phosphate removal from either type of substrate. Thus, both activities are attributable to the wt µ2 protein. Moreover, both activities involve the same putative nucleotide binding region of µ2 in which the alanine substitutions were located. An asparagine substitution for Lys-192 in Semliki Forest virus (Alphavirus) Nsp2 (see Fig. 2A) has been shown to abrogate both its NTPase and its RTPase activities (28). Similarly, alanine substitutions that inactivate the baculovirus and vaccinia virus RTPases also abrogate their NTPase activities (25, 26, 29).
Further evidence that the NTPase and RTPase activities are attributable to µ2 is that 3 mildly stimulated both activities of wt but not mutant, µ2. A viral component such as 3 would be unlikely to have a stimulatory effect on the activities of a cellular contaminant, but since 3 interacts with µ2 (Fig. 5), its stimulatory effect on the NTPase and RTPase activities of µ2is not surprising. Because (i) 3 alone did not show these activities, (ii) no increase in activities was seen when 3 was incubated with mutant µ2, and (iii) 3 interacts with both wt and mutant µ2, the increase in activities when 3 was incubated with wt µ2 cannot be attributed to an effect of µ2 binding on latent activities of 3. The new evidence for µ2-3 interaction in vitro also supports the hypothesis that 3 and µ2 interact within mORV cores, as previously suggested by electron cryomicroscopy and genetic results (5, 31). Such an interaction would further manifest the close juxtaposition of different transcription and capping enzymes through protein-protein contacts within viral particles (4, 5) (see Table I), as has been recently shown for analogous cellular enzymes (for review, see Ref. 50 and 51).
Our estimated values for the kinetic properties of µ2 in NTP hydrolysis (Table II) are similar to published ones for several other divalent cation-dependent viral NTPases. For example, the apparent Km for ATP hydrolysis by NTPase/RNA helicase NPH-II from vaccinia virus has been reported as 1.2 mM (52), and the Kcat for ATP hydrolysis by NTPase/RTPase D1 from vaccinia virus has been reported as 606 min-1 (27). In addition, we found apparent Km and Kcat values for the triphosphatase activities of µ2 to be much lower for RNA than for NTP substrates (Table II), and this trend has also been seen with other viral proteins. For example, the apparent Km for the RTPase activity of vaccinia virus D1 has been reported as 1.0 µM compared with 0.8 mM for ATP hydrolysis (27), and the Kcat for the RTPase activity of Semliki Forest virus Nsp2 has been reported as 5.5 min-1, compared with 230 min-1 for GTP hydrolysis (28).
The literature indicates that it is common for RTPases to exhibit NTPase activities (25–29). The reciprocal statement is harder to support, however, because NTPases have been less routinely tested for RTPase activity. We nonetheless expect that many NTPases could act as RTPases in vitro without the latter being a normal aspect of their functions in cells. Thus, our demonstration that µ2 exhibits RTPase activity in vitro does not necessarily mean that it acts as an RTPase in cores. The same logic applies to the in vitro RTPase activity reported for the mORV 1 protein (12).
The findings in this report corroborate previous genetic evidence for a role of µ2 in mORV core NTPase activities (24). Future investigations can now be focused on the specific role(s) of the triphosphatase activities of µ2 in core functions. For example, it will be important to dissect whether µ2 functions as an NTPase, an RTPase, or both within cores. The 1 shell protein has been proposed to be the capping RTPase in cores based on its reported in vitro RTPase activity (12). However, given our new evidence for in vitro RTPase activity of µ2, there appears to be little reason from enzymatic or genetic data to conclude that 1 is more likely than µ2 to represent the capping RTPase. The apparent Km values for RNA substrate reported for 1 and µ2 are similarly low (0.26 and 0.32 µm, respectively), although the reported Kcat of the 1-RNA reaction (3.1 min-1) is 10-fold higher than that of the µ2-RNA reaction (0.3 min-1) (Ref. 12; this study, Table II). An in vitro system for coupled transcription and capping reconstituted from wt or mutant mORV proteins would be useful for dissecting the relative roles of 1 and µ2 but has not been reported to date. We are currently testing core-like particles assembled in insect cells from baculovirus-expressed mORV proteins (39)3 for this purpose. New information about the precise structural positions and orientations of proteins within the core may also be informative since whether the catalytic regions of 1 or µ2 have access to the triphosphorylated 5' end of nascent mRNA within the crowded core interior may be a key determinant of whether either protein truly acts as an RTPase during viral mRNA synthesis (4, 9, 10).4
What other types of functions might µ2 or 1 mediate that could be associated with NTP hydrolysis? Many enzymes are known to couple NTP hydrolysis to protein conformational changes; RNA and DNA helicases, myosins, kinesins, and G proteins are well known examples. In many cases the conformational changes allow movement of the NTPase along a nucleic acid or protein polymer track (53, 54). The 3 crystal structure suggests some such possible roles for µ2 or 1 in melting, translocating, and/or reannealing the genomic plus-strand RNA as it is passed around the outside of the 3 polymerase, whereas the genomic minus-strand RNA is passed through the central cavity (and catalytic site) of 3 during transcript elongation (9). Alternatively, an NTP-dependent activity by either µ2or 1 could be required during a limited part of each transcription cycle, such as for template melting or translocation before the transcript has reached a certain length or for reinitiation after termination. To understand the functions of the mORV core as a molecular machine for mRNA synthesis, the roles of µ2 and 1 must be characterized in more detail. The capacity of µ2 to bind to microtubules (this study, Fig. 3B; also see Refs. 35 and 36) dictates that we must also be alert to possible NTP-dependent functions of µ2 during mORV genome replication and assembly in infected cells.
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FOOTNOTES |
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¶ Present address: James A. Baker Institute for Animal Health, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853.
|| To whom correspondence should be addressed: Dept. of Microbiology and Molecular Genetics, 200 Longwood Ave., Boston, MA 02115. Tel.: 617-645-3680; Fax: 617-738-7664; E-mail: mnibert{at}hms.harvard.edu.
1 The abbreviations used are: mORV, mammalian Orthoreovirus; RTPase, RNA 5'-triphosphatase; NTPase, nucleoside triphosphatase; T1L, type 1 Lang; T3D, type 3 Dearing; wt, wild type; Sf21 cells, S. frugiperda 21 cells; A655, absorbance at 655 nm; MES, 2-(N-morpholino)-ethanesulfonic acid.
2 X. Lu, J. Kim, M. L. Nibert, and S. C. Harrison, unpublished observation.
3 J. Kim and M. L. Nibert, unpublished observation.
4 J. Kim, Y. Tao, K. M. Reinisch, S. C. Harrison, and M. L. Nibert, submitted for publication.
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ACKNOWLEDGMENTS |
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3 purification, Teresa Broering for use of her µNS antiserum and expression constructs, Michael Farzan and Steve Buratowski for suggestions about enzyme assays, and other members of our laboratory for helpful discussions. We also thank Teresa Broering and Cathy Miller for reviewing preliminary drafts of the manuscript. |
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