An important determinant of the ability of Turnip mosaic virus to infect Brassica spp. and/or Raphanus sativus is in its P3 protein

Noriko Suehiro, Tomohide Natsuaki, Tomoko Watanabe and Seiichi Okuda

Faculty of Agriculture, Utsunomiya University, Mine-machi 350, Utsunomiya 321-8505, Japan

Correspondence
Tomohide Natsuaki
natsuaki{at}cc.utsunomiya-u.ac.jp


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Turnip mosaic virus (TuMV, genus Potyvirus, family Potyviridae) infects mainly cruciferous plants. Isolates Tu-3 and Tu-2R1 of TuMV exhibit different infection phenotypes in cabbage (Brassica oleracea L.) and Japanese radish (Raphanus sativus L.). Infectious full-length cDNA clones, pTuC and pTuR1, were constructed from isolates Tu-3 and Tu-2R1, respectively. Progeny virus derived from infections with pTuC induced systemic chlorotic and ringspot symptoms in infected cabbage, but no systemic infection in radish. Virus derived from plants infected with pTuR1 induced a mild chlorotic mottle in cabbage and infected radish systemically to induce mosaic symptoms. By exchanging genome fragments between the two virus isolates, the P3-coding region was shown to be responsible for systemic infection by TuMV and the symptoms it induces in cabbage and radish. Moreover, exchanges of smaller parts of the P3 region resulted in recombinants that induced complex infection phenotypes, especially the combination of pTuC-derived N-terminal sequence and pTuR1-derived C-terminal sequence. Analysis by tissue immunoblotting of the inoculated leaves showed that the distributions of P3-chimeric viruses differed from those of the parents, and that the origin of the P3 components affected not only virus accumulation, but also long-distance movement. These results suggest that the P3 protein is an important factor in the infection cycle of TuMV and in determining the host range of this and perhaps other potyviruses.

The nucleotide and amino acid sequence data reported in this article have been submitted to the DDBJ and assigned the accession numbers AB105134 and AB105135.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Systemic infection of plants occurs when a virus establishes genome amplification, then movement cell-to-cell and over long distances via the plasmodesmata and phloem cells (Carrington et al., 1996; Lucas & Gilbertson, 1994). The factors that influence virus host range, symptomatology and/or pathogenicity have been studied using various virus mutants as well as recombinants constructed from closely related viruses (Rao, 1999). In many cases these factors involve one or more proteins involved in virus replication and/or transport, including coat protein (CP), movement protein and/or other proteins conferring the function necessary for virus movement.

The genus Potyvirus contains more than 200 members or possible members and belongs to the largest plant virus family, Potyviridae (Berger et al., 2000). Potyviruses have flexuous filamentous particles that contain an approximately 10 kb positive-sense single-stranded RNA that is covalently linked to a virus genome-linked protein (VPg) at the 5' end and polyadenylated at the 3' end (Revers et al., 1999). The RNA has a single open reading frame that is translated into a large polyprotein, which is proteolytically cleaved into mature proteins by three virus-encoded proteinases (Riechmann et al., 1992; Urcuqui-Inchima et al., 2001).

The potyvirus P3 protein is proteolytically cleaved from polyprotein by HC-Pro and NIa-Pro, resulting in either the P3 protein or its precursor P3-6K1 (Riechmann et al., 1992). Among potyviruses there is relatively little similarity in P3 proteins compared to other proteins (Urcuqui-Inchima et al., 2001). The P3 proteins are thought to be involved in virus replication (Merits et al., 1999), accumulation (Klein et al., 1994), symptomatology (Chu et al., 1997; Sáenz et al., 2000), resistance breaking (Hjulsager et al., 2002; Jenner et al., 2002, 2003; Johansen et al., 2001) and cell-to-cell movement (Dallot et al., 2001; Johansen et al., 2001).

Turnip mosaic virus (TuMV) is a member of the genus Potyvirus (Berger et al., 2000) and infects cruciferous plants throughout the world (Walsh & Jenner, 2002). Cruciferous plants include a large variety of economically important crops belonging to different genera (Gómez-Campo, 1999). TuMV has adapted to such diversified cruciferous crops, and the many TuMV isolates in the world have been classified into several strains or pathotypes (Fujisawa, 1990; Green & Deng, 1985; Jenner & Walsh, 1996; Provvidenti, 1980; Stavolone et al., 1998; Stobbs & Shattuck, 1989). Recently, many TuMV isolates have been collected around the world and grouped into four lineages by sequencing and phylogenetic analysis of sequences encoding P1 and CP (Ohshima et al., 2002), CP (Sánchez et al., 2003), or the entire polyprotein (Tomimura et al., 2003). These classifications were based on host range, but the factors affecting the virus host range in cruciferous plants are not clear.

In this study we used two Japanese TuMV isolates, Tu-2R1 and Tu-3. Tu-2R1 can systemically infect not only Japanese radish (Raphanus sativus L.) but also Brassica spp., although Tu-2R1 induces very mild symptoms in cabbage (Brassica oleracea L.). On the other hand, Tu-3 infects Brassica spp. but not radish. To identify the genetic determinant that plays a role in systemic infection and symptomatology of TuMV in cabbage and radish, we constructed two infectious full-length cDNA clones and a series of chimeric viruses between them. Examination of the pathogenicity of these chimeric viruses in cabbage and radish showed that the determinant defining the differential infection phenotype is the P3 protein.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Virus isolates.
TuMV isolates Tu-2R1 and Tu-3 were used in this study. Tu-2R1 was isolated from field-grown Japanese radish (R. sativus) and was propagated in turnip (Brassica rapa subsp. rapa) cv. Fuyutoyo (Sakata Seed Co., Yokohama, Japan). Tu-3 was isolated from a diseased cabbage (B. oleracea var. capitata) and maintained in cabbage cv. Haruhikari No. 7 (Takii Seed Co. Ltd, Kyoto, Japan).

Construction of TuMV infectious full-length cDNA clones.
Four cDNA fragments were amplified separately to construct full-length infectious clones of TuMV by immunocapture–reverse transcription–polymerase chain reaction (IC-RT-PCR) (Table 1, fragments 5'-S, S-B, B-X, X-3'). These cDNA fragments were cloned between a Cauliflower mosaic virus (CaMV) 35S promoter and a nopaline synthetase (NOS) terminator in a pUC-based plasmid vector modified from pBI121 (Clontech). First, 5'-S fragments were blunt-ended at the 5' terminus and ligated, without non-virus nucleotides, downstream of the CaMV 35S promoter. In a second step, B-X (position 6130-8435) and X-3' [position 8435-poly(A)] fragments were combined at a XhoI site, the resulting fragments were inserted upstream of the NOS terminator, and finally the S–B fragment was digested with SalI and BamHI, and inserted between the 5'-S and B-3' fragments. To confirm the nucleotide sequence of the clones, named pTuR1 and pTuC, plasmids were digested with a suitable restriction enzyme, subcloned and sequenced.


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Table 1. Oligonucleotide primers used to amplify Tu-2R1 and Tu-3 cDNA

Restriction sites are underlined. Silent mutation sites on Tu-2R1 cDNA are shown in bold.

 
Almost all recombinant clones between pTuR1 and pTuC were prepared using unique corresponding restriction enzyme sites including SalI (position 2491), SnaBI (position 3305) and BamHI (position 6130) in virus cDNA, KpnI upstream of the CaMV 35S promoter, and SmaI downstream of the NOS terminator. The SalI (position 2491) site in pTuR1 was created in oligonucleotide primers (Table 1, Tu-2460F2 and Tu-2480R) used in IC-RT-PCR and had no coding effect.

Virus inoculation and detection.
TuMV cDNA constructs were inoculated to 3-week-old seedlings of turnip cv. Fuyutoyo by the Helios Gene Gun System (Bio-Rad). The bombardment conditions were: microcarrier (gold particles), 0·6, 1·0, or 1·6 µm in diameter; microcarrier loading quantity, 1 mg per shot; DNA loading ratio, 2·0 µg (mg gold)–1, 2·0 µg per shot; final concentration of polyvinylpyrrolidone (PVP), 0·05 mg (ml ethanol)–1; helium pressure 180–200 p.s.i.

At 3 weeks post-inoculation (p.i.) the upper leaves of individual turnip plants showing a systemic mosaic were ground in phosphate buffer (0·1 M, pH 8·0) and mechanically inoculated simultaneously on cotyledons of both cabbage cv. Shikidori and Japanese radish cv. Awashinbansei. In each inoculation test four to six plants were used and the simultaneous inoculation experiments were repeated three or four times. Inoculated plants were grown separately to prevent cross-contamination. Systemic infection was determined by Western blotting analysis at 15 days p.i. The percentage infectivity values were compiled from separate inoculation tests (see Fig. 2). To confirm the validity of chimeric viruses used as inocula, restriction fragment-length polymorphism analysis of IC-RT-PCR products was carried out with restriction enzymes that could differentiate the sequences of parental viruses. Several progeny viruses in systemic leaves of inoculated cabbage and radish plants were also confirmed by sequence analysis.



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Fig. 2. (A) Schematic representation of parental TuMV and relevant chimeric viruses showing their systemic infectivities in cabbage and radish. Segments derived from TuR1 or TuC are depicted as solid or open boxes, respectively. Restriction sites used to generate the chimeric viruses and their positions are indicated below the map. Systemic symptoms were recorded and samples from upper leaves of each plant were collected at 15 days p.i. to be used in Western blotting with an antiserum to TuMV. CS, chlorosis and ringspots; cm, mild chlorotic mottle; M, mosaic; –, no symptoms. Inf. (%), mean percentage of plants with systemic infections detected by Western blotting. (B) Western blotting analysis of accumulation of TuMV CPs in upper leaves from cabbage and radish inoculated with parental viruses and the relevant chimeric viruses 9 to 14 at 15 days p.i.

 
Seeds of the cabbage and radish cultivars were obtained from Takii Seed Co. Ltd. All plants were grown under constant conditions in growth chambers maintained at 14 h per day photoperiod at 22 °C/19 °C (day/night).

IC-RT-PCR.
PCR tubes (0·5 ml, polypropylene) were coated with 50 µl of an anti-TuMV rabbit antiserum diluted 1000-fold in phosphate-buffered saline (PBS), and were incubated at 37 °C for 1 h. After incubation the tubes were washed twice with PBS containing 0·05 % Tween 20 (PBST) and 50 µl plant tissue extract ground at 1 : 3 (w/v) in PBST was added. Then the tubes were incubated at room temperature for 15 min in order to trap virus particles before washing twice with PBST, and 30 µl RNase-free water was added. The tubes were incubated at 95 °C for 1 min, and cooled on ice immediately. From the resulting solution, 4 µl containing released virus RNA was used as the template for cDNA synthesis with M-MLV reverse transcriptase of the First-strand cDNA Synthesis Kit (Amersham Biosciences). cDNA was amplified by PCR using KOD-plus-DNA polymerase (Toyobo) according to the manufacturer's protocol. Primers used in this study are listed in Table 1.

Protein analysis.
Western blotting analysis of TuMV CP in the upper leaves of cabbage and radish was done using a modified protocol of Sambrook et al. (1989). Total protein extracts were tested for the presence and/or accumulation of TuMV CPs. Leaf tissue ground 1 : 6 (w/v) with 1xSDS-PAGE sample buffer [2 % (w/v) SDS, 2 % 2-mercaptoethanol, 0·05 M Tris/HCl pH 6·8] was boiled for 5 min, and centrifuged at 10 000 r.p.m. for 5 min. The supernatant fluid was mixed with 6xgel-loading dye [0·25 % (w/v) bromophenol blue, 0·25 % (w/v) xylene cyanol, 30 % glycerol in H2O]. Samples of 8 µl were separated in a 10 % SDS-polyacrylamide gel and electroblotted onto nitrocellulose membranes (Advantec) using a semi-dry trans-blotter (Nihon Eido). Membranes were then blocked for 30 min in TTBSPB [20 mM Tris/HCl pH 7·5, 500 mM NaCl (TBS) containing 0·05 % Tween 20, 2 % (w/v) PVP and 2 % (w/v) bovine serum albumin]. The membranes were incubated for 1 h at room temperature with an anti-TuMV rabbit antiserum diluted 1 : 1000 in TTBSPB and washed twice before incubation with a goat anti-rabbit IgG-AP conjugate (Bio-Rad) diluted 1 : 3000 in TTBSPB for 1 h at room temperature. The protein–antibody complexes were visualized by incubation with 5-bromo-4-chloro-3-indolyl phosphate and nitro blue tetrazolium.

Distributions of TuMV in cabbage and radish were analysed by tissue immunoblotting analysis (Srinivasan & Tolin, 1992). Briefly, inoculated leaves at 12 days p.i. were pressed between two pieces of 3 mm filter paper (Whatman). Residual green colour was removed from the filter by rinsing in 2 % Triton X-100 prior to blocking with TTBSPB. Detection of virus CP was performed as described above.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Viral progeny of infectious full-length cDNA clones derived from TuMV isolates of Tu-2R1 and Tu-3 exhibit different pathogenicity in B. oleracea and R. sativus
Full-length cDNA clones of Tu-2R1 and Tu-3 (named pTuR1 and pTuC, respectively) were constructed by combining four cDNA fragments. Each clone comprised 9833 nt excluding the poly(A) tail of 18 nt, and contained a single large open reading frame encoding 3164 aa (first initiation codon positioned at 130–132 and terminal codon positioned at 9622–9624). The nucleotide and amino acid sequences of pTuR1 and pTuC were 96 and 97 % identical, respectively. The nucleotide sequences of the 3'-UTRs and the amino acid sequences of 6K1 and 6K2 were 100 % identical between the two clones.

Biolistic inoculation with either pTuR1 or pTuC resulted in all inoculated turnip plants becoming infected. Symptoms appeared 9 to 10 days p.i. and were followed by severe mosaic symptoms.

The progeny viruses derived from pTuC or pTuR1 (hereafter denoted TuC and TuR1, respectively) in turnip plants were used as inoculum sources, and inoculated simultaneously onto cabbage (B. oleracea var. capitata) or Japanese radish (R. sativus). TuC caused systemic chlorosis and ringspots in cabbage plants (Fig. 1B1) but could not be detected in systemic tissue of radish plants (Fig. 1B2). In contrast, TuR1 infection induced mild chlorotic mottle in cabbage (Fig. 1A1) and mosaic symptoms in radish (Fig. 1A2). TuR1 caused a very mild symptom in cabbage, clearly distinguishable from the more severe symptoms provoked by TuC (Fig. 1A1 and B1). Results of Western blotting (Fig. 2) showed that the concentration of TuR1 in cabbage was approximately half that of TuC.



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Fig. 1. Phenotypic comparison of upper leaves of cabbage (B. oleracea var. capitata, A1–G1) and radish (R. sativus, A2–G2) inoculated with the progeny viruses of TuMV infectious clones pTuR1, pTuC and relevant chimeric viruses, and mock-inoculated (G1 and G2). All leaves shown were photographed at 3–4 weeks p.i.

 
Exchanges of the P3 genes alter infection phenotype of TuMV
In order to map the determinants that define the host-specific infection phenotypes, a series of reciprocal chimeric clones between pTuR1 and pTuC was constructed (Fig. 2, chimeras 1 to 14). These chimeric cDNA clones were inoculated into turnip, which is a common susceptible host for the two isolates. All constructs were able to infect turnip systemically, inducing severe mosaic symptoms, and all accumulated to similar levels as determined by Western blotting (data not shown). The crude sap from turnip plants infected with either of the chimeric clones was used as an inoculum.

Initially we exchanged 6130-3' regions of the two parental isolates containing most of NIa and all of NIb, CP and 3'-UTR (Fig. 2), because the sequences encoding host-specific virulence and/or systemic infection determinants have been shown to be in the central region of VPg in other potyviruses (Nicolas et al., 1997; Schaad et al., 1997; Rajamäki & Valkonen, 1999; Borgstrøm & Johansen, 2001). Chimeric virus 1 was able to infect both cabbage and radish. However, the symptoms induced in cabbage were the same as those of TuR1 infection. On the other hand, the reciprocal chimeric virus 2 exhibited a phenotype identical to that of TuC, and did not infect radish systemically. These results indicated that infectivity and symptomatology of the two isolates of TuMV in cabbage and radish did not involve the 6130-3' region.

When chimera 3 and the reciprocal virus 4 were prepared by changing the 5'-2491 region containing all of the 5'-UTR and P1, and almost all of HC-Pro, and were then inoculated, this exchange did not alter the pathogenicity to cabbage and radish. These results indicated that the 5'-2491 regions did not play a significant role in the differential infection phenotype between TuC and TuR1, suggesting that the central region (positions 2491 to 6130) might affect host-specific infection and symptomatology of TuMV. The central region of potyvirus polyproteins has been reported to affect pathogenicity and symptomatology in specific hosts or one plant species (Chu et al., 1997; Dallot et al., 2001; Hjulsager et al., 2002; Jenner et al., 2000, 2002, 2003; Johansen et al., 2001; Sáenz et al., 2000). As expected, the progeny virus 5 caused the same host responses as TuC, and the reciprocal recombinant 6 caused the same host responses as TuR1. These results showed that determinant(s) for host-specific virus infection phenotypes are located in the 2491–6130 region, presumably P3, CI and/or the N terminus of VPg. No difference in amino acid sequences was found between pTuR1 and pTuC for the C terminus of HC-Pro nor in all of 6K1 and 6K2.

Next, the 2491–6130 regions of pTuR1C(2491-6130) and pTuCR1(2491-6130) were divided by XhoI at nucleotide 3680 and four chimeric viruses TuR1C(2491-3680), TuR1C(3680-6130), TuCR1(2491-3680) and TuCR1(3680-6130) (Fig. 2, chimeras 7 to 10) were generated. Chimeric virus 7 had the same infection phenotype as TuR1. Similarly, chimera 8 induced a TuC infection phenotype in cabbage and radish. These results suggested that the 3680–6130 region, which encodes the CI protein and the N-terminal 76 amino acids of VPg, was unable to change TuMV infection phenotypes. The results of inoculation tests of chimeras 9 and 10 clearly indicated that a heterologous 2491–3680 genomic region conferred on TuC the ability of TuR1 to infect cabbage and radish (Fig. 1C1 and C2), and conferred on TuR1 the differential infection phenotype of TuC (Fig. 1D1 and D2). The 2491–3680 region encodes the C terminus of HC-Pro, all of P3 and the N terminus of 6K1, but the only differences in amino acid sequence between pTuR1 and pTuC were in the P3 gene. This suggests that the TuMV P3 gene encodes determinant(s) of host-specific infection phenotype and symptomatology in cabbage and radish.

Recombinant P3 genes cause infection different from those of either parent in Brassica and Raphanus
To locate more precisely the host-specific infection determinant in the P3 gene, the 2491–3680 region was analysed further. Chimeric virus 12, in which the C-terminal TuR1 P3 fragment of TuCR1(2491-3680) was exchanged with the corresponding fragment of TuC using the SnaBI site, induced a host response of TuC-like symptoms in cabbage, but in radish infection could not be detected by Western blotting which could detect virus at concentrations about 2 to 3 % of that reached in infected cabbage (data not shown). The reciprocal virus 11 induced mosaic symptoms in radish (Fig. 1E2) but no symptoms in cabbage (Fig. 1E1), in which no virus was detected by Western blotting (Fig. 2). This infection phenotype was completely different from that of either parental virus. However, a comparison of the results of two sets, chimera 9 with 11 and chimera 10 with 12, showed that the C terminus of the TuMV P3 gene has an important role in systemic infection and/or symptomatology in cabbage and radish. Chimeric virus 13, containing the TuC 3305–3680 fragment in a TuR1 background, induced the same infection phenotype as TuCR1(2491-3305) (Fig. 1F1 and F2). This result indicates that the C-terminal region of the TuC P3 gene was a crucial domain for TuC-like infection. Notably, the reciprocal chimera 14, containing TuR1 3305–3680 region in a TuC background, could not infect either cabbage or radish despite the high virus concentration reached in turnip (data not shown). The P3 genes of chimeras 11 and 14 were identical; however, neither virus was able to infect cabbage, in contrast to the parents TuR1 and TuC. It is possible that the C-terminal region of the TuR1 P3 gene alone is insufficient to induce TuR1-like infection in cabbage, and both the N and C termini of the TuR1 P3 gene are required.

Although the C-terminal region of the TuR1 P3 gene played an important role in the infection and subsequent induction of symptoms by chimeras 11 and 14 in radish, it may depend on background genes other than P3. This behaviour of TuCR1(3305-3680) was different from that of other chimeric viruses (Fig. 2, chimeras 1 to 13).

Tissue immunoblotting analysis of the distribution of TuMV chimeric viruses
Because chimera viruses containing the recombinant P3 molecules can systemically infect turnip, but not cabbage and/or radish, we performed tissue immunoblotting analysis of inoculated cotyledons of cabbage and radish to visualize the extent of infection. Parental and chimeric TuMV (chimeras 9 to 14; constructs are shown in Fig. 2) were detected in both cabbage and radish cotyledons, but their distributions were different at 12 days p.i. (Fig. 3).



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Fig. 3. Tissue immunoblotting analysis of inoculated cotyledons of cabbage (A–D and I–M) and radish (E–H and N–R) at 12 days p.i. Cotyledons were inoculated with TuR1 (D and H), TuC (I and N), or chimeras 9 (A and E), 10 (L and Q), 11 (C and G), 12 (J and O), 13 (B and F), 14 (K and P), or mock-inoculated (M and R). Virus numbers 9–14 correspond to those in Fig. 2. TuC and 9, 11 and 14, 12 and 13, and TuR1 and 10 encode the same P3 component. Halves of radish cotyledons were blotted. Bars, 5 mm. Schematic representation of rectangles between upper and lower panels indicates constructs of P3 gene from the TuR1 and/or TuC genomes.

 
In cabbage, chimeric viruses 11 and 14 were restricted to smaller lesions than those induced by other viruses (Fig. 3B and J). These two viruses encoding the same P3 component were unable to infect cabbage systemically (Fig. 2), suggesting that in this chimera P3 might affect TuMV spread in inoculated cotyledons and/or long-distance movement in cabbage. The other viruses induced systemic symptoms in cabbage and did not differ in their distributions in inoculated leaves (Fig. 3A, C, D, I, K and L). These results indicated that symptom differences in systemic leaves did not result from differences in virus propagation in inoculated cotyledons.

In radish, chimeric virus 10 encoding intact TuR1 P3 was able to distribute throughout inoculated cotyledons, like TuR1 (Fig. 3H and Q). The other viruses containing TuC P3 or chimera P3 showed restricted distributions (Fig. 3E–G and N–P). However, there was a tendency for larger lesions to form when the N terminus of P3 was from TuR1 (Fig. 3G and P). On the other hand, chimeric virus 11 was able to infect radish systemically (Fig. 2) but in inoculated cotyledons localized along the line of lateral veins (Fig. 3F). When the same P3 component was in a TuC background, in chimera 14, the virus distribution was similar to those of TuC and chimera 9 (Fig. 3E, N and O). Taken together, the results suggest that the different parts of TuMV P3 protein affect systemic infection.

Amino acid sequence comparisons among TuMV P3 proteins
When amino acid residues of the P3 protein were compared between TuR1 and TuC, 37 positions were different in the 355 amino acid residues. In its C-terminal region, which is thought to contain all the information necessary for determining TuC-like infection phenotype, 15 amino acid residues were different. To narrow down the candidate sites for differential infection phenotypes, the P3 sequences of the 18 other isolates of TuMV recorded in the database were also aligned and compared using the CLUSTAL W program (Fig. 4). Information on TuMV isolates, original host, country and strain is listed in Table 2. Amino acid differences were found throughout P3, but the most variable region was located between amino acids 200 and 300, containing the site used to generate recombinant P3 (Fig. 4). In this region amino acids at positions 203, 231, 268, 279–280 and 286 (asterisks) were identical among strains able to infect radish (BR strain); positions 268, 279 and 280 were clearly different between BR and non-BR strains (B strain) (Fig. 4, grey shading).



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Fig. 4. Comparative sequences of 355 amino acids of P3 proteins from TuR1, TuC and other isolates of TuMV. Dots indicate amino acid residues identical to consensus sequence. Asterisked and grey-shaded positions indicate that amino acid residues are highly conserved among BR strains and divided into two strains (see Table 2).

 

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Table 2. List and characterization of TuMV isolates used for comparing P3 protein amino acid sequences

 

   DISCUSSION
Top
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
The TuMV isolates TuR1 and TuC differ in host range and symptomatology in cruciferous plants belonging to different genera, Brassica and Raphanus. The results of our study show that the TuMV P3 protein is the factor determining these virus properties. Because the host ranges of most potyviruses are restricted (Hollings & Brunt, 1981), classifications of their strains or pathotypes are based on symptoms of different cultivars in inoculation tests.

In this study all chimeric viruses infected turnip and induced severe symptoms. However, as shown in Fig. 2, chimeric viruses 9 and 10, which contained heterologous P3 genes, had the infection phenotypes of the cognate parental virus. Additional analysis of four chimeras, 11 to 14, containing P3 genes recombined between those of TuC and TuR1, showed complicated infection phenotypes that differed from that of either parental virus. These results suggest that a component of the P3 molecule contains a key determinant of its ability to infect hosts in different genera.

Chimeric viruses 11 to 14 showed very low systemic infectivity in cabbage and/or radish (Fig. 2). Sequence analysis of virus in the systemically infected leaves did not show any amino acid mutations in the exchanged region that could have caused recovery (data not shown). The viruses incapable of full systemic infection were able to propagate in the inoculated leaves, and their distributions in cabbage and radish plants differed (Fig. 3). These facts could be explained by a model in which full systemic infection of a plant involves a race between the rates of virus replication and movement, and the rate of growth of the plant (Dawson & Hilf, 1992). An alternative explanation is the genetic non-uniformity of the cabbage and radish cultivars used in this study. The relative rate of virus spread in inoculated cotyledons was apparently unrelated to systemic infectivities in radish plants (Fig. 3F, G, O and P).

Potyviruses do not encode a dedicated movement protein, but several proteins participate in virus movement functions: HC-Pro (Cronin et al., 1995; Kasschau et al., 1997; Rojas et al., 1997), CI (Carrington et al., 1998), VPg (Nicolas et al., 1997; Schaad et al., 1997) and CP (Dolja et al., 1994, 1995). It has been reported that these proteins are also involved in the ability of viruses to overcome host resistance (Hämäläinen et al., 2000; Jenner et al., 2000; Johansen et al., 2001; Nicolas et al., 1997; Schaad et al., 1997). In the yeast two-hybrid system, potyvirus proteins NIa of Pea seed-borne mosaic virus (PSbMV; Guo et al., 2001) and NIb of Potato virus A (PVA; Merits et al., 1999) have been shown to interact with P3, and in other in vitro assays other PVA proteins were found to interact with P3 (Merits et al., 1999). Potyvirus P3 proteins play a role in virus replication (Merits et al., 1999) and accumulation (Klein et al., 1994), but although an involvement in movement has been proposed (Dallot et al., 2001; Johansen et al., 2001) it remains to be demonstrated. Our results clearly indicate that the P3 protein influences the efficiency of virus spread, especially in long-distance movement. When a combination of the N and C termini of the P3 protein was derived from TuC and TuR1, respectively, the efficiency of systemic invasion was dependent on the other parts as genomic background (Fig. 2, chimeras 11 and 14). This result suggests that the C terminus of TuR1 P3 may facilitate long-distance movement and interact with movement-associated virus proteins directly or indirectly. However, it is not possible to say whether the P3 protein affects either or both steps in replication and cell-to-cell movement, because cell-to-cell spread is the combined result of the level and rate of replication and the rate of cell-to-cell movement. To address this question, it will be necessary to examine virus replication at the single-cell level using protoplasts of cabbage and radish.

Establishment of systemic infection is a complex process, requiring a balance of the rates of replication, cell-to-cell movement and long-distance movement. In each of these phases there are interactions between virus proteins and host components. The P3 protein contains a putative integral transmembrane domain (Rodríguez-Cerezo & Shaw, 1991) and has no RNA-binding activity (Merits et al., 1998). Expression of P3 proteins can have detrimental effects on the growth of Escherichia coli (Merits et al., 1998) or plants (Moreno et al., 1998). Therefore it is reasonable to speculate that the P3 proteins of TuR1 and TuC also interact with host component(s) during the virus infection cycle. Among potyvirus–host relationships, the P3 genes have already been reported as pathogenicity or resistance-breaking determinants. In Plum pox virus (PPV) the P3 gene with 6K1 influenced symptoms on systemic infection hosts Pisum sativum and Nicotiana clevelandii (Sáenz et al., 2000), or infections of plum and peach, both Prunus species (Dallot et al., 2001). In TuMV (Jenner et al., 2002, 2003) and PSbMV (Hjulsager et al., 2002; Johansen et al., 2001) the P3 genes affected infections by virus pathotypes of Brassica napus and P. sativum expressing dominant or recessive resistance, respectively.

If our results were to be attributed to relationships between the P3 proteins of TuR1/TuC and resistance genes in cabbage/radish which mode, dominant or recessive resistance, is conceivable? Dominant resistance alleles are associated with strong mechanisms induced after a recognition event of virus infection, whereas incompletely dominant resistance alleles are associated with mechanisms to constitutively inhibit virus replication or movement (Fraser, 1992). In our study all chimeric viruses were able to multiply in inoculated cotyledons, and no hypersensitive responses were observed in inoculated and systemic leaves. The TuMV P3 protein is an avirulence determinant for an extreme form (immunity) of resistance in B. napus TuRB03 and TuRB04 (Hughes et al., 2003; Jenner et al., 2002). The amino acid positions at 153 and 312 in TuMV P3 are related to the overcoming of resistance in TuRB03 and TuRB04 (Jenner et al., 2002, 2003); however, in TuR1 and TuC both amino acids are the same, isoleucine and phenylalanine (Fig. 4). On the other hand, in Arabidopsis thaliana–potyvirus interactions, dominant genes for the blockage of virus long-distance movement have been identified for Tobacco etch virus (TEV) (Mahajan et al., 1998; Whitham et al., 1999; Chisholm et al., 2001) and for Lettuce mosaic virus (Revers et al., 2003).

Recessive resistance alleles are associated with negative effects, such as being resistant, or a non-host lacking functions essential for the virus infection cycle (e.g. replication and/or movement), or a dominant negative regulator of resistance in susceptible hosts (Fraser, 1992; Revers et al., 1999). Several avirulence determinants for recessive resistances, especially some affecting cell-to-cell or long-distance movement functions of potyviruses, have been identified in P. sativum as the P3 of PSbMV (Johansen et al., 2001), and in several solanaceous species as the VPg of TEV (Schaad et al., 1997), PVA (Hämäläinen et al., 2000) and Tobacco vein mottling virus (Nicolas et al., 1997). Thus, although the potyvirus P3 has been reported to be involved in a gene-for-gene relationship with both dominant and recessive resistance genes, no genetic analysis is available to explain the interactions between TuR1/TuC P3 and host components (genes). However, this does not exclude the possibility that both dominant and recessive resistances are involved in controlling TuMV host range.

Comparisons among the amino acid sequences of P3 proteins of 20 TuMV isolates identified uniform amino acid residues in BR strains able to infect radish (Fig. 4). These are asterisked in Fig. 4 (positions 203, 231, 268, 279–280 and 286) within a most variable region encompassing a site used to construct recombinant P3. These conserved amino acids are not all markedly different between TuR1 and TuC. However, intact TuR1 P3 protein is required for the induction of a TuR1-like infection phenotype (Fig. 2, chimeras 9 to 14), suggesting that these six amino acids might be a core of a determinant for full systemic infection in radish. On the other hand, positions 268 and 279–280 in the C-terminal region of P3 (Fig. 4, grey shading) clearly divided the 20 viruses into two patterns, suggesting that these amino acids might play an important role in characterizing the TuC-like infection phenotype.

A few relevant analyses of potyvirus host-range limitation have been reported. Sáenz et al. (2002) suggested that HC-Pro might be a factor for controlling the host range of PPV, based on the result that transgenic tobacco plants expressing HC-Pro of TEV were able to complement long-distance movement of PPV. In another study, Tóbiás et al. (2001) constructed hybrid viruses by replacing the CP gene in an infectious PPV clone by that of Zucchini yellow mosaic virus, and indicated that CP genes had no effect on host range. It seems likely from our results that the P3 gene is one of the important determinants of potyvirus host range in different genera.


   ACKNOWLEDGEMENTS
 
This work was supported in part by Grants-in-Aid for Scientific Research (12052206) and Special Coordination Funds for Promoting Science and Technology, Leading Research Utilizing Potential of Regional Science and Technology of the Ministry of Education, Culture, Sports, Science and Technology of the Japanese Government. The authors gratefully acknowledge Dr M. A. Mayo for critically reading this manuscript.


   REFERENCES
Top
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Berger, P. H., Barnett, O. W., Brunt, A. A. & 14 other authors (2000). Family Potyviridae. In Virus Taxonomy: Seventh Report of the International Committee on Taxonomy of Viruses, pp. 703–724. Edited by M. H. V. van Regenmortel, C. M. Fauquet, D. H. L. Bishop and 8 others. San Diego: Academic Press.

Borgstrøm, B. & Johansen, I. E. (2001). Mutations in Pea seedborne mosaic virus genome-linked protein VPg alter pathotype-specific virulence in Pisum sativum. Mol Plant Microbe Interact 14, 707–714.[Medline]

Carrington, J. C., Kasschau, K. D., Mahajan, S. K. & Schaad, M. C. (1996). Cell-to-cell and long-distance transport of viruses in plants. Plant Cell 8, 1669–1681.[Free Full Text]

Carrington, J. C., Jensen, P. E. & Schaad, M. C. (1998). Genetic evidence for an essential role for potyvirus CI protein in cell-to-cell movement. Plant J 14, 393–400.[CrossRef][Medline]

Chisholm, S. T., Parra, M. A., Anderberg, R. J. & Carrington, J. C. (2001). Arabidopsis RTM1 and RTM2 genes function in phloem to restrict long-distance movement of tobacco etch virus. Plant Physiol 127, 1667–1675.[Abstract/Free Full Text]

Chu, M., Lopez-Moya, J. J., Llave-Correas, C. & Pirone, T. P. (1997). Two separate regions in the genome of the tobacco etch virus contain determinants of the wilting response of tabasco pepper. Mol Plant Microbe Interact 10, 472–480.[Medline]

Cronin, S., Verchot, J., Haldeman-Cahill, R., Schaad, M. C. & Carrington, J. C. (1995). Long-distance movement factor: a transport function of the potyvirus helper component proteinase. Plant Cell 7, 549–559.[Abstract/Free Full Text]

Dallot, S., Quiot-Douine, L., Sáenz, P., Cervera, M. T., García, J.-A. & Quiot, J.-B. (2001). Identification of Plum pox virus determinants implicated in specific interactions with different Prunus spp. Phytopathology 91, 159–164.

Dawson, W. O. & Hilf, M. E. (1992). Host-range determinants of plant viruses. Annu Rev Plant Physiol Plant Mol Biol 43, 527–555.[CrossRef]

Dolja, V. V., Haldeman, R., Robertson, N. L., Dougherty, W. G. & Carrington, J. C. (1994). Distinct functions of capsid protein in assembly and movement of tobacco etch potyvirus in plants. EMBO J 13, 1482–1491.[Abstract]

Dolja, V. V., Haldeman-Cahill, R., Montgomery, A. E., Vandenbosch, K. A. & Carrington, J. C. (1995). Capsid protein determinants involved in cell-to-cell and long distance movement of tobacco etch potyvirus. Virology 206, 1007–1016.[CrossRef][Medline]

Fraser, R. S. S. (1992). The genetics of plant–virus interactions: implications for plant breeding. Euphytica 63, 175–185.

Fujisawa, I. (1990). Turnip mosaic virus strains in cruciferous crops in Japan. Jpn Agric Res Q 23, 289–293.

Gómez-Campo, C. (1999). Taxonomy. In Biology of Brassica coenospecies, pp. 3–32. Edited by C. Gómez-Campo. Amsterdam: Elsevier Science.

Green, S. K. & Deng, T. C. (1985). Turnip mosaic virus strains in cruciferous hosts in Taiwan. Plant Dis 69, 28–31.

Guo, D., Rajamäki, M.-L., Saarma, M. & Valkonen, J. P. T. (2001). Towards a protein interaction map of potyviruses: protein interaction matrixes of two potyviruses based on the yeast two-hybrid system. J Gen Virol 82, 935–939.[Abstract/Free Full Text]

Hämäläinen, J. H., Kekarainen, T., Gebhardt, C., Watanabe, K. N. & Valkonen, J. P. T. (2000). Recessive and dominant genes interfere with the vascular transport of Potato virus A in diploid potatoes. Mol Plant Microbe Interact 13, 402–412.[Medline]

Hjulsager, C. K., Lund, O. S. & Johansen, I. E. (2002). A new pathotype of Pea seedborne mosaic virus explained by properties of the P3-6K1- and viral genome-linked protein (VPg)-coding regions. Mol Plant Microbe Interact 15, 169–171.[Medline]

Hollings, M. & Brunt, A. A. (1981). Potyviruses. In Handbook of Plant Virus Infections and Comparative Diagnosis, pp. 731–807. Edited by E. Kurstak. Amsterdam: Elsevier/North-Holland.

Hughes, S. L., Hunter, P. J., Sharpe, A. G., Kearsey, M. J., Lydiate, D. J. & Walsh, J. A. (2003). Genetic mapping of novel Turnip mosaic virus resistance gene TuRB03 in Brassica napus. Theor Appl Genet 107, 1169–1173.[CrossRef][Medline]

Jenner, C. E. & Walsh, J. A. (1996). Pathotypic variation in turnip mosaic virus with special reference to European isolates. Plant Pathol 45, 848–856.

Jenner, C. E., Sánchez, F., Nettleship, S. B., Foster, G. D., Ponz, F. & Walsh, J. A. (2000). The cylindrical inclusion gene of Turnip mosaic virus encodes a pathogenic determinant to the brassica resistance gene TuRB01. Mol Plant Microbe Interact 13, 1102–1108.[Medline]

Jenner, C. E., Tomimura, K., Ohshima, K., Hughes, S. L. & Walsh, J. A. (2002). Mutations in Turnip mosaic virus P3 and cylindrical inclusion proteins are separately required to overcome two Brassica napus resistance genes. Virology 300, 50–59.[CrossRef][Medline]

Jenner, C. E., Wang, X., Tomimura, K., Ohshima, K., Ponz, F. & Walsh, J. A. (2003). The dual role of the potyvirus P3 protein of Turnip mosaic virus as a symptom and avirulence determinant in brassicas. Mol Plant Microbe Interact 16, 777–784.[Medline]

Johansen, I. E., Lund, O. S., Hjulsager, C. K. & Laursen, J. (2001). Recessive resistance in Pisum sativum and potyvirus pathotype resolved in a gene-for-cistron correspondence between host and virus. J Virol 75, 6609–6614.[Abstract/Free Full Text]

Kasschau, K. D., Cronin, S. & Carrington, J. C. (1997). Genome amplification and long-distance movement functions associated with the central domain of tobacco etch potyvirus helper component-proteinase. Virology 228, 251–262.[CrossRef][Medline]

Klein, P. G., Klein, R. R., Rodríguez-Cerezo, E., Hunt, A. G. & Shaw, J. G. (1994). Mutational analysis of the tobacco vein mottling virus genome. Virology 204, 759–769.[CrossRef][Medline]

Lucas, W. J. & Gilbertson, R. L. (1994). Plasmodesmata in relation to viral movement within leaf tissues. Annu Rev Phytopathol 32, 387–411.[CrossRef]

Mahajan, S. K., Chisholm, S. T., Whitham, S. A. & Carrington, J. C. (1998). Identification and characterization of a locus (RTM1) that restricts long-distance movement of tobacco etch virus in Arabidopsis thaliana. Plant J 14, 177–186.[CrossRef][Medline]

Merits, A., Guo, D. & Saarma, M. (1998). VPg, coat protein and five non-structural proteins of potato A potyvirus bind RNA in a sequence-unspecific manner. J Gen Virol 79, 3123–3127.[Abstract]

Merits, A., Guo, D., Järvekülg, L. & Saarma, M. (1999). Biochemical and genetic evidence for interactions between potato A potyvirus-encoded proteins P1 and P3 and proteins of the putative replication complex. Virology 263, 15–22.[CrossRef][Medline]

Moreno, M., Bernal, J. J., Jiménez, I. & Rodríguez-Cerezo, E. (1998). Resistance in plants transformed with the P1 or P3 gene of tobacco vein mottling potyvirus. J Gen Virol 79, 2819–2827.[Abstract]

Nicolas, O., Dunnington, S. W., Gotow, L. F., Pirone, T. P. & Hellmann, G. M. (1997). Variations in the VPg protein allow a potyvirus to overcome {nu}a gene resistance in tobacco. Virology 237, 452–459.[CrossRef][Medline]

Ohshima, K., Tanaka, M. & Sako, N. (1996). The complete nucleotide sequence of turnip mosaic virus RNA Japanese strain. Arch Virol 141, 1991–1997.[Medline]

Ohshima, K., Yamaguchi, Y., Hirota, R. & 10 other authors (2002). Molecular evolution of Turnip mosaic virus: evidence of host adaptation, genetic recombination and geographical spread. J Gen Virol 83, 1511–1521.[Abstract/Free Full Text]

Provvidenti, R. (1980). Evaluation of Chinese cabbage cultivars from Japan and the People's Republic of China for resistance to turnip mosaic virus and cauliflower mosaic virus. J Am Soc Hortic Sci 105, 571–573.

Rajamäki, M.-L. & Valkonen, J. P. T. (1999). The 6K2 protein and the VPg of potato virus A are determinants of systemic infection in Nicandra physaloides. Mol Plant Microbe Interact 12, 1074–1081.[Medline]

Rao, A. L. N. (1999). Molecular basis of symptomatology. In Molecular Biology of Plant Viruses, pp. 201–210. Edited by C. L. Mandahar. Norwell: Kluwer Academic.

Revers, F., Le Gall, O., Candresse, T. & Maule, A. J. (1999). New advances in understanding the molecular biology of plant/potyvirus interactions. Mol Plant Microbe Interact 12, 367–376.

Revers, F., Guiraud, T., Houvenaghel, M.-C., Mauduit, T., Le Gall, O. & Candresse, T. (2003). Multiple resistance phenotypes to Lettuce mosaic virus among Arabidopsis thaliana accessions. Mol Plant Microbe Interact 16, 608–616.[Medline]

Riechmann, J. L., Laín, S. & García, J. A. (1992). Highlights and prospects of potyvirus molecular biology. J Gen Virol 73, 1–16.[Medline]

Rodríguez-Cerezo, E. & Shaw, J. G. (1991). Two newly detected nonstructural viral proteins in potyvirus-infected cells. Virology 185, 572–579.[CrossRef][Medline]

Rojas, M. R., Zerbini, F. M., Allison, R. F., Gilbertson, R. L. & Lucas, W. J. (1997). Capsid protein and helper component-proteinase functions as potyvirus cell-to-cell movement proteins. Virology 237, 283–295.[CrossRef][Medline]

Sáenz, P., Cervera, M. T., Dallot, S., Quiot, L., Quiot, J.-B., Riechmann, J. L. & García, J. A. (2000). Identification of a pathogenicity determinant of Plum pox virus in the sequence encoding the C-terminal region of protein P3+6K1. J Gen Virol 81, 557–566.[Abstract/Free Full Text]

Sáenz, P., Salvador, B., Simón-Mateo, C., Kasschau, K. D., Carrington, J. C. & García, J. A. (2002). Host-specific involvement of the HC protein in the long-distance movement of potyviruses. J Virol 76, 1922–1931.[Abstract/Free Full Text]

Sako, N. (1980). Loss of aphid transmissibility of turnip mosaic virus. Phytopathology 70, 647–649.

Sambrook, J., Fritsch, E. F. & Maniatis, T. A. (1989). Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.

Sánchez, F., Wang, X., Jenner, C. E., Walsh, J. A. & Ponz, F. (2003). Strains of Turnip mosaic potyvirus as defined by the molecular analysis of the coat protein gene of the virus. Virus Res 94, 33–43.[CrossRef][Medline]

Schaad, M. C., Lellis, A. D. & Carrington, J. C. (1997). VPg of tobacco etch potyvirus is a host genotype-specific determinant for long-distance movement. J Virol 71, 8624–8631.[Abstract]

Srinivasan, I. & Tolin, S. A. (1992). Detection of three viruses of clovers by direct tissue immunoblotting. Phytopathology 82, 721.

Stavolone, L., Alioto, D., Ragozzino, A. & Laliberté, J.-F. (1998). Variability among turnip mosaic potyvirus isolates. Phytopathology 88, 1200–1204.

Stobbs, L. W. & Shattuck, V. I. (1989). Turnip mosaic virus strains in southern Ontario, Canada. Plant Dis 73, 208–212.

Tóbiás, I., Palkovics, L., Tzekova, L. & Balázs, E. (2001). Replacement of the coat protein gene of plum pox potyvirus with that of zucchini yellow mosaic potyvirus: characterization of the hybrid potyvirus. Virus Res 76, 9–16.[CrossRef][Medline]

Tomimura, K., Gibbs, A. J., Jenner, C. E., Walsh, J. A. & Ohshima, K. (2003). The phylogeny of Turnip mosaic virus; comparisons of 38 genomic sequences reveal a Eurasian origin and a recent ‘emergence’ in east Asia. Mol Ecol 12, 2099–2111.[CrossRef][Medline]

Tomlinson, J. A. & Ward, C. M. (1978). The reactions of swede (Brassica napus) to infection by turnip mosaic virus. Ann Appl Biol 89, 61–69.

Urcuqui-Inchima, S., Haenni, A.-L. & Bernardi, F. (2001). Potyvirus proteins: a wealth of functions. Virus Res 74, 157–175.[CrossRef][Medline]

Walsh, J. A. (1989). Genetic control of immunity to turnip mosaic virus in winter oilseed rape (Brassica napus ssp. oleifera) and the effect of foreign isolates of the virus. Ann Appl Biol 115, 89–99.

Walsh, J. A. & Jenner, C. E. (2002). Turnip mosaic virus and the quest for durable resistance. Mol Plant Pathol 3, 289–300.[CrossRef]

Whitham, S. A., Yamamoto, M. L. & Carrington, J. C. (1999). Selectable viruses and altered susceptibility mutants in Arabidopsis thaliana. Proc Natl Acad Sci U S A 96, 772–777.[Abstract/Free Full Text]

Received 19 November 2003; accepted 12 February 2004.