| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Donald Danforth Plant Science Center, 975 North Warson Road, St Louis, MO 63132, USA
Correspondence
Roger N. Beachy
rnbeachy{at}danforthcenter.org
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
Present address: Fraunhofer USA Center for Molecular Biotechnology, 9 Innovation Way, Newark, DE 19711, USA. 
Present address: Department of Biotechnology, Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita 565-0871, Osaka, Japan.
Present address: W. M. Keck Center for Comparative and Functional Genomics, 329 Edward R. Madigan Laboratory (MC 051), 1201 West Gregory Drive, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA.
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Oparka et al., 1999; Lazarowitz & Beachy, 1999). Infection by Tobacco mosaic virus (TMV) leads to production of a virus-encoded 30 kDa movement protein (MP) that is required for cell-to-cell movement of the complex that contains viral RNA. MP induces a transient modification of Pd, resulting in an increase in size exclusion limit (SEL) of the Pd from 0.5 to over 10 kDa (Deom et al., 1987; Oparka et al., 1997).
Virus replication complexes (VRCs) of many different viruses are associated with one or more cellular organelles, including lysosomal or endosomal membranes (Froshauer et al., 1988; Kujala et al., 2001), peroxisomes (Bleve-Zacheo et al., 1997), mitochondrial outer membranes (Miller et al., 2001), chloroplast envelope (Prod'homme et al., 2003) or vacuolar membranes (van der Heijden et al., 2001; Hasegawa et al., 2003). In the case of TMV, the VRCs assemble on ER membranes. VCRs contain viral RNA, viral replicase and MP plus many unidentified host proteins, including elements of the cell cytoskeleton such as microtubules and actin filaments (Mas & Beachy, 1999; Lazarowitz & Beachy, 1999; Beachy & Heinlein, 2000; Asurmendi et al., 2004). More recently, real-time imaging of TMV infection that monitored the accumulation of MP–green fluorescent protein (GFP) fusion protein provided evidence that TMV infection spreads from cell to cell as intact VRCs (Kawakami et al., 2004). Because of the intimate association of MP with ER and its role in forming VRCs on ER, it is important to characterize the association of MP with ER. The hypothesis that MP contains two transmembrane domains that might anchor the protein (Berna, 1995) was supported by recent studies using purified recombinant MP (Brill et al., 2000, 2004).
In a previous study, deletion mutagenesis was carried out to create an incomplete series of 3 aa deletion (TAD) mutants of the TMV-MP; the purpose of the study was to identify domains of the protein that are involved in its function and subcellular localization (Kahn et al., 1998). The mutations were introduced at intervals of 10 aa throughout the protein (contains 268 aa). TAD 1, for example, has a deletion of aa 9–11; the TAD 2 mutant lacks aa 19–21 and so on. TADs 1–6, 8–16 and 23 were non-functional in this study, whereas TADs 19, 21, 22, 24, 25 and 26 supported cell-to-cell movement of TMV infection. TADs 7, 17, 18 and 20 were not included in this study. When the TAD mutants were fused to GFP, these researchers reported altered patterns of accumulation of MP–GFP fusion proteins on ER (using fluorescence microscopy) for mutants that were non-functional in TMV infection; an exception was TAD 14, which accumulated in a pattern like wild-type (wt) MP–GFP but was not functional.
The current study provides additional information on the association of TAD mutants of MP with cellular membranes. Using several biochemical approaches, confocal microscopy and protein co-localization techniques, other studies from this laboratory confirmed that MP is associated with ER in leaf tissues as well as in protoplasts derived from BY-2 cell cultures (Mas & Beachy, 1998, 1999; Asurmendi et al., 2004). Here, we report that some TAD mutants, including those previously uncharacterized, have profound effects on the association of MP with ER. Possible topology of MP in the ER membrane is discussed based on the results from the TAD mutants and mutants that affect phosphorylation of the MP.
|
METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
). Construction of TAD mutants 1–6 and 8–16 was described previously (Kahn et al., 1998). Additional TAD mutants constructed in the present study were made as follows. Primers with the sequence of the region upstream of AatII sites of TMV (5'-GAGCGGGGACGTCACGACGT-3') and TAD-specific 3' primers (5'-GACGACCAAACAGACGTATCCACTATC-3' for TAD 7 and 5'-CAAACCTAAATTTCTATAAACAATACA-3' for TAD 17) were used to amplify the fragments carrying the deletion. 5' primers partially complementary to the corresponding TAD-specific 3' primers (5'-TACGTCTGTTTGGTCGTCACGGGCGAG-3' for TAD 7 and 5'-TATAGAAATTTAGGTTTGAGAGAGAAG-3' for TAD 17) and a primer with sequence of the region downstream of NcoI site of MP (5'-CCTTTACTCATCTTGAGCTCTAGAAT-3') were used to amplify the fragment carrying the deletion, which annealed to the PCR fragments with the AatII sites described above. A second PCR was then carried out with the two PCR products to obtain fragments that consist of 5' AatII, 3 aa deletion and 3' NcoI site. The PCR product of each TAD was digested with AatII and NcoI and the fragments were cloned into the AatII/NcoI site of pU3/12 to give infectious clones of TMV with TAD mutants. TMV-MP (37A/126S) and TAD4/126S were created by an identical approach. The primers specific for the mutation A126S are 5'-CCAATTATTCTATAACC-3' and 5'-GGTTATAGAATAATTGG-3'. Presence of the deletion and the mutation was confirmed by sequencing the region from AatII and NcoI sites of the resulting TMV clones.
Cell culture and electroporation.
Tobacco BY-2 suspension cells were maintained and prepared for isolation of protoplasts as described elsewhere (Watanabe et al., 1987). BY-2 protoplasts were infected by electroporation with in vitro transcribed RNA from TMV clones using the MegaScript T7 kit (according to the manufacturer's instruction; Ambion). Approximately 106 protoplasts were infected with 1 µg RNA and were incubated at 26 °C without light and harvested after 12–14 h. Protoplasts were collected by centrifugation at low speed (500 g for 4 min) and kept frozen at –20 °C.
Isolation of membrane vesicles.
Frozen samples of protoplasts were thawed on ice, resuspended in 400 µl breaking buffer [250 mM Tris/HCl, pH 8.0, 10 mM KCl, 1 mM EDTA, protease inhibitor mix (Sigma P-9599, 1 : 100 dilution into the buffer), 10 µg ALLM (Sigma A-6060) ml–1, 25 µM MG115 (Sigma C-6706)] and lysed by passing suspended cells through a 26G needle twice. Cell lysates were centrifuged at 4000 g for 5 min. The supernatant was then centrifuged at 100 000 g for 20 min. The 100 000 g supernatant was labelled ‘S’ and the pellet (P) was resuspended in buffer (equal to the original volume) containing 50 mM Tris/HCl, pH 8.0, 10 mM KCl, 1 mM EDTA. Equal volumes of each sample (S and P) were mixed with SDS-PAGE sample buffer and boiled for 4 min immediately prior to electrophoresis. For discontinuous sucrose gradients, protoplasts were lysed in the breaking buffer with either 1 mM EDTA (–Mg2+) or 5 mM MgCl2 (+Mg2+). The 100 000 g P was resuspended in 50 mM Tris/HCl, pH 8.0, 10 mM KCl and 1 mM EDTA (–Mg2+) or 5 mM MgCl2 (+Mg2+). Membrane samples were loaded onto discontinuous sucrose gradients with blocks of 40, 50, 60 and 70 % (w/v) prepared in 50 mM Tris/HCl, pH 8.0, 10 mM KCl and 1 mM EDTA (–Mg2+) or 5 mM MgCl2 (+Mg2+). Gradients were centrifuged for 2 h at 100 000 g and the interfaces of the sucrose blocks were collected. Collected membranes were analysed by SDS-PAGE and Western blot assays.
SDS-PAGE and Western blot analyses.
Protein samples were separated on standard SDS-PAGE (12.5 % acrylamide) gel (Laemmli, 1970) and subsequently transferred to nitrocellulose membrane (Protran BA83; Schleicher & Schuell). After transfer, membranes were probed with polyclonal rabbit anti-MP antibodies. MP was detected using horseradish peroxidase-conjugated goat anti-rabbit antibodies and the SuperSignal West Dura Extended Duration substrate (Pierce). Intensity of each band on the film was quantified using 1D gel analysing software (Phoretix 1D; Nonlinear Dynamics) after scanning the film with a Umax Astra 600S (UMAX Technologies).
Plant materials.
To confirm infectivity of virus constructs, in vitro transcribed RNA (described above) was inoculated to the hypersensitive hosts Nicotiana tabacum cv Xanthi NN and transgenic Xanthi NN line 2005. Plant line 2005 produces wt TMV-MP (Deom et al., 1991). The infectivity of the mutants was determined by the presence of local lesions after 5–7 days.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
. TAD mutants of MP used in this study are indicated by bold letters. The model is based upon the results of experiments showing that the C terminus of MP in ER-containing vesicles is susceptible to digestion by protease (Reichel & Beachy, 1998) and on physical analyses of recombinant MP produced in Escherichia coli (Brill et al., 2000, 2004). The model predicts that MP is an integral ER membrane protein with the N and C termini exposed to the cytoplasm and approximately 70 aa in the lumen of ER (Brill et al., 2000). This membrane topology was also suggested when the TMV-MP sequence was subjected to prediction by TMpred (http://www.ch.embnet.org; Hofmann & Stoffel, 1993), the algorithm based on the statistical analysis of a database of naturally occurring transmembrane proteins. Based on this model, we conducted a series of experiments to investigate the impact of TAD mutations on association of MP with ER.
|
demonstrated that TMV-MP is associated with rough ER isolated from infected tobacco leaves. Prior to initiating studies on the TAD mutants in BY-2 protoplasts, we confirmed that the protocol used in the previous study gave similar results. BY-2 protoplasts were infected with TMV that encodes MP–GFP fusion protein (TMV-MP–GFP) and at 15 h post-inoculation (p.i.) the microsomal vesicles collected in P at 100 000 g were isolated and subjected to centrifugation through sucrose step gradients constructed with 40, 50, 60 and 70 % sucrose. As shown in Fig. 2, in the presence of Mg2+, MP–GFP was detected throughout the gradient, as was BiP, an ER-resident protein. When Mg2+ was omitted from the lysis buffer and other solutions, the ER membrane was highly enriched at the interface of 40 and 50 % sucrose (Fig. 2; anti-BiP). Similarly, most of the MP–GFP was enriched at the 40/50 interface in the absence of Mg2+, indicating that the protein was associated with rough ER in BY-2 protoplasts. This shift was reproduced when the BY-2 protoplasts were infected with TMV with the full-length MP without GFP (data not shown). Therefore, we concluded that MP recovered in the 100 000 g P (microsomal fraction) was associated with rough ER in BY-2 protoplasts. This was in good agreement with its subcellular localization in tobacco plants (Reichel & Beachy, 1998). Because of the ease of achieving near synchrony of infection and of preparing membrane fractions from protoplasts, we used BY-2 protoplasts to study the effects of TAD mutations on association with ER.
|
demonstrated that TAD mutants 1–6 and 8–16 were not functional in cell-to-cell spread of TMV infection. Other mutants affected the subcellular location of MP–GFP fusion proteins. To complete the set of TAD mutants, TAD 7 and TAD 17 mutants were constructed. TAD 7 and TAD 17 are of significant interest in the present study because of their proximity to the predicted transmembrane sequences (Fig. 1
). To determine whether or not TAD 7 and TAD 17 mutants support cell-to-cell spread of infection, in vitro transcripts of TMV clones that contain the mutants were inoculated to leaves of N. tabacum Xanthi NN: no infection was observed in either case. When transcripts were inoculated to leaves of transgenic line 2005, which accumulates wt MP, necrotic local lesions were developed within 5 days of infection for both mutants, as anticipated (data not shown). These results indicate that the viruses that encode TAD 7 or TAD 17 mutants of MP are capable of replication but are unable to move from cell to cell in wt tobacco plants.
Association of TAD mutants with cellular membranes
In consideration of the model in Fig. 1, we predicted that deletions within or near domains that are responsible for membrane insertion or association would affect the interaction(s) of the protein with membrane. Fluorescence microscopy was used in a study by Kahn et al. (1998) to establish subcellular localization of selected TAD–GFP fusion proteins and confirmed that mutations can affect membrane association. It was pointed out, however, that some of the TAD–GFP fusion proteins were less stable than others (Kahn et al., 1998). Therefore, we chose to use non-fused MP and TAD mutants to avoid possible artefacts that may be associated with GFP fusion proteins. We chose to use biochemical analyses similar to those reported by Laporte et al. (2003) to address the physical interaction of MP with ER.
BY-2 protoplasts were infected with in vitro transcribed TMV RNA that encodes wt MP or TAD mutants, and at 12–14 h p.i. cells were collected. Previous studies indicated that during this period MP accumulated in VRCs but was not associated with microtublules; the latter is a late event in TMV infection (Heinlein et al., 1998; Mas & Beachy, 1999). Protoplast lysates were centrifuged for 5 min at 4000 g. The amounts of MP recovered in the 100 000 g S and P (microsomal fraction) from the 4000 g S were analysed by Western blot using anti-MP antibody (Fig. 3).
|
, wt MP lanes) as anticipated (Fig. 2). Similar studies were conducted with TMV that encodes TAD mutants 1–17. MP present in S represents soluble protein, i.e. having lost interaction with the membrane, while MP recovered in P presumably represents MP associated with ER membranes.
These studies were repeated at least twice and up to five times for each TAD mutant. Representative results for each TAD mutant are shown in Fig. 3, and it is readily evident that some mutant proteins partition in the S and P fractions similar to wt MP, while others are significantly different. For example, TAD 2 exhibited a similar degree of association with microsomes as does wt MP, while TAD 6 and TAD 16 were recovered mostly in the S fraction. To gain more quantitative information from these studies, the blots were scanned and the amount of bound antibody was determined in the S and P fractions. The data are presented as percentages of total MP (S+P) recovered in the P fraction (Fig. 4). TAD mutations 1, 2 and 3 had little impact on membrane association, and more than 70 % of MP was recovered in the P fraction, whereas TAD 4 showed less than 40 % of the protein in the P fraction. TAD 5 had a modest effect on membrane association, but more than 60 % of the protein was recovered in the P fraction. TAD mutants 6 and 7, which are within the first predicted transmembrane domain (Fig. 1), severely affected association with the membrane and less than 40 % of MP remained associated with the P fraction.
|
, aa 82–149) had mixed effects on membrane association of the protein. TAD mutations 8 and 9 resulted in less than 50 % of the proteins recovered with the P fraction. TAD mutations 12 through 15 resulted in more than 70 % of MP in the P fraction. TAD 16, which is located within the predicted second transmembrane domain (Fig. 1), resulted in less than 20 % of MP in the P fraction, indicating loss of membrane association. Finally, TAD 17, which is predicted to be near the cytosolic side of the membrane (Fig. 1) also affected membrane association, and less than 50 % of the protein remained associated with the membrane fraction.
The methods used here would not distinguish whether the TAD mutants were recovered in P as a consequence of protein aggregation versus physical interaction with the membrane. Nevertheless, it is clear that TAD mutations 4, 6, 7, 8, 9, 16 and 17 resulted in more than 50 % of the protein partitioning to the soluble fraction, suggesting that these deletions had strong impacts on interaction of MP with the ER membrane. The effect of the TAD mutations on membrane interaction did not appear to be caused by significant partial or complete degradation of the protein since Western blot analyses showed that all proteins had an apparent molecular mass of the predicted size (Fig. 3).
Effect of phosphorylation on membrane association
Phosphorylation of tomato mosaic tobamovirus (ToMV) MP is required for its function in cell-to-cell spread of virus infection. Substitution of one of the sites of phosphorylation, Ser at position 37 with Ala (S37A), prevented phosphorylation at aa 37, and resulted in dysfunctional MP that no longer supported cell-to-cell movement of infection (Kawakami et al., 1999). Furthermore, when the S37A mutation was introduced in an MP–GFP fusion protein, fluorescence was diffuse throughout the cytoplasm of infected cells. The authors concluded that the mutant MP–GFP was not associated with specific cellular structures. Kawakami et al. (2003) subsequently identified several functional revertants of the S37A mutation. One of these has a single amino acid substitution at Gly126 to Ser (G126S). The ToMV-MP with S37A and G126S is phosphorylated and provides partial recovery of cell-to-cell spread of infection. To determine the effect of MP phosphorylation on membrane association, the membrane partitioning assay was carried out after infection of BY-2 protoplasts with ToMV RNA carrying the S37A mutation in MP. In this experiment more than 90 % of the mutant MP was in the S fraction (Fig. 5, lanes 1, 2 and 3). This is consistent with the previous observation of distribution of MP-S37A–GFP fusion protein (Kawakami et al., 1999). In infections that produced MP-S37A that contained G126S, the ratio of MP in the S and P fractions was changed compared with MP-S37A. As shown in Fig. 5 (lanes 4, 5 and 6), the revertant MP was equally partitioned in the P and S fractions and indicates that the second site mutation partially restores membrane association. This coincides with the report that infectivity of the S37A/G126S mutant is partially restored relative to wt virus (Kawakami et al., 2003).
|
, lanes 7–12). However, TMV carrying the S37A/A126S double mutation did not produce infection when inoculated to Xanthi NN plants (data not shown), unlike the result with the double mutant of ToMV. We did not determine whether the double mutant protein is phosphorylated.
TAD mutants in the membrane fraction
In the case of non-functional mutants that retained partial or complete association with membranes, it is possible that interaction with the membrane is different from the membrane association of wt MP. To address this possibility, we treated representative TAD mutants with trypsin to assess susceptibility of MP to digestion compared with wt MP (Fig. 6). The P fractions collected from protoplasts infected with wt MP, TAD 12 or TAD 17 were treated to incomplete digestion with trypsin after which they were analysed by Western blot assays using anti-MP antibodies. Treatment of samples containing wt MP produced a primary proteolytic fragment of approximately 19 kDa (Fig. 6, asterisk). In contrast, protease treatment of the P fraction of TAD 12 (70 % of TAD 12 is in the P fraction) did not produce comparable amounts of the fragment. In the case of TAD 17, which caused more than 50 % of the protein to be in the S fraction, the protein in the P fraction was susceptible to protease treatment and little or no 19 kDa peptide was produced. These data suggest that at least some of the TAD mutants that remain in the membrane fraction are associated with the membrane in a manner that is different from wt MP.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
). As shown in Fig. 2, TMV-MP associates with ER in BY-2 protoplasts; similar results were obtained from studies in tobacco leaf tissue during the early to mid stages of infection cycle (Reichel & Beachy, 1998). Since the focus in our current study is on MP interaction with ER membranes, the data confirm the legitimacy of using BY-2 protoplasts for such studies.
The results from studies of the TAD mutants of the TMV-MP show that these mutations affect the subcellular localization of MP and support a model of MP as an integral or tightly associated membrane protein. Previous studies reported that MP was not released from cellular membranes by treatments that included high concentrations of NaCl or urea (Reichel & Beachy, 1998), indicating a high degree of interaction of MP with membranes. The results of the studies of the TAD mutants described here are largely consistent with studies of TAD–GFP fusion proteins (Kahn et al., 1998). For example, GFP fusions of TAD 2, 3 and 14 showed subcellular distributions similar to wt MP–GFP; in the present study TAD 2, 3 and 14 accumulated in the P fraction as does wt MP, indicating that the mutants retained the ability to interact with ER membranes. Similarly TAD 5–GFP was found in Pd (these structures contain ER) as well as in cortical bodies; more than 60 % of TAD 5 was recovered in the P fraction (Fig. 4). TAD mutants 6 and 16 were recovered in the S fraction in our current study; TAD 6–GFP and TAD 16–GFP showed diffused distribution, indicating lack of association with ER (Kahn et al., 1998).
In contrast, TAD mutants 12 and 13 showed quite different subcellular localizations in the two studies. Kahn et al. (1998) reported that TAD 12–GFP and TAD 13–GFP were diffuse in the cytosol. However, the Western blot analysis revealed significant degradation of the fusion protein, which may explain the diffused fluorescence (Kahn et al., 1998). We repeated the study of TAD 12–GFP using the sucrose-step gradient assay and found that the majority of GFP was at or near the top of the gradient (data not shown), indicating lack of association with membranes. In contrast, the analyses of the S and P fractions isolated from BY-2 protoplasts infected with TMV that encoded TAD 12 or TAD 13 revealed that each mutant was associated with the P fraction (Fig. 4). We conclude that the biochemical approach used in the present study provides more quantification of the effects of mutation on MP than the study of MP–GFP fusion proteins.
The TAD mutants that had the most significant effects on partitioning of MP in the S and P fractions are clustered around the predicted transmembrane domains (compare Figs 1 and 4). TAD 4 (predicted to be in a cytosolic domain of MP) and TAD 9 (predicted to be in the lumen of the ER) also reduced membrane association: interestingly, both deletions are in regions predicted to be weakly hydrophobic (Fig. 4b). Indeed, there is a strong correlation between the effect of mutations in areas predicted to be hydrophobic in nature and membrane association of MP.
TAD 15 is proximal to the second predicted membrane-spanning domain but does not destroy its association with the P fraction, in contrast to TAD 8. The differences between TAD 8 and TAD 15 may imply that the actual amino acids that were deleted contributed to anchoring MP to ER.
A number of different algorithms are used to predict transmembrane helices from primary amino acid sequences, often yielding different results (Cserzo et al., 1997; Krogh et al., 2001; Sonnhammer et al., 1998; Tusnády & Simon, 2001; Rost et al., 2004). Other plant virus MPs have been accurately predicted to have transmembrane helices. For example, the 9 kDa MP from carmovirus Carnation mottle virus was predicted to have transmembrane domains and was biochemically confirmed to be an ER membrane protein (Vilar et al., 2002). Recent studies showed that p9 is integrated into ER membranes by an signal recognition particle-dependent mechanism (Saurì et al., 2005). Some algorithms used in the present studies, including HMMTOP (Tusnády & Simon, 2001) and TMHMM (Krogh et al., 2001), did not predict that MP sequences are typical of an integral MP; a third algorithm, TMpred (Hofmann & Stoffel, 1993), predicted membrane association.
A survey of the 30 kDa superfamily of viral MPs predicted that the core structure consists of a series of 
-elements flanked by an 
-helix at each end (Melcher, 2000). The experimental data presented here showed that mutations in regions that are predicted to be hydrophobic disrupt association of MP with a membrane fraction. Therefore, one may conclude that such regions are involved in anchoring MP to the membrane rather than being integral membrane regions. Nevertheless, the predicted transmembrane sequences represented by aa 59–81 and 149–170 were critically important for association with the P fraction. These results may indicate that membrane association involves specific interactions with a membrane-associated feature or component that anchors MP to the membrane.
Kawakami et al. (2003) reported that mutations of Ser37 to Ala in MP of TMV and ToMV removed a site of phosphorylation of MP, and destroyed the capability of MP to function in cell-to-cell spread of infection. These mutant MPs did not partition in the P fractions in the present study. Amino acid residues Ser258, Thr261 and Ser265 were shown to be phosphorylated in another study (Citovsky et al., 1993). Although their role in MP interaction with Pd was suggested (Waigmann et al., 2000), the importance of the phosphorylation of these residues is not clear since MP from which C-terminal 55 aa (from aa 214 to 268) were deleted retained biological function (Berna et al., 1991; Gafny et al., 1992). Nevertheless, the role of phosphorylation of other amino acids may affect MP functions that are not yet described.
A second site mutation in MP that resulted in a double mutant, S37A/G126S, results in MP that is more stable than the S37A mutant, is partially functional and is associated with the P (membrane) fraction in the present studies. Kawakami et al. (2003) suggested that the G126S mutation restored the function of MP-S37A by increasing the stability of the protein. The present study suggests that the apparent increase of stability of MP is due to association with cellular membranes.
Since TAD mutants can be associated with membrane but not function in cell-to-cell spread of infection, the results support the hypothesis that there are multiple functional domains in MP. Boyko et al. (2000) identified a domain that is involved in interaction of MP with microtubules. It is likely that TAD mutations can affect a number of activities/functions of MP, including: (i) targeting MP to the ER membrane; (ii) insertion and retention in the membrane; (iii) anchoring of MP with actin/myosin during movement of VRCs in the cell (e.g. Kawakami et al., 2004); (iv) anchoring of VRCs to ER (e.g. Kawakami et al., 2004); and (v) ubiquitination and degradation of MP (e.g. Reichel & Beachy, 1998).
The experiments reported here were carried out in protoplasts for technical reasons, and thus do not address the relationship between association of MP with ER and accumulation in Pd. Kawakami et al. (1999, 2003) reported that mutations that eliminate phosphorylation of aa 37 reduced membrane association and accumulation in Pd. The correlation between association of TAD mutants with cellular membranes and accumulation in Pd can be made by comparing the results of the current studies with those reported by Kahn et al. (1998). In some TAD mutants the correlation is positive, while in others the correlation is not complete. TAD mutants 2, 3, 5 and 14 are largely associated with ER (Fig. 4) and TAD–GFP fusion proteins with these TADs accumulate in Pd; in contrast TAD 12 and 13 accumulate on ER but do not accumulate in Pd. However, TAD–GFP fusion proteins of TAD 12 and 13 are unstable (Kahn et al., 1998; Fig. 4), thus mitigating some of the apparent differences in the results of these studies. In general, there is a strong positive correlation between association with ER and accumulation in Pd.
In summary, the results from the membrane association assays demonstrate a correlation between the predicted transmembrane and hydrophobic domains of MP and association with membranes. Further investigations of the interactions of MP with cellular membranes using MP mutants should lead to more complete understanding of the role of MP in virus infection and cell-to-cell spread.
|
ACKNOWLEDGEMENTS |
|---|
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
Beachy, R. N. & Heinlein, M. (2000). Role of P30 in replication and spread of TMV. Traffic 1, 540–544.[CrossRef][Medline]
Berna, A. (1995). Involvement of residues within putative -helix motifs in the behavior of the alfalfa and tobacco mosaic virus movement proteins. Phytopathology 85, 1441–1448.
Berna, A., Gafny, R., Wolf, S., Lusas, W. J., Holt, C. A. & Beachy, R. N. (1991). The TMV movement protein: role of the C-terminal 73 amino acids in subcellular localization and function. Virology 182, 682–689.[CrossRef][Medline]
Bleve-Zacheo, T., Rubino, L., Melillo, M. T. & Russo, M. (1997). The 33K protein encoded by cymbidium ringspot virus localizes to modified peroxisomes of infected cells and of uninfected transgenic plants. J Plant Pathol 79, 197–202.
Boyko, V., van der Laak, J., Ferralli, J., Suslova, E., Kwon, M.-O. & Heinlein, M. (2000). Cellular targets of functional and dysfunctional mutants of tobacco mosaic virus movement protein fused to green fluorescent protein. J Virol 74, 11339–11346.
Brill, L. M., Nunn, R. S., Kahn, T. W., Yeager, M. & Beachy, R. N. (2000). Recombinant tobacco mosaic virus movement protein is an RNA-binding, -helical membrane protein. Proc Natl Acad Sci U S A 97, 7112–7117.
Brill, L. M., Dechongkit, S., DeLaBarre, D., Stroebel, J., Beachy, R. N. & Yeager, M. (2004). Dimerization of recombinant tobacco mosaic virus movement protein. J Virol 78, 3372–3377.
Citovsky, V., McLean, B. G., Zupan, J. R. & Zambryski, P. (1993). Phosphorylation of tobacco mosaic virus cell-to-cell movement protein by a developmentally regulated plant cell wall-associated protein kinase. Genes Dev 7, 904–910.
Cserzo, M., Wallin, E., Simon, I., von Heijne, G. & Elofsson, A. (1997). Prediction of transmembrane -helices in prokaryotic membrane proteins: the dense alignment surface method. Protein Eng 10, 673–676.
Deom, C. M., Oliver, M. J. & Beachy, R. N. (1987). The 30-kilodalton gene product of tobacco mosaic virus potentiates virus movement. Science 237, 389–394.
Deom, C. M., Wolf, S., Holt, C. A., Lucas, W. J. & Beachy, R. N. (1991). Altered function of the tobacco mosaic virus movement protein in a hypersensitive host. Virology 180, 251–256.[CrossRef][Medline]
Eisenberg, D., Wilcox, W. & McLachlan, A. D. (1986). Hydrophobicity and amphiphilicity in protein structure. J Cell Biochem 31, 11–17.[CrossRef][Medline]
Froshauer, S., Kartenbeck, J. & Helenius, A. (1988). Alphavirus RNA replicase is located on the cytoplasmic surface of endosomes and lysosomes. J Cell Biol 107, 2075–2086.
Gafny, R., Lapidot, M., Berna, A., Holt, C. A., Deom, C. M. & Beachy, R. N. (1992). Effects of terminal deletion mutations on function of the movement protein of tobacco mosaic virus. Virology 187, 499–507.[CrossRef][Medline]
Hagiwara, Y., Komoda, K., Yamanaka, T., Tamai, A., Meshi, T., Funada, R., Tsuchiya, T., Naito, S. & Ishikawa, M. (2003). Subcellular localization of host and viral proteins associated with tobamovirus RNA replication. EMBO J 22, 344–353.[CrossRef][Medline]
Heinlein, M., Padgett, H. S., Gens, J. S., Pickard, B. G., Casper, S. J., Epel, B. L. & Beachy, R. N. (1998). Changing patterns of localization of the tobacco mosaic virus movement protein and replicase to the endoplasmic reticulum and microtubules during infection. Plant Cell 10, 1107–1120.[Medline]
Hofmann, K. & Stoffel, W. (1993). TMbase – a database of membrane spanning proteins segments. Biol Chem Hoppe-Seyler 374, 166.
Holt, C. A. & Beachy, R. N. (1991). In vivo complementation of infectious transcripts from mutant tobacco mosaic virus cDNAs in transgenic plants. Virology 181, 109–117.[CrossRef][Medline]
Kahn, T. W., Lapidot, M., Heinlein, M., Reichel, C., Cooper, B., Gafny, R. & Beachy, R. N. (1998). Domain of the TMV movement protein involved in subcellular localization. Plant J 15, 15–25.[CrossRef][Medline]
Kawakami, S., Padgett, H. S., Hososkawa, D., Okada, Y., Beachy, R. N. & Watanabe, Y. (1999). Phosphorylation and/or presence of serine 37 in the movement protein of tomato mosaic tobamovirus is essential for intracellular localization and stability in vivo. J Virol 73, 6831–6840.
Kawakami, S., Hori, K., Hosokawa, D., Okada, Y. & Watanabe, Y. (2003). Defective tobamovirus movement protein lacking wild-type phosphorylation sites can be complemented by substitutions found in revertants. J Virol 77, 1452–1461.[Medline]
Kawakami, S., Watanabe, Y. & Beachy, R. N. (2004). Tobacco mosaic virus infection spreads cell to cell as intact replication complexes. Proc Natl Acad Sci U S A 101, 6291–6296.
Krogh, A., Larsson, B., von Heijne, G. & Sonnhammer, E. L. L. (2001). Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J Mol Biol 305, 567–580.[CrossRef][Medline]
Kujala, P., Ikaheimonen, A., Ehsani, N., Vihinen, H., Auvinen, P. & Kaariainen, L. (2001). Biogenesis of the Semliki Forest virus RNA replication complex. J Virol 75, 3873–3884.
Kyte, J. & Doolittle, R. (1982). A simple method for displaying the hydropathic character of a protein. J Mol Biol 157, 105–132.[CrossRef][Medline]
Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685.[CrossRef][Medline]
Laporte, C., Vetter, G., Loudes, A.-M., Robinson, D. G., Hillmer, S., Stussi-Garaud, C. & Ritzenthaler, C. (2003). Involvement of secretory pathway and the cytoskeleton in intracellular targeting and tubule assembly of Grapevine fanleaf virus movement protein in tobacco BY-2 cells. Plant Cell 15, 2058–2075.
Lazarowitz, S. G. & Beachy, R. N. (1999). Viral movement proteins as probes for intracellular and intercellular trafficking in plants. Plant Cell 11, 535–548.
Mas, P. & Beachy, R. N. (1998). Distribution of TMV movement protein in single living protoplasts immobilized in agarose. Plant J 15, 835–842.[CrossRef]
Mas, P. & Beachy, R. N. (1999). Replication of tobacco mosaic virus on endoplasmic reticulum and role of the cytoskeleton and virus movement protein in intracellular distribution of viral RNA. J Cell Biol 147, 945–958.
Melcher, U. (2000). The ‘30K’ superfamily of viral movement proteins. J Gen Virol 81, 257–266.
Miller, D. J., Schwarts, M. D. & Ahlquist, P. (2001). Flock house virus RNA replicates on outer mitochondrial membranes in Drosophila cells. J Virol 75, 11664–11676.
Oparka, K. J., Prior, D. A., Santa Cruz, S., Padgett, H. S. & Beachy, R. N. (1997). Gating of epidermal plasmodesmata is restricted to the leading edge of expanding infection sites of tobacco mosaic virus (TMV). Plant J 12, 781–789.[CrossRef][Medline]
Oparka, K. J., Roberts, A. G., Boevink, P. & 7 other authors (1999). Simple, but not branched, plasmodesmata allow the nonspecific trafficking of proteins in developing tobacco leaves. Cell 97, 743–754.[CrossRef][Medline]
Prod'homme, D., Jakubiec, A., Tournier, V., Drugeon, G. & Jupin, I. (2003). Targeting of the turnip yellow mosaic virus 66K replication protein to the chloroplast envelope is mediated by the 140K protein. J Virol 77, 9124–9135.
Reichel, C. & Beachy, R. N. (1998). Tobacco mosaic virus infection induces severe morphological changes of the endoplasmic reticulum. Proc Natl Acad Sci U S A 95, 11169–11174.
Rost, B., Yachdav, G. & Liu, J. (2004). The PredictProtein server. Nucleic Acids Res 32, Web Server Issue W321–W326.
Saurì, A., Saksena, S., Salgado, J., Johnson, A. E. & Mingarro, I. (2005). Double-spanning plant viral movement protein integration into the endoplasmic reticulum membrane is signal recognition particle-dependent, translocon-mediated, and concerted. J Biol Chem 280, 25907–25912.
Sonnhammer, E. L. L., von Heijne, G. & Krogh, A. (1998). A hidden Markov model for predicting transmembrane helices in protein sequences. Proc Int Conf Intell Syst Mol Biol 6, 175–182.[Medline]
Staehelin, L. A. (1997). The plant ER: a dynamic organelle composed of a large number of discrete functional domains. Plant J 11, 1151–1165.[CrossRef][Medline]
Tusnády, G. E. & Simon, I. (2001). The HMMTOP transmembrane topology prediction server. Bioinformatics 17, 849–850.
Van der Heijden, M. W., Carette, J. E., Reinhoud, P. J., Haegi, A. & Bol, J. F. (2001). Alfalfa mosaic virus replicase proteins P1 and P2 interact and colocalize at the vacuolar membrane. J Virol 75, 1879–1887.
Vilar, M., Saurí, A., Monné, M., Marcos, J. F., von Heijne, G., Pérez-Payá, E. & Mingarro, I. (2002). Insertion and topology of a plant viral movement protein in the endoplasmic reticulum membrane. J Biol Chem 277, 23447–23452.
Waigmann, E., Chen, M. H., Bachmaier, R., Ghoshroy, S. & Citovsky, V. (2000). Regulation of plasmodesmal transport by phosphorylation of tobacco mosaic virus cell-to-cell movement protein. EMBO J 19, 4875–4885.[CrossRef][Medline]
Watanabe, Y., Meshi, T. & Okada, Y. (1987). Infection of tobacco protoplasts with in vitro transcribed tobacco mosaic virus RNA using an improved electroporation method. FEBS Lett 219, 65–69.
Received 10 February 2006;
accepted 12 April 2006.
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
| J MED MICROBIOL | ALL SGM JOURNALS | |
