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Department of Applied Chemistry and Microbiology, Latokartanonkaari 11, PO Box 27, University of Helsinki, FIN-00014 Helsinki, Finland
Correspondence
Kristiina Mäkinen
kristiina.makinen{at}helsinki.fi
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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), and different host factors participate in the process (Ahlquist et al., 2003; Whitham & Wang, 2004). Numerous studies have described the importance of post-translational modifications in regulating viral protein function during host–virus interplay, including phosphorylation, glycosylation and lipid modifications. The characterization of host–virus communication via the identification of critical protein interactions and their regulation by post-translational modifications is important to increase our fundamental knowledge of virus–host biology and for the design of virus control strategies.
Potyviral protein functions (reviewed by Rajamäki et al., 2004) have been assessed using several approaches, but there have been few experiments that allow purification and characterization of viral proteins and protein complexes produced in the context of a virus infection. As the natural infection within host cells is the only environment where viral proteins are definitely synthesized and modified to form functional holoenzymes and structures, purification of viral proteins from active infections can be expected to reveal important aspects of virus biology. At present, there are numerous affinity-tag systems available for purifying proteins from complex protein samples with high specificity and diverse applicability (Terpe, 2003). Tandem affinity purification, relying on tagging proteins with the IgG-binding units of protein A from Staphylococcus aureus and the calmodulin-binding peptide, has proved to be highly successful in purifying protein complexes that exist only in minor amounts in cells (Rigaut et al., 1999; Puig et al., 2001). This method was used for virus-derived protein complexes to identify components of borna disease virus ribonucleoprotein complexes (Mayer et al., 2005). The large tandem affinity purification tag used in those studies could easily disrupt virus infectivity if expressed from the infectious viral cDNA copy, whereas small peptide affinity tags are less likely to interfere with the critical functions of the engineered protein. Over 70 % of 1125 random insertions of a 15 bp sequence into the genome of potato virus A (PVA; genus Potyvirus) still allowed virus replication in tobacco protoplasts (Kekarainen et al., 2002), indicating that several sites in the PVA genome may tolerate small affinity-tag insertions. After an insertion of a histidine (His) tag onto the 6K2 protein of PVA, a spontaneous mutation adjacent to the His tag was required in order to restore infectivity (Spetz & Valkonen, 2004). After several host passages, four of the six His had mutated and thus the affinity tag was lost. This highlights that some problems may arise when engineering potyviral proteins, including loss of virus infectivity and genomic stability of the insertion.
Most potyviral proteins are multifunctional, participating in several steps of the viral life cycle. The multiple functions of individual viral proteins need to be regulated and here the host-cell regulatory system participates. Previously, it was shown by our group that phosphorylation of the PVA coat protein (CP) by the host protein kinase casein kinase II inhibits RNA binding of CP and that the observed phosphorylation is important for virus infectivity (Ivanov et al., 2003). The PVA genome-linked viral protein (VPg) is phosphorylated in vitro (Ivanov et al., 2001). The phosphorylation patterns of recombinant VPgs, corresponding to VPgs of PVA isolates B11 and M obtained with Solanum commersonii plant sap as the kinase source, differ in a manner correlating with the capacity of these viruses to cause systemic infection in this plant (Puustinen et al., 2002). VPg, the subject of this study, is important in all critical steps of viral infection, i.e. replication, movement and virulence (reviewed by Rajamäki et al., 2004), and interacts with different host proteins (Wittmann et al., 1997; Dunoyer et al., 2004; Léonard et al., 2004). Differential proteolytic processing of the potyviral polyprotein is probably an important mechanism regulating VPg functions, as individual potyviral proteins are initially synthesized as one polyprotein that is subsequently processed into 10 mature proteins (Riechmann et al., 1992). Evidence exists that some polyprotein intermediates are long-lived and that they serve specific functions (Restrepo-Hartwig & Carrington, 1994; Martín et al., 1995; Schaad et al., 1997; Merits et al., 2002). Our knowledge about the different roles and functions of VPg in the infection process has been collected either by following the infection process with mutated viral genomes or by in vitro analysis. The area lacking thorough investigation is the status and interactions of VPg in the context of virus infection in planta. In this paper, we have approached the complicated biology of potyviral VPg by developing a system to purify VPg from infected plants and by analysing the purified protein products.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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) tagged with green fluorescent protein (GFP) and cloned under the cauliflower mosaic virus 35S promoter (35S-PVA-GFP; Ivanov et al., 2003) using standard procedures. The previously described PVA infectious cDNA fragment (nt 5163–6370) in pBlueScript (Rajamäki & Valkonen, 1999) was engineered by site-directed PCR mutagenesis to introduce the coding sequence for the HisHA tag (HHHHHHYPYDVPDYA). The insertion was generated by PCR using primers 5'-TACCCGTACGACGTCCCGGACTACGCCGAGTCGGTTGAATTCGAGTC-3' and 5'-GTGGTGGTGGTGGTGGTGTTTCTTCTGTGGTAGTTCAGC-3'. The fragment containing the insert was transferred to 35S-PVA-GFP. In spite of the GFP tag, the resulting cDNA was named 35S-PVAHisHA, the virus PVAHisHA and the protein VPgHisHA for simplicity. The initial GFP-tagged construct in this paper is referred to as wt PVA following the same logic.
Virus inoculation and detection.
The conditions for growing Nicotiana tabacum cv. SR1 and N. benthamiana were 22 °C, 18 h day/18 °C, 6 h night. The 35S-PVAHisHA and 35S-PVA cDNAs were inoculated onto N. benthamiana leaves using a PDS-1000/He particle delivery system (Bio-Rad Laboratories). Systemically infected leaves from these plants were harvested 10 days post-infection (p.i.) and stored at –80 °C for subsequent use for mechanical inoculations. For mechanical inoculation, these leaves were ground in PBS (1 : 10, w/v) with a pestle and mortar and rubbed onto the leaves of N. benthaminana or N. tabacum using carborundum as an abrasive. Infected plants were inspected with a hand-held long-wave UV lamp (B-100; UVP) and GFP fluorescence was documented using a Caplio R4 digital camera using 8 s exposure times.
SDS-PAGE and immunoblotting.
SDS-PAGE was performed essentially according to Laemmli (1970). Proteins were detected either by silver staining according to standard protocols or by immunoblotting. For immunoblotting, proteins were electrophoretically transferred to PVDF membranes (Immobilon-P; Millipore). Subsequently, membranes were probed with anti-HA monoclonal antibody (mAb) HA.11 (diluted 1 : 1000; Covance Research Products), affinity-purified anti-VPg polyclonal antibodies as described in Puustinen et al. (2002), anti-CP mAb (1 : 10 000; Adgen), anti-NIa proteinase (NIa-Pro) antibodies (1 : 5000), anti-RNA polymerase (NIb) antibodies (1 : 5000) and anti-cylindrical inclusion protein (CI) antibodies (1 : 20 000), and with anti-helper-component proteinase (HC-Pro) antiserum (1 : 1000). The production of antigens and subsequent production of anti-PVA protein antisera has been described by Merits et al. (1999). Purification of the antibodies followed the method of Peränen (1992). Detection of the horseradish peroxidase (HRP)- or alkaline phosphatase-conjugated secondary antibodies was carried out using ECL Substrate (GE Healthcare), Immobilon Western Chemiluminescent HRP Substrate (Millipore) or Western Blue Stabilized Substrate for Alkaline Phosphatase (Promega). Protein concentrations were determined using a BCA Protein Assay kit (Pierce) according to the manufacturer's instructions.
RT-PCR and sequencing of the genomic affinity tag.
PVAHisHA infection was passaged twice through N. benthamiana plants, using systemically infected leaves (7 days p.i.) from the previous passage for mechanical inoculation. Systemically infected leaves from the third round were harvested from three independent plants and total RNA was extracted using a total plant RNA isolation kit (Qiagen) according to the manufacturer's instructions. Reverse transcription was carried out using 2 µg total RNA per reaction with Moloney murine leukemia virus reverse transcriptase (Promega), and the PCR was carried out with primers amplifying the HisHA tag encoding the VPg region of the PVA genome. The PCR-amplified fragment was subsequently sequenced.
Preparation of a heavy membrane fraction.
Infected leaves (10 days p.i.) of N. benthaminana were homogenized in pre-chilled native buffer [50 mM HEPES (pH 8), 100 mM NaCl, 5 mM MgCl2, 1 µM leupeptin, 1 µM pepstatin A, 1 % polyvinyl pyrrolidone (PVP) and 10 % glycerol] using 2 ml buffer (g tissue) –1 in a mixer at 4 °C. The homogenate was cleared by two sequential centrifugations at 4000 g for 10 min at 4 °C. A heavy membrane pellet was prepared from the cleared supernatant by centrifugation at 30 000 g for 20 min at 4 °C and was designated P30.
Denaturing purification of VPgHisHA.
All procedures were performed at room temperature. Proteins from infected N. benthamiana leaves (10 days p.i.) were extracted for 60 min into lysis buffer [20 mM NaH2PO4, 10 mM Tris base, 8 M urea (pH 8), 2 % PVP and 1 % Triton X-100] using 2 ml buffer (g leaf tissue) –1. P30 membrane pellets were suspended in 0.5 ml lysis buffer (g initial leaf tissue)–1. After extraction, lysates were cleared by centrifugation at 20 000 g for 20 min. The supernatant was mixed with 1 ml Ni-NTA resin (Qiagen) for each 50 g initial leaf material and incubated in a rotator for 2 h. The resin was collected by sedimentation (1000 g for 2 min) and transferred to a 5 ml column, washed twice with 5 ml lysis buffer, three times with 5 ml lysis buffer adjusted to pH 6.3 and finally eluted with lysis buffer adjusted to pH 4.5. Washing and elution buffers were without PVP and Triton X-100.
Two-dimensional (2D)-PAGE.
Protein samples obtained from denaturing purification were diluted 1 : 10 (v/v) in isoelectric focusing rehydration buffer [8 M urea, 0.5 % IPG buffer (NL pH 3–11; GE Healthcare), 20 mM DTT and 1 % CHAPS]. For total protein extracts, leaves were homogenized directly in rehydration buffer and the protein concentration was adjusted. Non-linear pH 3–11 IPG strips (7 cm; GE Healthcare) were used for isoelectric focusing according to the manufacturer's protocol. Before the second-dimension separation, strips were equilibrated for 20 min in SDS buffer [50 mM Tris/HCl (pH 7.4), 1 % SDS and 100 mM β-mercaptoethanol], positioned on 12 % SDS-PAGE gels and fixed by embedding in 0.5 % agarose in SDS-PAGE running buffer. SDS-PAGE and immunodetection with anti-VPg IgGs was carried out as described above.
Enzymic analysis of phosphorylation.
Protein samples from denaturing purification were dialysed against phosphatase buffer [50 mM Tris/HCl (pH 7), 100 mM NaCl, 0.025 % Tween 20 and 0.1 mM EGTA]. Complete protease inhibitor cocktail (EDTA-free; Roche diagnostics), 2 mM DTT, 10 mM MgCl2 and 2 mM MnCl2 were added to the dialysed sample. Precipitated proteins were removed by centrifugation at 16 100 g for 10 min at 4 °C. The sample was divided and incubated with a mixture of the following enzymes: 3 U recombinant lambda protein phosphatase (
-PPase; New England Biolabs) µl–1, 0.05 U protein phosphatase 2A (PPase 2A; New England Biolabs) µl–1 and 0.02 U calf intestine alkaline phosphatase (CIAP; Promega) µl–1 at 30 °C for 60 min. A control sample was incubated under the same conditions but without the phosphatases. After phosphatase treatment, the samples were separated by 2D-PAGE and detected by immunoblotting using anti-VPg IgGs. The ScanSite molecular mass and isolectric point calculator found at the Expasy server (http://scansite.mit.edu/calc_mw_pi.html) was used to calculate the influence of phosphorylation on pI.
Purification of VPgHisHA under native conditions.
All procedures were performed at 4 °C using pre-chilled buffers. For tandem purification, 50 g leaves systemically infected with either PVAHisHA or wt PVA were harvested at 10 days p.i. and homogenized in 100 ml tandem lysis buffer [50 mM NaH2PO4, 150 mM NaCl, 2 % PVP, 0.1 % Triton X-100, 0.1 % Tween 20, 13 % sucrose and one complete protease inhibitor cocktail tablet (EDTA-free; Roche Diagnostics) per 50 ml buffer, pH 8] using a kitchen rod mixer. The homogenate was cleared by centrifugation at 20 000 g for 20 min. The cleared protein extract was incubated with rotation for 2 h with 1 ml Ni-NTA resin (Qiagen). The resin was collected by sedimentation (1000 g for 2 min), transferred to 5 ml columns and washed as follows; twice with 5 ml PBS plus 0.1 % Tween 20 (PBS-Tween), once with 5 ml PBS-Tween plus 20 mM imidazole and twice with 1 ml PBS-Tween plus 50 mM imidazole. Proteins were eluted using four volumes of 1 ml PBS-Tween containing 250 mM imidazole. After elution, the sample was desalted 1 : 10 with regard to imidazole using an Amicon filtering device (Millipore) and precipitated proteins were removed by centrifugation 16 000 g for 10 min. Samples were taken for the analysis of His purification only. The samples were then incubated for 2 h with rotation with 0.5 ml anti-HA affinity matrix (HA.11), collected in 5 ml columns, washed twice with 5 ml PBS, once with 2 ml PBS diluted 1 : 10 in ddH2O and eluted with 0.1 % trifluoroacetic acid in ddH2O (pH 
2.5) using four 1 ml volumes. The elution was pooled, dried in a SpeedVac (Savant) and analysed by SDS-PAGE. For anti-HA purification alone, cleared protein extracts were obtained as for tandem purification except that the lysis buffer comprised PBS, 2 % PVP, 0.1 % Triton X-100, 0.1 % Tween 20, 13 % sucrose and protease inhibitors as in the tandem lysis buffer (pH 7.4). The cleared protein extracts were incubated for 2 h with rotation with 0.5 ml anti-HA affinity matrix. The affinity matrix was collected by sedimentation (1000 g for 2 min), washed once with 20 ml PBS-Tween, twice with 5 ml PBS and once with 2 ml PBS diluted 1 : 10 in ddH2O. Elution was carried out as for the tandem purification.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The engineered virus, PVAHisHA, infected both N. benthamiana and N. tabacum systemically, and the number of plants infected was the same as for wt PVA (95–100 %). The GFP pattern and intensity of the engineered virus were broadly similar to those of wt PVA (Fig. 2), with the following differences. In N. benthamiana, GFP was first observed in the veins of systemic leaves from which it spread to the leaf mesophyll. GFP fluorescence appeared 1 day later in the upper systemic veins for PVAHisHA (7 days p.i.) than for wt PVA (6 days p.i.). The mesophyll entry appeared also to be somewhat slower for the mutant. In N. tabacum, GFP appeared as spots in the systemic leaf mesophyll and by 14 days p.i., the amount and size of spots due to PVAHisHA were both reduced in comparison with those due to wt PVA. When a triple HisHA tag sequence encoding 53 aa and a GFP gene was introduced to the same insertion site, the mutant viruses were not able to establish systemic infection (data not shown).
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, lane 4), representing tagged VPg and NIa, respectively. The tagged proteins migrated at a position of a few kDa more than wt VPg, consistent with the presence of the affinity tag.
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, lanes 1 and 2). Due to the uniform staining of the lane containing the proteins purified from the PVAHisHA-infected plants, it was difficult to identify the specific VPg products (Fig. 4a, lane 2). To show a purification gain, the same quantity of total protein from wt-and PVAHisHA-infected plants before and after Ni-NTA purification was subjected to immunoblot analysis (Fig. 4b). The purified sample derived from PVAHisHA-infected plants resulted in a clear signal (Fig. 4b, lane 4), whereas hardly any signal was detected in the other samples (Fig. 4b, lanes 1–3).
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, lane 4) and anti-HA (lane 5), but not with anti-NIa-Pro IgGs (lane 6), and were concluded to represent two forms of VPg. A major band at 55 kDa was detected by all three antibodies and was concluded to be NIa. There were at least three high-molecular-mass protein bands detected by all antibodies above 120 kDa, probably representing polyprotein forms (Fig. 4b, lanes 4–6).
Analysis of purified VPgHisHA by 2D-PAGE
Separation by 2D-PAGE followed by immunodetection revealed that both VPg and NIa were present in multiple isoelectric isoforms (Fig. 5a). Based on the pH gradient map of the strip used in isoelectric focusing, we estimated that the observed isoforms ranged from pI 5.3 to 6.3. At least five major VPg isoforms were detected. The theoretical pI for both VPgHisHA and NIaHisHA is 8. Interestingly, a small amount of VPg was detected at this pI only when the amount of analysed protein was increased fivefold (Fig. 5b). The increased protein amount also revealed a new row of isoforms residing above the major one, starting from pI 7 and corresponding in size to unmodified VPg at pI 8. One possibility is that the two rows of isoforms corresponded to the 25 and 27 kDa bands observed in the immunoblots in Fig. 4. As replication and probably translation of viral proteins take place in association with membranes, we investigated the pattern of VPg isoforms in a membrane-enriched protein sample and found that the pattern of acidic isoforms was similar to that from the total protein purification, although the molar ratios differed. Two especially abundant isoforms were detected at approximately pI 6, whereas the amount of the more acidic forms was clearly reduced in the membrane fraction (Fig. 5c). Also, the relative concentration of VPg at pI 8 appeared higher for membrane-purified VPg. To rule out the possibility that the observed isoforms were because of the inserted affinity tag, similar 2D-PAGE analysis was carried out for a non-purified total protein sample from wt PVA-infected plants (Fig. 5d). A row of six different isoforms of VPg was present in this sample, verifying that both wt and VPgHisHA exist in multiple isoelectric forms during infection.
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-PPase and CIAP abolished three acidic VPg isoforms (Fig. 6a and b, box 2), indicating that these were very probably phosphorylated VPg forms. Two major isoforms were still present after PPase treatment at pI 6, seemingly corresponding to the abundant isoforms detected from heavy membranes (Fig. 5c). The intensity of the PPase-resistant isoforms increased after PPase treatment (Fig. 6a and b, box 1), indicating that dephosphorylation caused reversion to these isoforms. The pH coverage of PPase-sensitive isoforms in box 2 was estimated to range from 5.4 to 5.7. Conversion of the most acidic isoform (pI 5.4) to the PPase-insensitive isoform of pI 5.9 requires, in theory, the removal of approximately six phosphate groups according to the ScanSite results. Another cluster of 12 kDa isoforms, which was detected specifically with anti-VPg antibodies, ran just above the SDS-PAGE front (Fig. 6a, box 3). This protein product was also affected by PPase treatment. In this case, ScanSite counted four phosphate groups to cover the estimated pH range (5.0–5.3), corresponding to the number of spots observed. NIa was not detectable in the PPase analysis, possibly because of precipitation during dialysis, and therefore no statement concerning its phosphorylation can be made. Nevertheless, the acidic shift seen in Fig. 5 indicates the presence of post-translational modifications.
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). To show a purification gain, the same quantity of total protein from wt- and PVAHisHA-infected plants before and after Ni-NTA purification were subjected to immunoblot analysis (Fig. 7b). This showed that both wt NIa and NIaHisHA were purified to the same extent, whereas the purification of VPg was His-tag dependent. NIa of turnip mosaic virus has been shown to be purified by nickel–agarose resin without a His tag (Ménard et al., 1995; Plante et al., 2004), which appears also to be the case for PVA NIa. After Ni-NTA purification, the amount of total protein was reduced to 1/200 of that of the initial lysate. The second purification step using an anti-HA affinity matrix led to a further increase in purity (Fig. 7c and d). After tandem purification, the amount of total protein (13 µg) was reduced to 1/50 000 of that of the initial lysate (700 mg). Silver staining showed several protein bands that appeared to be specific for the VPgHisHA sample (Fig. 7c). These should represent proteins co-purifying due to interactions with VPg or VPg-containing molecular complexes in addition to VPgHisHA proteins. Purification using anti-HA matrix alone was also evaluated (Fig. 7e and f). Immunoblot analysis demonstrated specific purification (Fig. 7f, lane 4). From the silver-stained gel, some bands appeared to be specific for VPgHisHA, but there were still more proteins present in the wt PVA sample than after tandem purification (compare Fig. 7c and e, lanes 1).
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). PVA NIb, HC-Pro and CI co-purified specifically with VPgHisHA. From the total protein sample, anti-NIb readily detected a band of 50 kDa. This band was 10 kDa smaller than the calculated NIb (59 kDa). NIb was therefore most likely the less intense band corresponding in size to both the co-purified signal and recombinant NIb. The origin of the 50 kDa band in total protein lysate is not clear. CP was not detected with the anti-PVA CP mAbs, whereas the light chains of the anti-HA mAbs derived from the affinity matrix were detected with the secondary antibody used. The absence of CP indicated that PVA virions did not purify in the tandem affinity system, despite the fact that VPg is exposed at the virion end (Puustinen et al., 2002).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Blanc et al., 1999; Arazi et al., 2001). Successful purification under native conditions showed that the affinity tag, and hence the C terminus of VPg, was exposed on the surface of both VPg and NIa.
Phosphorylation of PVA VPg has been studied by our group for some time (Ivanov et al., 2001; Puustinen et al., 2002). The demonstration of phosphatase-sensitive VPg isoforms present in plants during infection supports and complements the previously obtained data. The possibility of using denaturing conditions in purification is of great convenience when reversible post-translational modifications such as protein phosphorylation or ubiquitinylation are studied, as undesired enzymic activities are not present after cell lysis (Tagwerker et al., 2006). After the pure protein has been obtained from infected plants, detailed characterization of post-translational modifications by mass spectrometry is possible (Witze et al., 2007). This is a future direction for VPg characterization.
The fact that the majority of VPg present in plants was found at pI 6 and not at pI 8, which is the theoretical pI of the tagged VPg, is interesting. The acidic shift is partially attributable to phosphorylation, but even after PPase treatment, VPg was found at pI 6, indicating that an additional modification is abundant in the pool of VPg proteins. A covalently linked ribonucleotide or several of them would be an expected modification (Puustinen & Mäkinen, 2004), but a pI shift from 8 to 6 would require an RNA extension that should be observed as an increase in the VPg size in the gel electrophoresis. Two VPg bands at 25 and 27 kDa were recognized by both anti-VPg and anti-HA antibodies from the sample obtained by denaturing purification. The 27 kDa VPg band apparently corresponded to the row of slightly larger isoforms in 2D gels. This row contained VPg at pI 8, which can be assumed to be the non-modified, intact VPgHisHA. The 27 kDa pI 8 form was more abundant in the sample derived from the membrane fraction. The first acidic isoform on this row was found already at pI 7 (see Fig. 5b), a shift that the addition of one phosphoryl group would cause. The slightly smaller form, probably corresponding to the 25 kDa VPg of the one-dimensional gels, was the major form detected in the total protein sample. One option is that a truncated 25 kDa form of VPg exists. A C-terminal cleavage can be excluded, as purification is dependent on the affinity tag being intact. When recombinant VPg is subjected to a partial trypsin digestion, an N-terminal cleavage of 14 aa is among the first taking place, demonstrating that this region is readily accessible to proteases (K. Rantalainen and K. Mäkinen, unpublished results). Removal of these 14 aa changes the theoretical pI of VPg to 6.2. Further investigation is required to prove the existence of a truncated VPg form. In turnip mosaic virus, the N-terminal 16 aa of VPg are needed for the important interaction with potyvirus VPg-interacting protein PVIP (Dunoyer et al., 2004). This interaction is required for systemic infection. It may therefore be assumed that the N terminus of VPg should be intact for this binding function. The phosphorylated polypeptide of 12 kDa, recognized specifically by anti-VPg antibodies, is also an interesting observation. The biological significance of the different VPg forms observed requires further studies.
The finding that PVA VPg is multiply phosphorylated in vivo sheds light upon the possible regulatory mechanisms controlling the diverse functions of VPg during potyvirus infection. Potyviruses replicate in association with cellular endomembranes (Martín & García, 1991; Martín et al., 1995; Schaad et al., 1997). The importance of regulation of positive-sense ssRNA virus replication by protein phosphorylation was reviewed recently (Jakubiec & Jupin, 2007). Phosphorylated isoforms of PVA VPg were not abundant in the membrane fraction, which may indicate that phosphorylation is a modification occurring after active replication. A recent report described VPg of potato virus Y as structurally highly disordered (Grzela et al., 2008). Phosphorylation has been observed to be abundant and important for functional regulation of intrinsically disordered proteins (Iakoucheva et al., 2004). In addition to the possible role of phosphorylation in structural adaptation, a role in targeted host defence and degradation, for example via the ubiquitin pathway, is another possibility.
Host proteins are assembled together with viral proteins to form functional protein complexes, and purification and characterization of these from active infections can be considered one of the important aims of future research. By developing a tandem affinity purification protocol, we were able to show the apparent co-purification of three viral proteins: HC-Pro, NIb and CI. NIb has been demonstrated to interact with VPg in the yeast two-hybrid system and in vitro (Hong et al., 1995; Li et al., 1997; Fellers et al., 1998; Darós et al. 1999; Guo et al., 2001). The VPg–NIb interaction should take place at least in the initiation of replication, when VPg is probably uridylylated by the catalytic activity of NIb (Puustinen & Mäkinen, 2004). The VPg–HC-Pro interaction has also been described in vitro and using the yeast two-hybrid system (Yambao et al., 2003; Roudet-Tavert et al., 2007). Furthermore, HC-Pro and VPg reside in close proximity in the tip structure present at the other end of potyviral particles (Torrance et al., 2006). To our knowledge, no direct interaction between CI and VPg has been shown. NIb, HC-Pro and CI all bind to RNA (Merits et al., 1998). Further experimentation is required to determine in which protein or ribonucleoprotein complex context the interactions detected take place. The interactions could be via VPg, via the viral RNA genome or via a third protein interacting with VPg. Nevertheless, the specific co-purification of these viral proteins is promising when considering the validity of our system and the demonstration that the purification protocol used was gentle enough to retain these interactions.
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ACKNOWLEDGEMENTS |
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Received 7 December 2008;
accepted 18 February 2008.
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