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Journal of Virology, November 2002, p. 11321-11328, Vol. 76, No. 22
0022-538X/02/$04.00+0     DOI: 10.1128/JVI.76.22.11321-11328.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Role of the Alfalfa Mosaic Virus Methyltransferase-Like Domain in Negative-Strand RNA Synthesis

A. Corina Vlot, Aymeric Menard,{dagger} and John F. Bol*

Gorlaeus Laboratories, Institute of Molecular Plant Sciences, Leiden University, Leiden, The Netherlands

Received 10 May 2002/ Accepted 15 August 2002


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ABSTRACT
 
RNAs 1 and 2 of the tripartite genome of alfalfa mosaic virus (AMV) encode the replicase proteins P1 and P2, respectively. P1 contains a methyltransferase-like domain in its N-terminal half, which has a putative role in capping the viral RNAs. Six residues in this domain that are highly conserved in the methyltransferase domains of alphavirus-like viruses were mutated individually in AMV P1. None of the mutants was infectious to plants. Mutant RNA 1 was coexpressed with wild-type (wt) RNAs 2 and 3 from transferred DNA vectors in Nicotiana benthamiana by agroinfiltration. Mutation of His-100 or Cys-189 in P1 reduced accumulation of negative- and positive-strand RNA in the infiltrated leaves to virtually undetectable levels. Mutation of Asp-154, Arg-157, Cys-182, or Tyr-266 in P1 reduced negative-strand RNA accumulation to levels ranging from 2 to 38% of those for the wt control, whereas positive-strand RNA accumulation by these mutants was 2% or less. The (transiently) expressed replicases of the six mutants were purified from the agroinfiltrated leaves. Polymerase activities of these preparations in vitro ranged from undetectable to wt levels. The data indicate that, in addition to its putative role in RNA capping, the methyltransferase-like domain of P1 has distinct roles in replication-associated functions required for negative-strand RNA synthesis. The defect in negative-strand RNA synthesis of the His-100 and Cys-189 mutants could be complemented in trans by coexpression of wt P1.


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INTRODUCTION
 
Alfalfa mosaic virus (AMV) is a positive-strand RNA virus belonging to the family Bromoviridae. It has a tripartite genome, with RNAs 1 and 2 encoding the replicase proteins P1 and P2, respectively. Of these, P1 contains regions with homology to known methyltransferase domains in its N-terminal half, as well as a superfamily I helicase-like domain in its C-terminal half (10, 33) (Fig. 1A). P2 is the viral RNA-dependent RNA polymerase protein (RdRp) (27). AMV RNA 3 codes for the movement protein (P3) as well as the coat protein (CP), which is translated from subgenomic mRNA 4. A mixture of the genomic RNAs of AMV is infectious only when either CP or RNA 4 is added to the inoculum in a process called genome activation (7, 13). However, it was recently shown that addition of a poly(A) tail to the normally unpolyadenylated 3'-untranslated regions (UTR) of the AMV RNAs can substitute for the requirement for CP in the inoculum (25). All AMV RNAs are capped, with uncapped RNAs 1 and 2 being noninfectious. In addition, uncapped RNA 4 is unable to activate the genome.



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  		FIG. 1. (A) Schematic representation of AMV P1. Black boxes, N-terminal methyltransferase (MT)-like and C-terminal helicase (HEL)-like domain motifs. (B) Amino acid sequence alignment of the AMV methyltransferase-like domain and the methyltransferase domains of BMV, HEV, SFV, and SIN (adapted from reference 33). The conserved residues that are mutated in this study are boxed. {Delta}, sequence corresponding to 55 amino acids in AMV P1 that is not shown in the alignment.

Viruses that belong to the alphavirus-like superfamily of viruses replicate in the cytoplasm of infected cells. They use the host translation machinery for translation of their RNAs, which are capped with a 7-methyl-G(5')ppp moiety. Since cellular mRNAs are capped in the nucleus, the host capping proteins are not accessible to alphavirus-like viruses. Consequently, these viruses encode their own capping enzymes. The capping reaction of alphavirus RNAs can be divided into four steps (3, 6): (i) dephosphorylation of the viral RNA 5'-pppN end, resulting in a 5'-ppN terminus, (ii) methylation of GTP, resulting in m7GTP, (iii) complex formation of m7GTP and the viral methyltransferase protein (MT), resulting in m7GMP-MT, and (iv) transfer of the m7GMP moiety of m7GMP-MT to the 5'-ppN terminus of the RNA, resulting in m7GpppN(pN)n. It has been shown that steps ii and iii, the methyltransferase and guanylyltransferase reactions, are supported in vitro by the proposed capping protein domains of Semliki Forest virus (SFV), tobacco mosaic virus, bamboo mosaic virus (BaMV), hepatitis E virus (HEV), and brome mosaic virus (BMV) (1, 4, 15, 20, 22, 23). In addition, the proposed capping protein of Sindbis virus (SIN) can support at least the methyltransferase reaction of the capping process in vitro (43). Recently, RNA 5'-triphosphatase activities (step i of the capping process) were assigned to the helicase-like proteins of SFV, SIN, and BaMV (21, 41). Moreover, it was shown that BaMV RNA can be capped in vitro by sequential treatment of the template with the BaMV helicase-like and methytransferase domains (21).

Despite a very low sequence homology between the proposed capping proteins of alphaviruses and alphavirus-like viruses, four highly conserved sequence motifs were distinguished (33). Several conserved motifs in the proposed capping proteins of AMV, BMV, HEV, SFV, and SIN are shown in Fig. 1B. Motif I contains an invariant His residue (His-100 in AMV P1), motif II contains a conserved AspXXArg sequence (Asp-154 and Arg-157 in AMV P1), and motif IV is characterized by an invariant Tyr residue (Tyr-266 in AMV P1). In addition, two conserved Cys residues have been recognized (Cys-182 and Cys-189 in AMV P1) (33). Mutation of these conserved residues affected infectivity, methyltransferase activity, and/or guanylyltransferase activity of BMV, SFV, and SIN (1, 2, 4, 43).

Previously, we showed that the replicase proteins P1 and P2 of AMV could be transiently expressed in Nicotiana benthamiana leaves by agroinfiltration (42). Expression of RNAs 1 and 2 from a transferred-DNA (T-DNA) vector resulted in formation of an RdRp complex that was active both in vivo and in vitro. Accumulation of P1 and P2 in this system was not affected by mutations that inhibited replication of RNA 1 or 2 (42). In the present study, agroinfiltration was employed to express AMV RNAs 2 and 3 in N. benthamiana together with RNA 1 sequences encoding a mutation of one of six conserved residues in the methyltransferase-like domain of P1 (Fig. 1B). The mutations abolished infectivity of the virus and affected the accumulation of negative- and positive-strand RNA to various degrees. Several mutations that affected negative-strand RNA synthesis could be complemented in trans by coexpression of wild-type (wt) P1. Partial purification of the transiently expressed mutant replicases yielded preparations supporting transcription of positive- and negative-strand AMV template RNAs in vitro at levels ranging from 0 to 100% of those for the wt. The data indicate that, in addition to causing a putative defect in capping activity, the mutations affected other replication-associated functions.


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MATERIALS AND METHODS
 
DNA constructs. The construction of plasmids pBS-R1, pMOGR2, pMOGR12, pMOGR12-C189S, pMOGR3, and pMOGR1{Delta}/2{Delta} was described previously (42). pBS-R1 contains AMV cDNA 1 cloned between the cauliflower mosaic virus 35S promoter and the terminator sequence of the nopaline synthase gene (Tnos). pMOG800 is the binary vector used to express cDNA 1 (R1), cDNA 2 (R2), and cDNA 3 (R3) upon agroinfiltration. The empty vector was used as a negative control (42). pMOGR12 contains both cDNA 1 and cDNA 2. In pMOGR12-C189S cDNA 1 encodes the C189S mutation. pMOGR1{Delta}/2{Delta} contains cDNAs 1 and 2 with the 3'-UTRs deleted.

Mutations were introduced by PCR-mediated site-directed mutagenesis. To introduce H100A, nucleotides 214 to 407 of AMV cDNA 1 were amplified by PCR with primers pCo10 (5'CGACACAACTGCAATCGG3') and pCo11 (5'CCGCAAAGCATGCAGATGAACTCGAACTG3'). Nucleotides 389 to 1248 were amplified with primers pCo12 (5'CATCTGCATGCTTTGCGGCTGCCCATC3') and pCL10 (5'GCGGAATGAGCATCAGCG3'). Due to the complementarity of pCo11 and pCo12, the two fragments could be fused by PCR with primers pCo10 and pCL10. The SalI-NdeI fragment of the product of this PCR was exchanged with the corresponding fragment of pBS-R1, yielding pBS-R1H100A. Together with the coding sequence change producing H100A, a silent mutation was introduced into codon 101 of cDNA 1 to create an SphI site.

To introduce D154A, nucleotides 389 to 573 of AMV cDNA 1 were amplified with primers pCo12 and pCo17 (5'GTGAGCCTAGCTCCGGCTCTAG3'). Nucleotides 551 to 904 were amplified with primers pCo16 (5'CTAGAGCCGGAGCTAGGCTCAC3') and pCo13 (5'GACTAGCTCCCAAATTGGGCTCG3'). The two fragments were fused by PCR using primers pCo12 and pCo13. An HpaII site was created by the coding sequence change producing the D154A mutation. To introduce R157G, nucleotides 389 to 573 of AMV cDNA 1 were amplified with primers pCo12 and pCo19 (5'GTGAGCCCTGCTCCGTCTCTAG3'). Nucleotides 551 to 904 were amplified with primers pCo18 (5'CTAGAGACGGAGCAGGGCTCAC3') and pCo13. The two fragments were fused by PCR using primers pCo12 and pCo13. Together with the introduction of R157G, an AluI site was disrupted by a silent mutation in codon 156 of cDNA 1. To introduce coding sequence changes producing D154A and R157G into full-length cDNA 1, the ApaI-SspI fragments of the respective PCR products were first ligated to the SspI-EcoRI fragment of cDNA 1 and cloned in pBluescript SK (+). The NdeI-EcoRI fragments of these constructs were then exchanged with the corresponding fragment of pBS-R1, yielding pBS-R1D154A and pBS-R1R157G.

To introduce C182S, nucleotides 389 to 655 of AMV cDNA 1 were amplified with primers pCo12 and pCo23 (5'GAAACGTATCCATGCTGTAATC3'). Nucleotides 633 to 904 were amplified with primers pCo22 (5'GATTACAGCATGGATACGTTTC3') and pCo13. The two fragments were fused by PCR using primers pCo12 and pCo13. Together with the introduction of C182S, an AflIII site was disrupted by a silent mutation in codon 684 of cDNA 1. To introduce coding sequence changes producing C182S into full-length cDNA 1, the SspI-SstI fragment of the PCR product was first ligated to the SspI-SalI fragment of cDNA 1 and cloned in pBluescript SK(+). The SalI-BglII fragment of this construct was then exchanged with the corresponding fragment of pBS-R1, yielding pBS-R1C182S.

To introduce Y266A, nucleotides 389 to 904 of AMV cDNA 1 were amplified with primers pCo12 and pCo13. Nucleotides 895 to 1248 were amplified with primers pCo14 (5'GGAGCTAGTCATCGGTTTTCATTG3') and pCL10. The two fragments were fused by PCR with primers pCo12 and pCL10. The BglII-HpaI fragment of the product of this PCR was exchanged with the corresponding fragment of pBS-R1, yielding pBS-R1Y266A. An AluI site was disrupted by the coding sequence change producing the Y266A mutation.

All PCR-derived fragments were sequenced (T7 sequencing kit; Amersham Pharmacia Biotech). A silent mutation had been introduced into codon 159 of pBS-R1R157G. All cDNA 1 constructs that were cloned in pBS-R1 were subsequently transferred to pMOGR2 with KpnI and SstI, yielding pMOGR12-H100A, pMOGR12-D154A, pMOGR12-R157G, pMOGR12-C182S, and pMOGR12-Y266A.

Agrobacterium tumefaciens-mediated transient expression. Agroinfiltration experiments were done with N. benthamiana as transient expression of genes from the T-DNA vector is most efficient in this plant species. Moreover, N. benthamiana is the preferred host for the isolation of AMV RdRp. Transformation of the pMOG T-DNA constructs to A. tumefaciens strain LBA4404 by electroporation and agroinfiltration of leaves was done as described previously (42). Bacterial suspensions were infiltrated in N. benthamiana through incisions in the leaves by using a syringe without a needle. To express mutant RNA 1 with wt RNAs 2 and 3, the leaves were coinfiltrated with a mixture of two bacterial suspensions, one expressing RNAs 1 and 2 from a pMOGR12 derivative and the other expressing RNA 3 from pMOGR3. The optical density at 600 nm (OD600) of the mixed suspension was between 1.0 and 1.5, and the OD600 of each strain in the mixture was at least 0.5 (42). Leaves were harvested for total RNA isolation 2 or 5 days after infiltration.

Infectivity studies. RNAs accumulating in agroinfiltrated N. benthamiana leaves were assayed for infectivity by inoculation of RNA extracts from these leaves on the leaves of nontransgenic tobacco plants (Nicotiana tabacum cv. Samsun NN) or transgenic tobacco expressing the AMV P1 and P2 genes (P12 plants). Each tobacco leaf was inoculated with an RNA extract obtained from 50 mg of infiltrated tissue, to which 10 mg of CP had been added (36). The inoculated leaves were harvested for RNA isolation 5 days after inoculation.

Isolation and analysis of RNA. RNA was extracted from 250 mg of plant tissue, either agroinfiltrated N. benthamiana or inoculated N. tabacum, and analyzed by Northern blot hybridization as described previously (42). All Northern blotting was performed with nylon membranes (Roche). RNA from 5 mg of agroinfiltrated N. benthamiana tissue was analyzed with digoxigenin-labeled riboprobes specific for minus strand AMV RNAs 1, 2, and 3. RNA from 0.1 mg of agroinfiltrated N. benthamiana or inoculated tobacco tissue was analyzed with digoxigenin-labeled riboprobes specific for plus strand AMV RNAs 1, 2, and 3. To permit quantification of the RNAs, RNA from 5 mg of agroinfiltrated N. benthamiana tissue was analyzed with 32P-labeled transcripts specific for plus or minus strand AMV RNAs 1, 2, and 3. The radiolabeled transcripts were obtained with T7 RNA polymerase. The RNAs were visualized by autoradiography and quantified with a PhosphorImager (Bio-Rad).

RdRp isolation and analysis. Two days after agroinfiltration, RdRp was isolated from approximately 10 g of N. benthamiana leaf tissue as described previously (30). In short, leaves were homogenized, and large debris and nuclei were spun down. The supernatant was subsequently centrifuged at 30,000 x g for 20 min. To obtain template-dependent RdRp, the 30,000 x g pellet (P30) was solubilized by using a high-salt solution and detergent. The isolate was further purified on a glycerol gradient. The gradients were harvested in 19 fractions of 2 ml each. Fifteen microliters of each fraction was analyzed in an in vitro RdRp assay using plus or minus strand AMV RNA 3 as a template (11). The 32P-labeled RNA products synthesized in vitro by the RdRp were extracted with phenol-chloroform, precipitated, treated with nuclease S1, run on a 1.5% agarose gel, and analyzed by autoradiography.

The template RNAs used in the RdRp assays were transcribed with T7 RNA polymerase (11). Positive-strand AMV RNA 3 was transcribed from pAL3 (26). Negative-strand RNA 3 was transcribed from pT71-301 (38). Therefore, pAL3 and pT71-301 were linearized with PstI, which created overhangs that were made blunt with T4 DNA polymerase.

Protein analysis. Accumulation of P1, P2, and CP in agroinfiltrated N. benthamiana was visualized by Western blot analysis. Five hundred milligrams of tissue was homogenized in 0.5 ml of PEN buffer (10 mM NaH2PO4 [pH 7.0], 1 mM EDTA) 5 days after infiltration. For the detection of CP, an aliquot of the homogenate corresponding to 2.5 mg of infiltrated tissue was loaded onto the blot and analyzed with a rabbit polyclonal CP-specific antibody. For the detection of P1 and P2, the homogenate was centrifuged at 30,000 x g, the pellet (P30 fraction) was resuspended in PEN buffer, and an aliquot corresponding to 50 mg of infiltrated tissue was loaded onto the blot. P1 and P2 were detected by using rabbit polyclonal antibodies directed against the C-terminal 20 amino acids of P1 or the N-terminal half of P2 (37, 39). Western blotting was performed with Hybond-P polyvinylidene difluoride membranes (Amersham Pharmacia Biotech).


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RESULTS
 
Agroinfiltration of AMV P1 mutants. Six mutants were constructed each containing a single substitution of a conserved residue in the putative AMV methyltransferase domain (Fig. 1B). His-100, Asp-154, and Tyr-266 were replaced by Ala residues (mutants H100A, D154A, Y266A), Arg-157 was replaced by a Gly residue (mutant R157G), and Cys-182 and Cys-189 were replaced by Ser residues (mutants C128S and C189S). Full-length mutant cDNA 1 sequences were cloned between the cauliflower mosaic virus 35S promoter and the nos terminator in the T region of the binary vector pMOGR2, which already contained cDNA 2 between the 35S promoter and the nos terminator (42). The resulting pMOGR12 derivatives were transformed to A. tumefaciens. Previously, we showed that infiltration of N. benthamiana leaves with a mixture of two A. tumefaciens suspensions, one containing pMOGR12 and the other containing pMOGR3, resulted in the accumulation of positive-strand RNA, negative-strand RNA, and CP at levels comparable to those of a wt infection (42). In this agroinfection, RNAs 1 and 2 are expressed from pMOGR12 whereas RNA 3 is expressed from pMOGR3. CP is not required to initiate agroinfection, which is probably due to the poly(A) tail of the T-DNA transcripts (25).

The six P1 mutants were expressed in N. benthamiana leaves by infiltration with a mixture of two bacterial suspensions, one containing a mutant pMOGR12 derivative and the other containing pMOGR3. Mixtures of bacteria with the wt construct pMOGR12 and pMOGR3 were used as a positive control, whereas mixtures containing the empty vector pMOG800 and pMOGR3 served as a negative control. Figure 2 shows the accumulation in the agroinfiltrated leaves of viral positive-strand RNA (A), negative-strand RNA (B), and CP (C). Note that lane 8 of Fig. 2A with the wt control was loaded with 33-fold-less material than the other lanes of Fig. 2A. Lanes 1 to 8 of Fig. 2B and C were loaded with equal amounts of RNA or protein extracts. With radiolabeled probes, accumulation of positive- or negative-strand RNA by mutants H100A and C189S was not detectable. For mutant H100A, low levels of positive- and negative-strand accumulation could be visualized when the blots were probed with more-sensitive digoxigenin-labeled probes and exposed to X-ray films for a longer time (data not shown), but signals obtained with this type of label are difficult to quantify. In agreement with their defect in RNA accumulation, mutants H100A and C189S did not induce the accumulation of detectable amounts of CP (Fig. 2C, lanes 2 and 6).



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  		FIG. 2. Accumulation of viral RNA and CP in agroinfiltrated leaves. Leaves were infiltrated with a mixture of a suspension of A. tumefaciens containing the pMOGR3 construct and a suspension of A. tumefaciens containing either the empty T-DNA vector (pMOG800) or construct pMOGR12-H100A, pMOGR12-D154A, pMOGR12-R157G, pMOGR12-C182S, pMOGR12-C189S, pMOGR12-Y266A, or pMOGR12, as indicated above the lanes. (A and B) Viral RNA accumulation. RNA was extracted from the leaves 5 days after infiltration and analyzed by Northern blot hybridization using radiolabeled probes detecting plus strand RNAs 1 to 4 (A) or digoxigenin-labeled probes detecting minus strand RNAs 1, 2, and 3 (B). Lane 8 of panel A was loaded with RNA that was extracted from 33-fold-less leaf material than that from which the RNA loaded onto lanes 1 to 7 was extracted; lanes 1 to 8 of panels B and C were equally loaded. The positions of plus strand RNAs 1 to 4 and minus strand RNAs 1 to 3 are indicated at the left. (C) CP accumulation. Leaves were homogenized 5 days after infiltration, and the homogenates were analyzed by Western blotting using antiserum to CP. The position of CP is indicated at the left. In each panel, the lanes were derived from a single gel.

The other four P1 mutants (D154A, R157G, C182S, and Y266A) induced the accumulation of significant amounts of positive-strand RNA, negative-strand RNA, and CP. As shown in Fig. 2A and as summarized in Table 1, accumulation of positive-strand RNA by these mutants ranged between 0.3 and 2% of that of the wt control. As discussed in the next paragraph, the bands of positive-strand RNAs 1, 2, and 3 in Fig. 2A may represent de novo synthesis of viral RNA and/or nuclear transcripts from the T-DNA vectors that are protected from degradation by the viral CP. Therefore, the levels of RNA 4 were measured to quantify positive-strand accumulation. Accumulation of negative-strand RNA by mutant C182S was detectable only after a longer exposure of the blot of Fig. 2B (result not shown) and was quantified at 2% of that of the wt control (Table 1). In contrast, levels of negative-strand RNA accumulation by mutants D154A, R157G, and Y266A were relatively high and ranged between 27 and 38% of that of the wt control (Fig. 2B and Table 1). Because for some mutants accumulation of negative-strand RNA 1 or 2 was difficult to quantify, the data in Table 1 reflect the accumulation of negative-strand RNA 3. For unknown reasons, some mutations in the pMOGR12 derivatives, particularly H100A, R157G, and Y266A, caused the bacteria to grow to a lower OD600 than the wt control in a few experiments. Therefore, the experiments were repeated four or five times to obtain the data shown in Table 1; Fig. 2 represents the data from a typical experiment.


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  		TABLE 1. RNA accumulation in N. benthamiana leaves agroinfiltrated with AMV mutants

Accumulation of CP in leaves agroinfiltrated with mutants D154A, R157G, C182S, and Y266A (Fig. 2C) paralleled the accumulation of RNA 4 by these mutants (Fig. 2A). Although CP was not quantified, the Western blot data indicated that CP accumulation by these mutants was strongly reduced compared to that of the wt control. By analogy to the data with other alphavirus-like viruses, the mutant RNAs are probably not capped. We have shown that uncapped AMV RNA 4 is translated in transfected protoplasts, albeit with considerably lower efficiency than capped RNA 4 (25). Furthermore, we have shown that positive-strand AMV RNAs, either transcribed from a T-DNA vector or synthesized de novo in agroinfiltrated leaves, are rapidly degraded in the absence of CP (42). Therefore, it is possible that one ore more mutants support the synthesis of a relatively large amount of positive-strand RNA, which is subsequently degraded because of the limited availability of CP. Thus, the synthesis of CP that is visualized in Fig. 2C could have been driven by relatively large amounts of uncapped RNA 4. The possibility that inefficient translation of uncapped RNA 4 serves as a bottleneck in the accumulation of positive-strand AMV RNAs makes it difficult to assess the effect of the P1 mutations on positive-strand RNA synthesis. Our previous data demonstrated that the absence of CP does not affect accumulation of negative-strand AMV RNAs in agroinfiltrated leaves (42). Thus, we limited our conclusions concerning the effects of the P1 mutations on replication in vivo to the synthesis of negative-strand RNA.

Primer extension analyses have been successfully used to discriminate between capped and noncapped RNA of BMV and tobacco mosaic virus (2, 18). However, in our experiments DNA fragments of similar sizes were obtained irrespective of whether primer extension toward the 5' end of the RNA was done with native RNA 4, in vitro-transcribed capped or noncapped RNA 4, or RNA 4 synthesized by the mutant replicase complexes in the infiltrated leaves (data not shown). Thus, we did not obtain definite proof that the positive-strand RNAs produced by our mutants were uncapped.

Inoculation of N. tabacum with RNAs from the infiltrated N. benthamiana leaves. Fragments of RNA 1 spanning the D154A, R157G, C182S, and Y266A mutations were amplified by reverse transcription-PCR on RNA isolated from the infiltrated N. benthamiana leaves. The resulting PCR products were cloned, and, when at least two clones per PCR were sequenced, no revertants were found. Subsequently, nontransgenic N. tabacum as well as P12 tobacco plants were inoculated with RNA from the infiltrated N. benthamiana leaves. P12 plants express P1 and P2 from nuclear transgenes, and inoculation of these plants with RNA 3 results in replication of RNA 3 and synthesis of RNA 4 (36). AMV RNAs from the N. benthamiana leaves with RNA 1 encoding the D154A, R157G, C182S, or Y266A mutation were not infectious to nontransgenic tobacco plants (Fig. 3A, lanes 2 to 5). However, the RNA 3 species that accumulated in the infiltrated N. benthamiana leaves were fully infectious to P12 plants (Fig. 3B, lanes 2 to 5). Note that RNA 3 does not need to be capped to be infectious and that genome activation is not required for infection of P12 plants with RNA 3 (26, 36). So, the amount of RNA isolated from the infiltrated leaves was sufficient to initiate an infection (Fig. 3B), but the mutations introduced into the methyltransferase-like domain of P1 abolished infectivity of the RNA (Fig. 3A). In conclusion, RNAs encoding the D154A, R157G, C182S, or Y266A mutations in the methyltransferase-like domain of P1 could most likely support a single round of replication in agroinfiltrated N. benthamiana, after which the resulting RNAs were not able to sustain an infection.



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  		FIG. 3. Accumulation of viral RNA in leaves of nontransgenic N. tabacum (A) and in leaves of P12 tobacco plants, which express P1 and P2 from nuclear transgenes (B). The leaves were inoculated with RNA isolated from N. benthamiana leaves that had been infiltrated with a mixture of A. tumefaciens containing the pMOGR3 construct and A. tumefaciens containing either the empty T-DNA vector (pMOG800) or construct pMOGR12-D154A, pMOGR12-R157G, pMOGR12-C182S, pMOGR12-Y266A, or pMOGR12, as indicated above the lanes. RNA isolated from leaves that had been infiltrated with A. tumefaciens containing pMOGR3 and A. tumefaciens containing pMOGR12 was diluted 100-fold before inoculation. RNA was isolated from the leaves 5 days after inoculation and analyzed by Northern blot hybridization using digoxigenin-labeled probes detecting plus strand RNAs 1 to 4. The positions of RNAs 1 to 4 are indicated at the left.

Effects of methyltransferase domain mutations on RdRp activity in vitro. Leaves were coinfiltrated with A. tumefaciens containing pMOGR12 derivatives encoding one of the six P1 mutations and A. tumefaciens containing pMOGR3. Two days after infiltration, the leaves were homogenized, after which RdRp was solubilized from the 30,000 x g membrane fraction (P30) and purified on a glycerol gradient. Glycerol gradients were harvested in 19 2-ml fractions starting at the bottom of the gradient, after which RdRp activity was detected by conversion of positive- or negative-strand RNA 3 templates into double-stranded radiolabeled products (Fig. 4A and B). Several mutants induced detectable amounts of RdRp activity, and the sedimentation behavior of this RdRp resembled that of wt RdRp, with most activity being present in fraction 11 of the glycerol gradients. Therefore, fraction 11 was used in the RdRp assays with all mutants. Note that positive-strand RNA 4 was synthesized in the RdRp assays when negative-strand RNA 3 was used as a template (Fig. 4B). Samples from the P30 fractions that were collected during the RdRp isolations were analyzed with antiserum to P1 and P2 to show the relative levels of accumulation of these proteins in the infiltrated leaves (Fig. 4C). The antisera detected minor amounts of host proteins (Fig. 4C, lane 1) that comigrated with P1 and P2, but the viral proteins were consistently detected at above-background levels (Fig. 4C, lanes 2 to 8). P1 and P2 were not detectable in the supernatant (S30 fraction) obtained after centrifugation at 30,000 x g of the homogenates of leaves infiltrated with bacteria containing the wt or mutant constructs (result not shown).



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  		FIG. 4. In vitro activity of RdRp preparations isolated from agroinfiltrated N. benthamiana leaves. Leaves were infiltrated with a mixture of a suspension of A. tumefaciens containing the pMOGR3 construct and a suspension of A. tumefaciens containing either the empty T-DNA vector (pMOG800) or construct pMOGR12-H100A, pMOGR12-D154A, pMOGR12-R157G, pMOGR12-C182S, pMOGR12-C189S, pMOGR12-Y266A, or pMOGR12, as indicated above the lanes. Two days after infiltration, the leaves were homogenized, after which RdRp was solubilized from the 30,000 x g membrane fraction (P30) and sedimented in a glycerol gradient. The gradients were harvested in 19 fractions of 2 ml each with fraction 1 at the bottom of the gradient. (A and B) In vitro RdRp assays. Samples from fraction 11 of the glycerol gradients were used in RdRp assays with plus strand AMV RNA 3 (A) or minus strand RNA 3 (B) as the templates. Radiolabeled products synthesized by the RdRp on these templates were analyzed by gel electrophoresis and autoradiography. The wt RdRp copies plus strand RNA 3 into full-length minus strand RNA 3 and copies minus strand RNA 3 into plus strand RNA 4. The positions of double-stranded RNA (dsRNA) 3 and dsRNA 4 are indicated at the left. (C) Accumulation of P1 and P2. Samples from the P30 fractions of the RdRp isolation procedure were analyzed by Western blotting using antisera to P1 and P2. The positions of P1 and P2 are indicated at the left.

Similar to the defects in RNA synthesis in vivo of mutants H100A and C189S (Fig. 2) was the inactivity of RdRp preparations of these mutants in RNA synthesis in vitro (Fig. 4A and B, lanes 2 and 6). Mutants D154A, R157G, and Y266A induced similar levels of negative-strand RNA accumulation in vivo (27 to 38% of that of the wt control; Table 1 and Fig. 2), but the in vitro RNA-synthesizing activities of the purified RdRp preparations of these mutants were rather different. Synthesis of negative- and positive-strand RNA in vitro (Fig. 4A and B) by the RdRp of mutant D154A (lane 3) resembled that of the wt control (lane 8), while the activity of the RdRp of mutant R157G was reduced 40- to 50-fold (lane 4) and the activity of the RdRp of mutant Y266A was barely detectable (lane 7). RNA synthesis in vitro by the RdRp preparation of mutant C182S (Fig. 4A and B, lane 5) resembled that of mutant R157G (Fig. 4A and B, lane 4) although levels of accumulation of negative-strand RNA by these mutants in vivo are rather different, being 2 and 28% of that of the wt, respectively (Table 1 and Fig. 2). Apparently, there is no clear correlation between the levels of RNA synthesis induced by these mutants in the infiltrated leaves and the activity in vitro of the RdRps isolated from these leaves. Possible explanations for this observation are discussed below.

Negative-strand synthesis of RNA 1 encoding H100A and C189S mutations can be supported in trans. Previously, we obtained evidence that AMV RNAs 1 and 2 require their encoded proteins in cis for replication (24, 40). In the present study, leaves were coinfiltrated with an A. tumefaciens strain carrying pMOGR12, which encodes the H100A or C189S mutation via cDNA 1 and an A. tumefaciens strain carrying pMOGR1{Delta}/2{Delta}, which encodes RNAs 1 and 2 with the 3' UTRs deleted (42). Coinfiltration of N. benthamiana with A. tumefaciens containing the pMOGR1{Delta}/2{Delta} and pMOGR3 constructs resulted in replication of RNA 3 at nearly wt levels, but no negative strands corresponding to the 3'-truncated RNAs 1 and 2 were synthesized (42). Thus, transient expression of P1 and P2 from the pMOGR1{Delta}/2{Delta} construct is sufficient to support replication of RNA 3 in trans. Figure 5 shows that this transient expression also supports in trans the synthesis of negative strands corresponding to RNA 1 encoding the H100A and C189S mutations (lanes 2 and 3). The apparent discrepancy with previous data pointing to a requirement of P1 in cis for replication are discussed below.



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  		FIG. 5. Complementation of P1 mutants by transiently expressed wt P1. Leaves were infiltrated with a mixture of a suspension of A. tumefaciens containing the pMOGR1{Delta}/2{Delta} construct and a suspension of A. tumefaciens containing either the empty T-DNA vector (pMOG800) or construct pMOGR12-H100A, pMOGR12-C189S, or pMOGR12, as indicated above the lanes. RNA was isolated from the leaves 2 days after infiltration and analyzed by Northern blot hybridization using digoxigenin-labeled probes detecting minus strand RNAs 1 and 2. The positions of minus strand RNAs 1 and 2 are indicated at the left.


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DISCUSSION
 
Agroinfiltration of AMV. Coinfiltration of N. benthamiana with A. tumefaciens suspensions containing pMOGR12 and pMOGR3 constructs results in levels of virus accumulation in the infiltrated leaf that are similar to those obtained in a wt infection (42). In the previous study, we provided evidence that the T-DNA is expressed in a large portion of the cells of the infiltrated leaf and that cell-to-cell transport plays no role in the transient expression of the P1 and P2 proteins (42). Here, we used this expression system to study mutations in the methyltransferase-like domain of P1. Six residues that are conserved in capping proteins of a number of alphavirus-like viruses and that are crucial to the capping activity of these proteins (1, 2, 4, 43) were mutated. Mutant viral T-DNA transcripts are capped by the cellular capping machinery, and Western blot data (Fig. 4C) indicated that the six mutations did not grossly affect translation of P1 from these nuclear transcripts or the stability of P1. The finding that none of the mutants was infectious showed that the mutated residues are essential for replication-associated functions of P1. Transient expression of the mutants revealed that some mutations blocked negative-strand RNA synthesis while other mutations mainly affected replication events following negative-strand RNA synthesis.

Replication of AMV with mutations in the methyltransferase-like domain of P1. Mutants H100A and C189S showed virtually no accumulation of negative- or positive-strand RNAs. The results with these mutants point to a role for the methyltransferase-like domain of AMV in the synthesis of viral negative-strand RNA. BMV encoding the equivalent of the H100A mutation replicated in Saccharomyces cerevisiae at 10% of the wt level, while mutation of one of three other residues in the methyltransferase domain of BMV reduced negative-strand synthesis to levels ranging from 1 to 2% of that of the wt to undetectable (2). Furthermore, the SIN capping protein nsP1 has been reported to play a role in negative-strand RNA synthesis (44). It is possible that mutations in the methyltransferase-like domain of AMV affect the functions of other domains in P1 or P2 by changing the tertiary structure of P1. Similar to what was found for the BMV 1a and 2a proteins, it has been shown that the C terminus of AMV P1 interacts with the N terminus of P2 (14, 28, 31, 32, 37). In addition, for the 1a proteins of BMV and cucumber mosaic virus, interactions between the capping and helicase-like domains have been demonstrated, while the capping domains were found to mediate 1a-1a dimerization (28, 29). Moreover, sequences in the capping domains of BMV and SFV (and flanking sequences) mediate membrane association, and it has been proposed that these domains guide RdRp/template complexes to the intracellular membranes, where RNA replication occurs (5, 9, 16, 17, 34).

In infected plant cells, the AMV P1 and P2 proteins accumulate at the vacuolar membrane, and it was suggested that this membrane represents the site of RNA replication (37). After fractionation of a leaf homogenate, the P1 and P2 proteins induced by the wt and mutants H100A and C189S were found in the P30 fraction (Fig. 4C). After solubilization from the P30 fraction, mutant P1 sedimented in the same fractions of the glycerol gradient as wt P1 (data not shown). This could indicate that the mutant replicase proteins are membrane associated but does not prove that they are correctly targeted to putative replication complexes at the vacuolar membrane. We have shown that formation in agroinfiltrated leaves of an RdRp that is active in vitro requires both transient expression of P1 and P2 and the presence of an actively replicating template RNA (42). Possibly, the absence of replication in the leaves infiltrated with mutants H100A and C189S prevented formation of an RdRp with in vitro activity.

Mutants D154A, R157G, and Y266A supported negative-strand RNA synthesis at 30 to 40% of the wt level, whereas accumulation of positive-strand RNA by these mutants was relatively low (2% or less). As discussed in Results, the low level of accumulation of positive-strand RNA could be (partially) due to inefficient translation of uncapped RNA 4 into CP, preventing protection of the RNAs by encapsidation. Alternatively, a defect in capping could promote degradation of the RNAs or the mutant enzymes could be defective in de novo synthesis of positive-strand RNA. Mutation of R136 in the BMV 1a protein (equivalent to R157 in AMV P1) reduced positive-strand RNA accumulation in yeast to less than 0.5% of the wt level (2). However, this accumulation increased dramatically when the mutant was expressed in yeast defective in degradation of uncapped RNAs (2). In addition, 1a stabilizes BMV RNAs in yeast, and it has been proposed that the binding of 1a to the RNAs reflects a role for the protein in recruitment of the RNAs to the membrane-bound replication complexes (8, 12, 34, 35). Interestingly, the R136A mutation in BMV 1a appeared to affect this recruitment function (2).

The observation that mutants D154A, R157G, and Y266A induced the synthesis of negative-strand RNA at 30 to 40% of the wt level suggested that at least some of the replicase proteins and template RNAs of these mutants were properly targeted to membrane-associated replication complexes. Possibly, these mutations mainly affect steps in the replication cycle which occur after negative-strand RNA synthesis, including the capping of the positive-strand RNAs. Because the capping of RNAs does not occur in the in vitro RdRp assay, we surmised that a defect in capping activity would not affect RNA synthesis by the mutant enzymes in vitro. However, only the RdRp containing the D154A mutation supported negative- and positive-strand RNA synthesis in vitro at wt levels (Fig. 4). The strong reduction in the in vitro activity of RdRp from mutants R157G and Y266A could suggest that the mutations affect functions that are involved in RNA synthesis in vitro. Another possibility, however, is that these mutations affect the stability of the RdRp and result in inactivation of the enzyme during the isolation procedure. To our knowledge, the results with mutant D154A represent the first assembly in a plant of an RdRp with full in vitro activity from a noninfectious virus mutant. Possibly, this mutant was defective in RNA capping only.

Mutant C182S supported synthesis of negative-strand RNA at 2% of the wt level. However, this was sufficient to permit both accumulation of positive-strand RNA and CP and formation of an RdRp that showed a low activity in the in vitro RdRp assay. Apparently, mutations in the methyltransferase-like domain of P1 can reduce negative-strand RNA synthesis to levels ranging from undetectable to 40% of the wt level.

P1 supports negative-strand synthesis of RNA 1 in trans. Previously, we showed that transgenic plants that express the AMV P1 protein (P1 plants) support replication of AMV RNAs 2 and 3 (36). In these plants wt RNA 1 was able to coreplicate with RNAs 2 and 3, but this coreplication was abolished by mutations in the P1 gene in RNA 1 (40). Similar results were obtained with protoplasts from P1 plants (24). It was proposed that either the P1 protein is required in cis for replication of RNA 1 or mutant RNA 1 is unable to compete with wt RNAs for the replication machinery. In these previous experiments, only accumulation of positive-strand RNA was analyzed. Here, we showed that the defect in negative-strand RNA synthesis of mutants H100A and C189S could be complemented by coexpression of wt P1 from the pMOGR1{Delta}/2{Delta} construct. Thus, negative-strand RNA 1 synthesis by at least mutants H100A and C189S could be complemented by wt P1 in trans. Similarly, a defect in negative-strand RNA 3 synthesis of a BMV methyltransferase domain mutant could be complemented in yeast by coexpression of wt 1a (2). Moreover, nsP1 could rescue in trans replication of a SIN strain that was conditionally defective in methyltransferase activity (19). A more extensive analysis of cis- and trans-acting functions of AMV P1 and P2 in negative-strand RNA synthesis will be published elsewhere (A. C. Vlot, unpublished data).


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FOOTNOTES
 
* Corresponding author. Mailing address: Institute of Molecular Plant Sciences, Gorlaeus Laboratories, Leiden University, Einsteinweg 55, 2333 CC Leiden, The Netherlands. Phone (31) 71-5274749. Fax: (31) 71-5274469. E-mail: j.bol{at}chem.leidenuniv.nl. Back

{dagger} Present address: Ecologie Microbienne, UMR 5557, Batiment Gregor Mendel, Université Claude-Bernard Lyon 1, 69622 Villeurbanne, France. Back


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REFERENCES
 
    1
  1. Ahola, T., and P. Ahlquist. 1999. Putative RNA capping activities encoded by brome mosaic virus: methylation and covalent binding of guanylate by replicase protein 1a. J. Virol. 73:10061-10069.[Abstract/Free Full Text]
    2. 2
  2. Ahola, T., J. A. den Boon, and P. Ahlquist. 2000. Helicase and capping enzyme active site mutations in brome mosaic virus protein 1a cause defects in template recruitment, negative-strand RNA synthesis, and viral RNA capping. J. Virol. 74:8803-8811.[Abstract/Free Full Text]
    3. 3
  3. Ahola, T., and L. Kääriäinen. 1995. Reaction in alphavirus mRNA capping: formation of a covalent complex of nonstructural protein nsP1 with 7-methyl-GMP. Proc. Natl. Acad. Sci. USA 92:507-511.[Abstract/Free Full Text]
    4. 4
  4. Ahola, T., P. Laakkonen, H. Vihinen, and L. Kääriäinen. 1997. Critical residues of Semliki Forest virus RNA capping enzyme involved in methyltransferase and guanylyltransferase-like activities. J. Virol. 71:392-397.[Abstract]
    5. 5
  5. Ahola, T., A. Lampio, P. Auvinen, and L. Kääriäinen. 1999. Semliki Forest virus mRNA capping enzyme requires association with anionic membrane phospholipids for activity. EMBO J. 18:3164-3172.[CrossRef][Medline]
    6. 6
  6. Bisaillon, M., and G. Lemay. 1997. Viral and cellular enzymes involved in synthesis of mRNA cap structure. Virology 236:1-7.[CrossRef][Medline]
    7. 7
  7. Bol, J. F. 1999. Alfalfa mosaic virus and ilarviruses: involvement of coat protein in multiple steps of the replication cycle. J. Gen. Virol. 80:1089-1102.[Medline]
    8. 8
  8. Chen, J., A. Noueiry, and P. Ahlquist. 2001. Brome mosaic virus protein 1a recruits viral RNA 2 to RNA replication through a 5' proximal RNA 2 signal. J. Virol. 75:3207-3219.[Abstract/Free Full Text]
    9. 9
  9. Den Boon, J. A., J. Chen, and P. Ahlquist. 2001. Identification of sequences in brome mosaic virus replicase protein 1a that mediate association with endoplasmic reticulum membranes. J. Virol. 75:12370-12381.[Abstract/Free Full Text]
    10. 10
  10. Gorbalenya, A. E., E. V. Koonin, A. P. Donchenko, and V. M. Blinov. 1989. Two related superfamilies of putative helicases involved in replication, recombination, repair, and expression of DNA and RNA genomes. Nucleic Acids Res. 17:4713-4729.[Abstract/Free Full Text]
    11. 11
  11. Haasnoot, P. C. J., F. T. Brederode, R. C. L. Olsthoorn, and J. F. Bol. 2000. A conserved hairpin structure in alfamovirus and bromovirus subgenomic promoters is required for efficient RNA synthesis in vitro. RNA 6:708-716.[Abstract]
    12. 12
  12. Janda, M., and P. Ahlquist. 1998. Brome mosaic virus RNA replication protein 1a dramatically increases in vivo stability but not translation of viral genomic RNA 3. Proc. Natl. Acad. Sci. USA 95:2227-2232.[Abstract/Free Full Text]
    13. 13
  13. Jaspars, E. M. J. 1999. Genome activation in alfamo- and ilarviruses. Arch. Virol. 144:843-863.[CrossRef][Medline]
    14. 14
  14. Kao, C. C., and P. Ahlquist. 1992. Identification of the domains required for direct interaction of the helicase-like and polymerase-like RNA replication proteins of brome mosaic virus. J. Virol. 66:7293-7302.[Abstract/Free Full Text]
    15. 15
  15. Kong, F., K. Sivakumaran, and C. Kao. 1999. The N-terminal half of the brome mosaic virus 1a protein has RNA capping-associated activities: specificity for GTP and S-adenosylmethionine. Virology 259:200-210.[CrossRef][Medline]
    16. 16
  16. Kujala, P., A. Ikäheimonen, N. Ehsani, H. Vihinen, P. Auvinen, and L. Kääriäinen. 2001. Biogenesis of the Semliki Forest virus RNA replication complex. J. Virol. 75:3873-3884.[Abstract/Free Full Text]
    17. 17
  17. Lampio, A., I. Kilpeläinen, S. Pesonen, K. Karhi, P. Auvinen, P. Somerharju, and L. Kääriäinen. 2000. Membrane binding mechanism of an RNA virus-capping enzyme. J. Biol. Chem. 275:37853-37859.[Abstract/Free Full Text]
    18. 18
  18. Lewandowski, D. J., and W. O. Dawson. 2000. Functions of the 126- and 183-kDa proteins of tobacco mosaic virus. Virology 271:90-98.[CrossRef][Medline]
    19. 19
  19. Li, M.-L., H.-L. Wang, and V. Stollar. 1997. Complementation of and interference with Sindbis virus replication by full-length and deleted forms of the nonstructural protein, nsP1, expressed in stable transfectants of Hela cells. Virology 227:361-369.[CrossRef][Medline]
    20. 20
  20. Li, Y.-I., Y.-J. Chen, Y.-H. Hsu, and M. Meng. 2001. Characterization of the AdoMet-dependent guanylyltransferase activity that is associated with the N terminus of bamboo mosaic virus replicase. J. Virol. 75:782-788.[Abstract/Free Full Text]
    21. 21
  21. Li, Y.-I., T.-W. Shih, Y.-H. Hsu, Y.-T. Han, Y.-L. Huang, and M. Meng. 2001. The helicase-like domain of plant potexvirus replicase participates in formation of RNA 5' cap structure by exhibiting RNA 5'-triphosphatase activity. J. Virol. 75:12114-12120.[Abstract/Free Full Text]
    22. 22
  22. Magden, J., N. Takeda, T. Li, P. Auvinen, T. Ahola, T. Miyamura, A. Merits, and L. Kääriäinen. 2001. Virus-specific mRNA capping enzyme encoded by hepatitis E virus. J. Virol. 75:6249-6255.[Abstract/Free Full Text]
    23. 23
  23. Merits, A., R. Kettunen, K. Mäkinen, A. Lampio, P. Auvinen, L. Kääriäinen, and T. Ahola. 1999. Virus-specific capping of tobacco mosaic virus RNA: methylation of GTP prior to formation of covalent complex p126-m7GMP. FEBS Lett. 455:45-48.[CrossRef][Medline]
    24. 24
  24. Neeleman, L., and J. F. Bol. 1999. cis-Acting functions of alfalfa mosaic virus proteins involved in replication and encapsidation of viral RNA. Virology 254:324-333.[CrossRef][Medline]
    25. 25
  25. Neeleman, L., R. C. L. Olsthoorn, H. J. M. Linthorst, and J. F. Bol. 2001. Translation of a nonpolyadenylated viral RNA is enhanced by binding of viral coat protein or polyadenylation of the RNA. Proc. Natl. Acad. Sci. USA 98:14286-14291.[Abstract/Free Full Text]
    26. 26
  26. Neeleman, L., A. C. van der Kuyl, and J. F. Bol. 1991. Role of alfalfa mosaic virus coat protein gene in symptom formation. Virology 181:687-693.[CrossRef][Medline]
    27. 27
  27. O'Reilly, E. K., and C. C. Kao. 1998. Analysis of RNA-dependent RNA polymerase structure and function as guided by known polymerase structures and computer predictions of secondary structure. Virology 252:287-303.[CrossRef][Medline]
    28. 28
  28. O'Reilly, E. K., J. D. Paul, and C. C. Kao. 1997. Analysis of the interaction of viral RNA replication proteins by using the yeast two-hybrid assay. J. Virol. 71:7526-7532.[Abstract]
    29. 29
  29. O'Reilly, E. K., Z. Wang, R. French, and C. C. Kao. 1998. Interactions between the structural domains of the RNA replication proteins of plant-infecting RNA viruses. J. Virol. 72:7160-7169.[Abstract/Free Full Text]
    30. 30
  30. Quadt, R., and E. M. J. Jaspars. 1990. Purification and characterization of brome mosaic virus RNA dependent RNA polymerase. Virology 178:189-194.[CrossRef][Medline]
    31. 31
  31. Restrepo-Hartwig, M. A., and P. Ahlquist. 1996. Brome mosaic virus helicase- and polymerase-like proteins colocalize on the endoplasmic reticulum at sites of viral RNA synthesis. J. Virol. 70:8908-8916.[Abstract]
    32. 32
  32. Restrepo-Hartwig, M. A., and P. Ahlquist. 1999. Brome mosaic virus RNA replication proteins 1a and 2a colocalize and 1a independently localizes on the yeast endoplasmic reticulum. J. Virol. 73:10303-10309.[Abstract/Free Full Text]
    33. 33
  33. Rozanov, M. N., E. V. Koonin, and A. E. Gorbalenya. 1992. Conservation of the putative methyltransferase domain: a hallmark of the ‘Sindbis-like’ supergroup of positive-strand RNA viruses. J. Gen. Virol. 73:2129-2134.[Abstract/Free Full Text]
    34. 34
  34. Schwartz, M., J. Chen, M. Janda, M. Sullivan, J. Den Boon, and P. Ahlquist. 2002. A positive-strand RNA virus replication complex parallels form and function of retrovirus capsids. Mol. Cell 9:505-514.[CrossRef][Medline]
    35. 35
  35. Sullivan, M. L., and P. Ahlquist. 1999. A brome mosaic virus intergenic RNA 3 replication signal functions with viral replication protein 1a to dramatically stabilize RNA in vivo. J. Virol. 73:2622-2632.[Abstract/Free Full Text]
    36. 36
  36. Taschner, P. E. M., A. C. van der Kuyl, L. Neeleman, and J. F. Bol. 1991. Replication of an incomplete alfalfa mosaic virus genome in plants transformed with viral replicase genes. Virology 181:445-450.[CrossRef][Medline]
    37. 37
  37. van der Heijden, M. W., J. E. Carette, P. J. Reinhoud, A. Haegi, and J. F. Bol. 2001. Alfalfa mosaic virus replicase proteins P1 and P2 interact and colocalize at the vacuolar membrane. J. Virol. 75:1879-1887.[Abstract/Free Full Text]
    38. 38
  38. van der Kuyl, A. C., K. Langereis, C. J. Houwing, E. M. J. Jaspars, and J. F. Bol. 1990. cis-acting elements involved in replication of alfalfa mosaic virus RNAs in vitro. Virology 176:346-354.[CrossRef][Medline]
    39. 39
  39. Van Pelt-Heerschap, H.,. 1987. Immunochemical analysis of the Alfalfa mosaic virus gene products. Ph.D. thesis. Leiden, The Netherlands.
    40. 40
  40. van Rossum, C. M. A., M. L. Garcia, and J. F. Bol. 1996. Accumulation of alfalfa mosaic virus RNAs 1 and 2 requires the encoded proteins in cis. J. Virol. 70:5100-5105.[Abstract/Free Full Text]
    41. 41
  41. Vasiljeva, L., A. Merits, P. Auvinen, and L. Kääriäinen. 2000. Identification of a novel function of the Alphavirus capping apparatus. J. Biol. Chem. 275:17281-17287.[Abstract/Free Full Text]
    42. 42
  42. Vlot, A. C., L. Neeleman, H. J. M. Linthorst, and J. F. Bol. 2001. Role of the 3'-untranslated regions of alfalfa mosaic virus RNAs in the formation of a transiently expressed replicase in plants and in the assembly of virions. J. Virol. 75:6440-6449.[Abstract/Free Full Text]
    43. 43
  43. Wang, H.-L., J. O'Rear, and V. Stollar. 1996. Mutagenesis of the Sindbis virus nsP1 protein: effects on methyltransferase activity and viral infectivity. Virology 217:527-531.[CrossRef][Medline]
    44. 44
  44. Wang, Y.-F., S. G. Sawicki, and D. L. Sawicki. 1991. Sindbis virus nsP1 functions in negative-strand RNA synthesis. J. Virol. 65:985-988.[Abstract/Free Full Text]

Journal of Virology, November 2002, p. 11321-11328, Vol. 76, No. 22
0022-538X/02/$04.00+0     DOI: 10.1128/JVI.76.22.11321-11328.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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