Architecture of the Flaviviral Replication Complex

PROTEASE, NUCLEASE, AND DETERGENTS REVEAL ENCASEMENT WITHIN DOUBLE-LAYERED MEMBRANE COMPARTMENTS*

Pradeep Devappa Uchil {ddagger} and Vijaya Satchidanandam §

From the Department of Microbiology and Cell Biology, Indian Institute of Science, Bangalore 560012, India

Received for publication, February 19, 2003 , and in revised form, March 27, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Flavivirus infection causes extensive proliferation and reorganization of host cell membranes to form specialized structures called convoluted membranes/paracrystalline arrays and vesicle packets (VP), the latter of which is believed to harbor flaviviral replication complexes. Using detergents and trypsin and micrococcal nuclease, we provide for the first time biochemical evidence for a double membrane compartment that encloses the replicative form (RF) RNA of the three pathogenic flaviviruses West Nile, Japanese encephalitis, and dengue viruses. The bounding membrane enclosing the VP was readily solubilized with nonionic detergents, rendering the catalytic amounts of enzymatically active protein component(s) of the replicase machinery partially sensitive to trypsin but allowing limited access for nucleases only to the vRNA and single-stranded tails of the replicative intermediate RNA. The RF co-sedimented at high speed from nonionic detergent extracts of virus-induced heavy membrane fractions along with the released individual inner membrane vesicles whose size of 75–100 nm as well as association with viral NS3 was revealed by immunoelectron microscopy. Viral RF remained nuclease-resistant even after ionic detergents solubilized the more refractory inner VP membrane. All of the viral RNA species became nuclease-sensitive following membrane disruption only upon prior trypsin treatment, suggesting that proteins coat the viral genomic RNA as well as RF within these membranous sites of flaviviral replication. These results collectively demonstrated that the newly formed viral genomic RNA associated with the VP are oriented outwards, while the RF is located inside the nonionic detergent-resistant vesicles.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Although replication of flaviviruses has been an extensively studied aspect, the precise mechanism adopted and intricate interactions among the factors involved are yet to be unraveled. The flavivirus genome is a single-stranded positive-sense RNA ~11-kb long, lacking a 3'-poly(A) tail but with a 5'-type I cap. This genomic RNA upon uncoating utilizes the host translational machinery to direct synthesis of an ~3,400 amino acid long polyprotein that is processed co-translationally and post-translationally by the host signalase and a virus-encoded proteinase to give three structural (capsid, premembrane/membrane, and envelope) and seven nonstructural (NS)1 proteins (NS1-NS5) (1). The replication of the viral genome is thought to take place using putative complexes composed of viral as well as hypothetical host protein(s) (2). This process is initiated by the synthesis of a negative strand RNA complementary to the viral genomic plus strand, resulting in a double-stranded (ds) replicative form (RF). Asymmetric and semi-conservative synthesis of RNA (3, 4) from the RF results in formation of replicative intermediates (RI) with nascent single-stranded RNA tails that resolve upon completion of strand synthesis to generate one molecule of single-stranded RNA and a RF.

Two decades of scientific effort have revealed the putative and/or actual functions of most of the nonstructural proteins in the flavivirus life cycle. NS5, the largest of all of the viral proteins, functions as the RNA-dependent RNA polymerase (RdRp) (57) and a methyl transferase (8), the latter implicating its role in capping of viral genomic RNA. The multifunctional protein NS3 manifests three activities. 1) The viral protease along with the cofactor NS2b is critical for proper processing of the viral polyprotein (911). 2) A helicase is required most probably for unwinding dsRF (12). 3) NTPase activity (13) presumably is required in the first step of capping the viral genomic RNA. The secreted NS1 protein is a soluble complement-binding factor for which a role in negative strand RNA synthesis has also been ascribed (14). NS4a, an integral membrane protein, is believed to serve as a protein bridge between NS1 with which it specifically interacts (14) and the flaviviral replication complex (RC), thus tethering the RC with its numerous proteins to the membrane (15). The small hydrophobic protein NS2a has been shown to specifically bind the 3'-untranslated region and together with NS5 and NS3 that independently bind the same region has been hypothesized to seed the formation of RC (16). Recent evidence has also revealed a surprising role for both NS3 and NS2a in virion morphogenesis (17). The role of NS4b is debatable as it localized more in the nucleus than at the sites of replication (18).

The RNA-synthesizing machinery of virtually all of the eukaryotic cytoplasmic single-stranded positive-sense RNA viruses including members of the togaviridae, flaviviridae, coronaviridae, arteriviridae, bromoviridae, and picornaviridae have been known to intimately engage the host intracellular membranes as platforms for viral replication (19, 20). Cryo-immunoelectron microscopy carried out on cells infected with Kunjin virus (KUNV) and dengue virus (DENV) has revealed an extensive rearrangement of host-derived membranes leading to the development of distinct structures termed as convoluted membranes (CM) that reversibly form alternate structures called paracrystalline arrays (PC) (15, 21). In addition, present at the periphery of and closely associated with CM/PC are clusters of several small vesicles in close apposition with each other called vesicle packets (VP) (21). Although the VP and CM/PC represent distinct cellular compartments, they appear to be interconnected via the bounding of rough endoplasmic reticulum (22). The CM/PC originate from membranes derived from intermediate compartments and are presumed to be the site for proteolytic cleavage of the nascent polyprotein by the viral protease complex NS2b-NS3 located therein (15). The VP on the other hand are derived from membranes of the trans-Golgi network (22), and flaviviral replication is thought to ensue in tight association with the VP since the dsRF. which is presumably the template for viral RNA synthesis, was associated with these structures (15, 23). Furthermore, RdRp activity also predominantly localized to heavy membrane fractions that contained smooth membrane vesicle-like structures (SMS) (24, 25), which may be synonymous with VP as noted earlier (26).

Thus, while there exists extensive literature on ultrastructure of virus-induced membrane structures and the identity of the host organelle whence these membranes originate, there still persists a dearth of information pertaining to the architecture of flaviviral RC housed within these membranes. The protease sensitivity of the major flaviviral replicase proteins NS5 and NS3 had suggested a cytoplasmic orientation for the membrane bound RC (27, 28). In contrast, electron microscopic analysis carried out on KUNV-infected cells as well as DENV-infected cells displayed a dominant association of the RF as well as replicase proteins with membranous vesicle packets that were in turn enclosed by an outer membrane (15, 21). In keeping with these observations, we have earlier shown that extensive protease treatment of heavy membrane fractions from Japanese encephalitis virus (JEV)-infected cells did not compromise the in vitro RNA-dependent RdRp activity, despite effecting near-complete destruction of the major replicase proteins NS3 and NS5 (29). This result highlighted two important features of the flaviviral RC: 1) the presence of a bounding membrane that protects the enzymatically active replicase from protease action and 2) the requirement for only catalytic amounts of replicase proteins. In this study, we extend these observations for the flaviviruses West Nile virus (WNV) and DENV and also provide definitive biochemical proof using protease, detergents, and nuclease as tools for location of the flaviviral RNA species as well as viral replicase proteins behind a membrane barrier that encloses the RC. Our results also suggest that flaviviral RC are associated with differentially detergent-sensitive double layered vesicular structures wherein the newly formed vRNA is extruded into the intermembrane space while the RF remains protected inside the inner vesicular compartment, tightly associated with proteins. The implications of such an organization that confers differential accessibility for the viral RNA species to the host cell environment are discussed.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Viruses and Cells—WNV strain E101, JEV strain P20778 [GenBank] (GenBankTM accession number AF080251 [GenBank] ), and DENV-2 strain TR 1751 (National Institute of Virology, Pune, India) were propagated in the Aedes albopictus cell line C6/36 (National Centre for Cell Science (NCCS), Pune, India) in minimum essential medium (Invitrogen) supplemented with 5% fetal bovine serum, 0.3% tryptose phosphate broth (Invitrogen), 0.22% NaHCO3, and 2 mM HEPES (pH 7.3). Confluent monolayers of C6/36 were infected with virus at a m.o.i. of 0.1 for routine expansion, and medium-containing virus was harvested at 5.5 days postinfection (p.i.), aliquoted, and stored at –80 °C until further use. The porcine kidney cell line PS (NCCS) maintained at 37 °C in minimum essential medium with 10% fetal bovine serum in a humidified atmosphere with 5% CO2 was used to determine viral titers by the TCID50 method (30). These cells infected with WNV, JEV, or DENV at a m.o.i. of 10 were used as a source of viral RC 18–22 h p.i.

Preparation of Flaviviral Replication Complexes and in Vitro RdRp Assay—Flavivirus-infected PS cells were harvested by centrifugation at 800 x g at 18–22 h p.i. and used to obtain heavy membrane fractions sedimenting at 16,000 x g (P16) as a source of RC in in vitro RdRp assays as previously described (4, 29). The in vitro RdRp assay, RNA extraction, and analysis using partially denaturing 7 M urea, 3% PAGE followed by autoradiography were carried out as described earlier (4). Results from lithium chloride fractionation and subsequent RNase A digestion of viral RNA species according to reported procedures (4, 29) showed that the RNA species produced in an in vitro RdRp assay using the P16 fraction from WNV- and DENV-infected cells are similar in their properties to those reported earlier for KUNV (31) and JEV (29). We further confirmed the viral origin of the labeled RNA species generated during the in vitro assays by hybridization to unlabeled strand-specific viral RNA probes followed by RNase protection assays (29, 32). The amount of each viral RNA species generated was estimated by scanning the gels on a Fuji BAS1000 phosphorimaging system and analyzed using the Fuji MacBAS V2.4 software.

Metabolic Labeling of Proteins—Subconfluent monolayers of PS cells were mock or flavivirus-infected at a m.o.i. of 10. At 16 h p.i., the cells were labeled with 50 µCi/ml [35S]methionine-cysteine (1175.0 Ci/mmol, EXPRE35S35S, PerkinElmer Life Sciences) as described previously (29). Protein samples after appropriate treatments were analyzed on SDS-10% PAGE. The gels were processed for fluorography using AMPLIFYTM (Amersham Biosciences) according to manufacturer's instructions, dried, and exposed. Antisera specific to the NS3, NS5, NS1, and envelope proteins were used to confirm the identity of labeled proteins for JEV.

In Vivo Labeling of Viral RNA—Mock- or flavivirus-infected (m.o.i. = 5) PS cells were labeled at 16 h p.i as described earlier (29) with 30 µCi/ml [32P]inorganic phosphate (PerkinElmer Life Sciences) for 1 h in the presence of 3 µg/ml actinomycin D. Homogenates were prepared from harvested cells and treated with micrococcal nuclease wherever required as described above. The extracted labeled viral RNA was resolved on a partially denaturing 7 M urea 3%-PAGE and visualized by autoradiography.

Micrococcal Nuclease and Trypsin Treatments—P16 fractions from WNV, JEV, or DENV-infected cells were treated either before or after carrying out the RdRp assays with 15 units/ml micrococcal nuclease (MNase, MBI Fermentas) and 20 units/ml DNase I (Roche Applied Science) in the presence of 1 mM CaCl2 at 30 °C for 30 min. The treatments were terminated by adding EGTA (pH 8.0) to 5 mM and holding on ice for 30 min. Trypsin (sequencing grade, Promega Corporation) treatment was carried out on ice for 15 min at the concentrations mentioned and terminated using soybean trypsin inhibitor (Invitrogen) and phenylmethylsulfonyl fluoride (Sigma) at final concentrations of 2 mg/ml and 1 mM respectively. The samples were incubated on ice for 30 min for complete inactivation of trypsin before further processing.

Detergent and Sodium Citrate Treatment of Virus-infected P16 Fractions—Detergent treatment of virus-infected P16 fractions was carried out at the appropriate concentrations on ice for 1 h. The nonionic detergent Triton X-100 (TX100) was used at a final concentration of 1% that has been reported to solubilize endoplasmic reticulum (ER) and ER-like membranes (33), while the ionic detergent sodium deoxycholate (DOC) was used at a final concentration of 1.5%. Gentle disruption of ER and ER-like membranes was achieved using 1% sodium citrate at 4 °C for 30 min (34). The protein concentration in the homogenates and P16 fractions during all of the treatments was maintained at 2 mg/ml. All of the detergents used were nuclease-free molecular biology grade obtained from Sigma.

Floatation Analysis—The P16 fraction after in vitro RdRp assay from mock- and flavivirus-infected cells was subjected to 1% TX100 or 1.5% DOC at 4 °C for 1 h followed by sedimentation at 16,000 x g for 15 min to obtain S16 fractions, which were used for floatation analysis. 0.5 ml (2 x 106 cells) of the S16 fraction mixed with 4 ml of 75% (wt/wt) sucrose was layered on 0.5 ml of 80% (wt/wt) sucrose and overlaid with 4 ml of 55% (wt/wt) sucrose and 1 ml of 5% (wt/wt) sucrose in buffer containing 10 mM Tris, pH 8.0, 10 mM sodium acetate, and 1.5 mM MgCl2. Gradients were then centrifuged for 18 h at 35,000 rpm in a Beckman L8–80 model Ultracentrifuge using a SW41 Ti rotor at 4 °C, and 1-ml fractions were collected from the top and RNA was extracted and analyzed as mentioned above.

Electron Microscopy of TX100-resistant Membrane Structures—Detergent-treated S16 fraction obtained as above was subjected to ultracentrifugation at 35,000 rpm (150,000 x g) for 5 h. The pellet (P150) obtained (detergent-resistant membrane fraction) was resuspended in ice-cold phosphate-buffered saline and deposited on Formvar-coated copper grids (Ted Pella Inc.) for 3 min and stained with 2% uranyl acetate in distilled water. The samples were visualized in a JEOL JEM-100CXII electron microscope operated at 80 kV.

Immunoelectron microscopy of TX100-treated P150 fractions and first two fractions after floatation analysis, obtained as mentioned above from mock- and JEV-infected cells, were processed for low temperature embedding in LR Gold (Ted Pella Inc.) according to manufacturer's instructions after fixing the samples with 3.7% paraformaldehyde (TAAB Laboratory Equipment) and 0.01% glutaraldehyde (EM grade, Sigma) in phosphate-buffered saline. Ultra thin sections were then incubated at room temperature as follows: 2 h in PBG (phosphate-buffered saline containing 0.1% (w/v) bovine serum albumin, 0.5% (w/v) gelatin (from cold water fish skin, Sigma) and 0.05% (v/v) in Tween 20 (Sigma)); 3 h in polyclonal rabbit anti-JEV NS3 serum diluted 1: 4000 in PBG; 5 x 10 min in PBG; 2 h in anti-rabbit IgG (H + L) antibodies coupled to either 15- or 10 nm-gold (Ted Pella Inc.) diluted 1:100 in PBG; and 5 x 10 min in PBG. The conditions mentioned above were empirically standardized using sections obtained from mock-infected and infected whole cells embedded similarly. Only specific binding of antibodies (both primary and secondary) was observed under these conditions. The sections were then stained with uranyl acetate and lead citrate and examined as mentioned above.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Flavivirus Replicase Proteins Are Required in Catalytic Amounts and Are Present Behind a Membrane Barrier—Extensive trypsin treatment of heavy membrane fractions from WNV and DENV-infected PS cells did not affect the in vitro RdRp activity (Fig. 1A, lanes 2 and 3 for WNV and lanes 5 and 6 for DENV), despite the near complete destruction of the metabolically labeled major replicase proteins NS5 and NS3 (Fig. 1B, lanes 1 and 2 for WNV and lanes 5 and 6 for DENV). We interpreted this to suggest that trace amounts of NS5 and NS3 that are protected from trypsin digestion, probably by a membrane barrier(s), suffice to manifest the total detectable RdRp activity in these two flaviviruses that was similar to the properties of JEV RC previously demonstrated by us (29). The suggested existence of a membrane barrier was tested using the nonionic and ionic detergents TX100 and DOC, respectively. Trypsin digestion of TX100-treated membrane fractions from WNV and DENV decreased the RdRp activity by ~50% (Fig. 1D, lanes 1–3 for WNV and lanes 4–6 for DENV) over and above the 30% reduction in activity observed due to the detergent treatment alone (Fig. 1C, lanes 1 and 2 for WNV and lanes 3 and 4 for DENV), indicating that TX100 caused vital protein components of RC to become partially exposed to trypsin. Specifically, decreased incorporation of label into vRNA and RI species was observed under these conditions (Fig. 1C, lanes 2 and 4), similar to that reported for KUNV (4). The total loss of vRNA in KUNV by this treatment could however be attributed to residual nuclease activity in cytoplasmic extracts used by these workers in contrast to the extensively washed heavy membrane fractions (P16) used by us, which reduces the burden of endogenous nuclease activity. Similar evaluation of the effect of trypsin on JEV RdRp after TX100 treatment could not be carried out because of the complete loss of activity suffered by JEV RC following detergent treatment alone (Fig. 1C, lanes 5 and 6 (29). This could be attributed either to the greater inherent inhibition of JEV RC compared with WNV and DENV by TX100 or selective loss of one or more factors from JEV RC, possibilities that are under investigation. Whereas DOC treatment did not adversely affect RdRp activity (Fig. 1D, lanes 7, 10, and 13), it however led to a complete loss of activity when followed by trypsin in all of the three flaviviruses under study. This suggested complete solubilization of the membrane barrier(s) by the ionic detergent, thereby rendering the functional replicase proteins NS5 and NS3 accessible to trypsin. Since most of the detectable major replicase proteins were degraded even in intact membranes (Fig. 1B, lanes 2 and 6), the exact orientation of the enzymatically active replicase proteins within the associated membranes was difficult to ascertain.



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FIG. 1.
Flaviviral replication complexes are present behind a membrane barrier. Heavy membrane (P16) fractions from WNV (lanes 1–3) and DENV (lanes 4–6) infected cells were subjected to increasing concentrations of trypsin (0–1 mg/ml) as depicted in the flow chart before carrying out the RNA-dependent RdRp assays using [{alpha}-32P]GTP. The labeled RNA products generated were resolved on a partially denaturing 7 M urea, 3% PAGE. B, effect of in vitro trypsin treatment on metabolically labeled flaviviral proteins. P16 fractions metabolically labeled with [35S]methionine-cysteine from actinomycin D-treated WNV (lanes 1–4) and DENV (lanes 5–8) infected cells were subjected to trypsin (1 mg/ml) without (lanes 2 and 6) or with prior treatment with 1% sodium deoxycholate (DOC, lanes 4 and 8) or 1% TX100 (lanes 3 and 7). Lane 9 represents labeled proteins from similarly treated mock-infected cells. The processed samples were electrophoresed on SDS-10% polyacrylamide gel followed by autoradiography. The dots indicate the locations of flavivirus-specific proteins with their putative identities mentioned on the left. The positions of the standard molecular size markers are mentioned on the right. C, effect of TX100 on in vitro flaviviral RdRp activity. P16 fractions from WNV (lanes 1 and 2), DENV (lanes 3 and 4), or JEV (lanes 5 and 6) were treated (T, lanes 2, 4, and 6) or not treated (N, lanes 1, 3, and 5) with 1% TX100 for 1 h on ice followed by in vitro RdRp assay using [{alpha}-32P]GTP. The labeled RNA products after extraction were analyzed using 7 M urea, 3% PAGE. D, P16 fractions from flavivirus-infected cells were processed as depicted in the flow chart, and RdRp assays were carried out after trypsin inactivation. The labeled RNA products after extraction were resolved as in A. Values below lane numbers denote total radioactivity incorporated by all of the three viral RNA species as a proportion of that detected in appropriate control assays shown in lanes represented as 1. The arrowheads in A, C, and D denote the position as well as the identity of the three viral RNA species RI, vRNA, and RF.

 

Flaviviral Replication Complexes Are Present in Micrococcal-Nuclease-resistant Compartments—In the next step of our analyses, we utilized flavivirus-induced membrane preparations from infected PS cells to ascertain the orientation of the three viral RNA species, namely RI, RF, and vRNA, in the membrane-bound RC based on sensitivity to a nonspecific nuclease such as MNase. MNase was the enzyme of choice over others such as RNase A since it is robust and, under the reaction conditions used, digests both single- and double-stranded nucleic acids. The strict dependence of its activity on the divalent cation calcium and consequent complete inactivation using EGTA made it possible to carry out RdRp assays after MNase treatment. The viral RNA species associated with membrane-bound RC can be oriented either toward the luminal space or the cytoplasmic compartment depending on their organization within the membrane. The susceptibility of some, all, or none of the three viral RNA species to MNase would thus help to decipher their organization within the membrane-bound RC.

Exhaustive pretreatment of the membrane preparations from WNV-infected cells with MNase (Fig. 2A, compare lanes 5 and 6, and B, lanes 1 and 2) did not result in any reduction in RdRp activity, suggesting that those species of viral RNA that functioned as template(s) for RNA synthesis were not accessible to nucleases. In addition, MNase treatment at the end of the assay period revealed nuclease resistance of all of the three newly synthesized labeled viral RNA species that were generated during the in vitro reaction (Fig. 2B, compare lanes 1 and 3). The MNase resistance of viral RNAs was not due to their secondary structure and/or double stranded nature, because labeled viral RNA species added exogenously to infected cell P16 fractions were completely digested by MNase (Fig. 2A, lane 8). EtBr staining also confirmed the selective MNase resistance of all of the endogenous viral RNAs but not the host RNA within the infected cell (Fig. 2A, lanes 1–3). Similar results were obtained for JEV and DENV RNA (Fig. 2B, lanes 4–6 and lanes 7–9, respectively). We also carried out in vivo labeling of viral RNAs using radiolabeled [32P]inorganic phosphate to assess the nuclease sensitivity profile of in vivo generated viral RNAs. Again, no reduction in the signal intensities due to the radiolabel in WNV, JEV, and DENV RNA species was evident following MNase treatment (Fig. 2C, lanes 1–6). In contrast, mock-infected cells processed similarly did not show presence of any MNase-resistant RNA species migrating in the gel (Fig. 2C, compare lanes 7 and 8). The residual label in these wells revealed the presence of nonspecific insoluble aggregates following these manipulations. Having thus confirmed that the properties of the in vitro and in vivo-labeled viral RNAs were similar, we confined the subsequent series of investigations to labeled RNA generated from in vitro RdRp assays.



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FIG. 2.
Resistance of flaviviral RNA species to micrococcal nuclease. A, heavy membrane fractions (P16) obtained from WNV-infected cells (I) and mock-infected cells (M) were either treated (+) or not treated (–) with MNase prior to carrying out in vitro RNA-dependent RdRp assay using [{alpha}-32P]GTP. The labeled RNA products were resolved using 7 M urea, 3% PAGE. Lanes 1–3 are ethidium bromide-stained gel photograph of the same gel whose autoradiogram is shown in lanes 4–6. Lanes 7 and 8 show the MNase susceptibility of labeled RNA products exogenously added to heavy membrane fractions of WNV-infected cells. B, MNase resistance of in vitro generated WNV, JEV, and DENV RNA products. WNV (lanes 2 and 3), JEV (lanes 5 and 6), and DENV (lanes 8 and 9) infected cell heavy membrane fractions were subjected to MNase digestion either after or before in vitro RdRp assays in the order shown by the numbers in the top panel. RNA products generated similarly from MNase-untreated controls for each virus are shown in lanes 1, 4, and 7. The labeled RNA products were analyzed as in A. C, MNase resistance of in vivo labeled viral RNA products from flavivirus-infected cells. Actinomycin D-treated WNV (lanes 1 and 2), JEV (lanes 3 and 4), DENV (lanes 5 and 6), and mock-infected cells (lanes 7 and 8) at 16 h p.i. were labeled with [32P]inorganic phosphate for 3 h. The homogenates obtained were either treated (+, lanes 2, 4, 6, and 8) or not treated (–, lanes 1, 3, 5, and 7) with MNase prior to RNA extraction. D, viral RNA products are resistant to MNase even after prior treatment with trypsin. The P16 fractions from WNV-infected cells were processed as depicted in the flow chart with a trypsin treatment included prior to MNase after labeling the viral RNA products with [{alpha}-32P]GTP. The labeled viral RNA products were analyzed as in A. The arrowheads in A–C denote the positions of RI, vRNA, and RF. Exposure times were 8 h for WNV and JEV and 24 h for DENV.

 

The observed nuclease resistance of flaviviral RNAs could be due either to a membrane barrier and/or proteins denying access to the RNA. We explored the potential role of proteins bound to viral RNA species in protecting them from degradation by performing trypsin digestion after the in vitro assay to degrade any bound proteins prior to MNase treatment. All of the three labeled WNV RNA species remained MNase-resistant, despite trypsin treatment (Fig. 2D, lanes 1–6). Activity of trypsin under these conditions was confirmed using actinomycin D-treated and [35S]methionine-labeled proteins from virus-infected cells as shown in Fig. 1B, lane 2. Similar nuclease resistance of all of the three JEV and DENV RNA species was also observed (data not shown). Thus, our results suggested that all of the three viral RNA species most probably reside in a membrane-enclosed and nuclease-resistant compartment that cannot be traversed or disrupted by trypsin in keeping with the trypsin resistance of the RdRp enzyme activity of the replication machinery.

Nonionic Detergent Treatment Exposes Nascent vRNA to Nuclease Degradation—In a manner similar to that used for protein analysis of RC, we used detergents to study the nature of the membranous barrier if any that might confer nuclease resistance on the viral RNAs within the RC. We carried out RdRp assay using P16 fractions obtained from WNV-infected cells followed by nonionic detergent treatment with TX100. The subsequent MNase digestion rendered the single-stranded vRNA and the single-stranded nascent tails of RI sensitive to nuclease action (Fig. 3A, compare lanes 1 and 2). The latter led to a loss of RI species from the origin, where it normally migrates, with its concomitant conversion to RF and consequent increase in the amount of RF in samples treated sequentially with nonionic detergent and MNase (Fig. 3A, lane 2) compared with samples treated with detergent alone (Fig. 3A, lane 1). The residual label in the wells following MNase treatment represents insoluble and nonspecific aggregates since labeled RI RNA free of membranes and proteins is fully susceptible to MNase as seen in Fig. 2A, lane 8. Furthermore, MNase treatment of exogenously added labeled viral RNAs to TX100-treated P16 membranes confirmed the complete susceptibility of RI to MNase action as well as the activity of the nuclease under these conditions (Fig. 3B, lanes 2 and 3). This differential MNase sensitivity pattern of the three different viral RNA species held up even after trypsin digestion of the nonionic detergent treated P16 fraction (Fig. 3A, lanes 3–6). The solubilizing activity of TX100 under the assay conditions was also confirmed by its ability to efficiently extract NS1 (Fig. 3C, compare lanes 2 and 3) and consequently renders it sensitive to trypsin (Fig. 1B, lanes 3 and 7). These results corroborated the data obtained for partial trypsin sensitivity of RdRp activity from TX100-treated P16 fractions and also suggested the presence of an additional membrane barrier resistant to nonionic detergents as well as impervious to trypsin and MNase that protected RF from degradation. The RF is probably present enclosed within the inner membrane while the free vRNA and the single-stranded nascent tails of RI extrude into the intermembrane space.



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FIG. 3.
Susceptibility of the viral RNA species to MNase after detergent, trypsin, and sodium citrate treatment. A, the P16 fractions from WNV-infected cells were processed as depicted in the flow chart after labeling the viral RNA products with [{alpha}-32P]GTP. The values on top of the panel and below lane numbers in A denote arbitrary pixel units obtained when RI and RF band areas, respectively, were quantitated using PhosphorImager. B, in vitro labeled and extracted RNA products were incubated with (lane 2) or without (lane 1) MNase in the presence of detergent alone (lanes 1 and 2) and in combination with inactivated trypsin (lane 3) as in A. C, [35S]Methionine-labeled proteins obtained from P16 fraction of WNV-infected cells were treated with TX100 (T) under RdRp assay conditions and fractionated at the end of the treatment period at 16,000 x g for 15 min to obtain pellet (P) and supernatant (S) fractions as a control for activity of TX100. The dots on the right represent flavivirus-specific proteins absent in mock-infected cells with their putative identities mentioned alongside. The positions of standard molecular size markers are shown on the left. D, RdRp assays were carried out using P16 fractions of WNV-infected cells, treated with 1% sodium citrate, and processed as described in the flow chart. The RNA samples after extraction were analyzed using 7 M urea, 3% PAGE. The arrowheads in A, B, and D denote the positions of RI, vRNA, and RF.

 

The nonionic detergents used in addition to solubilizing membranes may have also perturbed or destroyed RNA-protein interactions, which in turn may have resulted in the susceptibility of vRNA to MNase digestion upon nonionic detergent treatment even in the absence of trypsin digestion (Fig. 3A, lane 2), thus masking any protein-vRNA interaction that might have existed. The more gentle agent sodium citrate, which is also known to disrupt ER and ER-like membranes (34), did not render the viral RNAs sensitive to MNase (Fig. 3D, compare lanes 1 and 2). However, digestion of sodium citrate-treated P16 fractions after RdRp assay with increasing concentrations of trypsin followed by MNase treatment rendered the vRNA increasingly susceptible to degradation by MNase (Fig. 3D, lanes 3–6). These results collectively demonstrated that proteins bound to vRNA protected it from degradation and also revealed that nonionic detergents could remove these weakly bound proteins.

Complete Solubilization of the P16 Fractions with Ionic Detergents Does Not Expose the RF to Nuclease—The data presented thus far suggested that the flaviviral RC reside within membrane compartments with at least two membrane layers, the outer of which has a different detergent solubilization profile from that of the inner layer. DOC, an ionic detergent, was again employed to further probe the architecture of the RC. DOC treatment released most of the RdRp activity into the supernatant fractions (Fig. 4A, compare lanes 1 and 4). However, as shown in Fig. 4A, lanes 2 and 3, the template RF was still resistant to MNase following DOC treatment. On the other hand, pretreatment of DOC-solubilized P16 fractions with trypsin rendered RF susceptible to MNase beginning at 0.5 mg/ml trypsin with a complete loss of full-length intact RF achieved at the highest concentration of trypsin used (Fig. 4B, lanes 1–6). This was in contrast to the inability of trypsin to facilitate access to the RF for MNase following nonionic detergent treatment of the P16 fractions. However, susceptibility of vRNA and the single-stranded tails of RI to MNase without pre-exposure to trypsin were observed following treatment with both types of detergents (Fig. 4A, lanes 3 and 6). These results showed that RF in addition to being present within the inner membrane of the double membranous structure was also shielded completely by proteins whose tight association with RF was resistant to disruption by detergents. The identity and properties of the proteins that bind viral RNA species are currently being investigated.



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FIG. 4.
A, P16 fractions treated with 1.5% DOC were sedimented at 16,000 x g, and the pellet and supernatant fractions obtained were used for RdRp assays either before (lanes 3 and 6) or after (lanes 2 and 5) MNase treatment. Lanes 1 and 4 represent controls not treated with MNase. The extracted RNA samples were analyzed using 7 M urea, 3% PAGE. B, DOC-treated P16 fractions were subjected to increasing concentrations of trypsin following [{alpha}-32P]GTP incorporation in an in vitro RdRp assay. Samples were either not treated (–, lanes 1, 3, and 5) or treated (+, lanes 2, 4, and 6) with MNase. The extracted RNA samples were analyzed using 7 M urea, 3% PAGE. The upper panel shows the ethidium bromide-stained samples while the lower panel shows the autoradiogram of the same gel. C, 1% TX100-treated P16 fraction labeled with [{alpha}-32P]GTP (T, lane 1) were sedimented at 16,000 x g to obtain pellet (P, lane 2) and supernatant (S, lane 3) fractions. 1.5% DOC (lane 4) or 1% TX100-solubilized (lane 7) P16 fractions were subjected to ultracentrifugation at 150,000 x g in a SW41 rotor using a Beckman L8–80 centrifuge for 5 h. Equivalent amounts of supernatants (S, lanes 5 and 8) and pellet (P, lanes 6 and 9) fractions were processed for RNA that were analyzed using 7 M urea, 3% PAGE. D, the metabolically labeled proteins released into the supernatant fractions from TX100-treated heavy membranes from WNV, DENV, and JEV were subjected to ultracentrifugation at 150,000 x g to obtain proteins associated with detergent-resistant vesicles (UP, pellet) and those that were completely solubilized (US, supernatant) by the detergent. Equivalent amounts of pellet and supernatant fractions were analyzed, and the proteins were visualized by autoradiography. The dots on the right represent the major replicase proteins NS3 and NS5. The positions of standard molecular size markers are shown on the left.

 

Thus, our results obtained by analyses of both the replicase proteins and the viral RNAs are in agreement with the presence of RC within vesicle packets as shown for KUNV (15, 23) and additionally suggest that the CM/PC and VP with its bounding ER form a closed compartment. These membranes are sufficiently heavy to sediment at 16,000 x g (24, 25). The differential solubility of the outer ER-like membranes alone to nonionic detergent should as a result release all of the inner vesicles, which being smaller would no longer be expected to sediment at 16,000 x g. Indeed, treatment of P16 fractions with 1% TX100 followed by sedimentation at 16,000 x g showed that ~60–80% of labeled viral RNA species remained in the supernatant fraction (Fig. 4C, lanes 2 and 3). The demonstrated resistance of the RF in these membranes to trypsin and MNase (Fig. 3A) pointed to its presence inside these intact nonionic detergent-resistant membrane structures. Successful co-sedimentation of the major replicase proteins NS3 and NS5 in all of the three flaviviruses studied with RF at 150,000 x g (Fig. 4C, lanes 7–9) from S16 fractions of TX100 extracts (Fig. 4D, lanes 1–6) further indicated that these membrane structures were intact and were associated with the RC. In contrast, DOC-solubilized RNA from P16 fractions of WNV-infected cells did not sediment at 150,000 x g, proving that this detergent completely solubilized membranes housing the RC (Fig. 4C, lanes 4–6). In contrast, DOC-solubilized RNA from P16 fractions of WNV-infected cells did not sediment at 150,000 x g, proving that this detergent completely solubilized membranes housing the RC (Fig. 4C, lanes 4–6). The selective loss of vRNA during these prolonged manipulations following detergent treatment is, in keeping with its heightened sensitivity to degradation, shown earlier (Fig. 3A). Our results are in contrast to that for KUNV RC, which was fully solubilized by nonionic detergents (35). However, we were unable to verify this difference in our laboratory under similar conditions because KUNV is a human pathogen that is not endemic to the Indian subcontinent.

Floatation Analysis and Electron Microscopy of Membrane Structures from Detergent Extracts of P16 Fractions—We next attempted to characterize the detergent-resistant membrane structures by subjecting them to both membrane floatation as well as electron microscopic analysis. P16 fractions and their detergent extracts from WNV-infected cells obtained after RdRp assay were studied by floatation gradient analyses in which intact or detergent-resistant membranes with the associated radiolabeled RF would float to a lower density (i.e. top fractions) based on their buoyancies in a sucrose gradient, whereas free RF not bound to membrane or following dissolution of membranes with DOC would remain at the bottom of the gradient containing the denser sucrose solution. P16 membranes prior to detergent treatment floated as expected to the top fractions (Fig. 5A, bottom panel), whereas DOC extracts of membranes remained at the bottom of the gradient, denoting complete solubilization of membranes as monitored by the presence of radiolabeled RF in these fractions (Fig. 5A, middle panel). However, >70% of radiolabeled RF from TX100 extracts was found in the top two fractions clearly denoting association with detergent-resistant intact membrane structures. The trailing of radiolabeled RF in the lower fractions could be the result of damage suffered during the extensive manipulations by a small proportion of these membrane structures, thereby influencing their buoyancy. Once again, vRNA was absent in detergent-treated samples because of its increased sensitivity. Similar results were obtained for TX100 extracts of P16 membranes from JEV-infected cells (data not shown).



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FIG. 5.
Characterization of TX100-resistant membranous structures. A, P16 fraction (lower panel) and its TX100 (top panel, S16TX100) and DOC (middle panel, S16DOC) extracts after RdRp assay with [{alpha}-32P]GTP were subjected to floatation analysis using sucrose step gradients. Labeled RNA products obtained from 1-ml fractions collected from top of the gradients were analyzed using 7 M urea, 3% PAGE. Fraction numbers are indicated above the top panel. B–D, electron micrographs of the pellet obtained following ultracentrifugation of TX100-treated P16 fractions from WNV-infected cells (B) and mock-infected cells (C). D, electron micrograph of pellet obtained following ultracentrifugation of DOC treated-P16 fractions from WNV-infected cells. Negative staining of these samples deposited on formvar-coated copper grids with uranyl acetate clearly showed the presence of intact membrane vesicles only in TX100-treated fractions. E–H, immunoelectron microscopy of TX100-resistant vesicles obtained as above from JEV (E and F) and mock-infected (H) cells using rabbit anti-JEV NS3 antibodies and visualized using anti-rabbit antibodies conjugated to 15-nm (G) or 10-nm (H) gold particles. The TX100-resistant vesicle obtained from top two fractions after floatation analysis from JEV-infected cells processed as in E is shown in G. The bars represent 100 nm.

 

Electron microscopic analysis revealed vesicular structures measuring 75–100 nm (Fig. 5, B) in ultrasedimented fractions of TX100 extracts from WNV-infected cells, a size similar to that previously reported for structures enclosed within bounding rough endoplasmic reticulum in KUNV- and DENV-infected cells (15, 36). These vesicles were devoid of the outer bounding membrane that held them together in clusters, supporting our biochemical data that suggested its solubilization by nonionic detergents (Fig. 3A). Fractions obtained from mock-infected cells following the same treatment did not contain any TX100-resistant structures (Fig. 5, C and H). In addition, we also did not observe any vesicular structures when DOC-treated membrane fractions were sedimented at 150,000 x g (Fig. 5D). Similar structures were also observed in P150 fractions as well as top fractions of sucrose floatation gradients of TX100 extracts from JEV-infected cells (data not shown). Additionally, we confirmed the virus-induced nature of these JEV-derived structures by resin-embedding the P150 fractions as well as those obtained from the top two fractions of sucrose floatation gradients and immunostaining with rabbit antibodies to JEV NS3, a major replicase protein (Fig. 5, E–G).

Vesicles that harbor viral RNA have been in fact observed previously in closely related togaviruses, mouse hepatitis, and poliovirus (3739). The mechanisms by which these vesicles interact with their host environment for obtaining precursors for and releasing products of RNA synthesis remain to be elucidated. Our results with the three viruses we investigated leads to a model (Fig. 6, inset), wherein the flaviviral RC that associate with the VP form an enclosed double membrane structure impermeable to MNase and trypsin. This model is in excellent accordance with the congregation of vesicles bounded by an additional membrane observed by cryo-immunoelectron microscopy inside KUNV- and DENV-infected cells (15, 21).



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FIG. 6.
Proposed model for flaviviral RNA architecture within RC showing the template RF enclosed within two layers of virus-induced membranes. The inset represents multiple VP-bearing RF being utilized as a template by the viral RC with the synthesized vRNA extruding outward. The RF and vRNA are shown bound to as yet unidentified proteins. Replication occurs within VP (inset), and the outwardly oriented vRNA is released by the RC. The as yet unexplained exit of vRNA into the cytosol of the infected cell to gain access to the ribosomes for translation as well as for packaging and subsequent morphogenesis is also shown.

 


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The replication and transcription of eukaryotic plus-strand RNA viruses is mediated by virus-encoded replicases through a distinctive process of RNA-dependent RNA synthesis. The intimate association of the viral RNA-synthesizing machinery with the host intracellular membranes is a common but poorly understood phenomenon. Membranes have been suggested to play a structural and/or organizational role in the RC, possibly by offering a suitable microenvironment for viral RNA synthesis and/or by facilitating the availability of membrane-bound host enzymes (40). Such an arrangement could also concentrate and compartmentalize viral products by targeting them to a common structure, provide key lipid constituents, and physically support the viral RC (41). The choice of host membranes nevertheless appears to be quite variable for each virus group with BMV (42) and tobacco etch potyvirus utilizing ER-derived structures (43), alphaviruses using the cytosolic surface of endocytic organelles (39), and rubella virus exploiting host lysosomal membranes (44) as the site of assembly for their RC. Extensive modification of host cell membranes and induction of specific vesicular membrane structures bearing viral RC are also common (38). For instance, poliovirus induces formation of a complex of vesicles or "rosettes" from the anterograde membrane-trafficking pathway on the surface of which polio viral RC functions (41, 45). Recruitment of the viral RC to these membrane vesicles appears to be mediated by the intrinsic property of one or more membrane-targeted viral nonstructural proteins, which have been shown in certain instances to induce the membrane alterations even in the absence of viral RNA synthesis (4651). In case of the flavivirus KUNV, induction of intricate membranous structures were proposed to require high levels of both viral RNA and protein synthesis (26). However, studies to address the architecture of the flaviviral RC within these membranes have not been undertaken to date. Thus, this study represents, to the best of our knowledge, the first that explores the organization and orientation of viral RNAs and, to a limited extent, also the proteins constituting the flaviviral RC using a combination of probes.

We were unable to decipher the orientation of the individual replicase proteins responsible for RdRp activity because trypsin treatment, even in the absence of detergents, destroyed most of the major replicase proteins NS5 and NS3 and other small nonstructural proteins known to be involved in replication without concomitant loss of replicase activity. The catalytic amounts of NS5 and NS3 required for the measurable RdRp activity was too low to be detected even by metabolic labeling with [35S]methionine. However, the partial loss of RdRp activity upon trypsin treatment of TX100-treated P16 fractions from WNV and DENV-infected cells revealed the presence of one or more proteins on the surface of TX100-resistant vesicles that were required for complete replicase activity. In contrast, the RdRp activity of alfalfa mosaic virus could be totally destroyed by treatment of intact chloroplasts with trypsin, showing that an "essential part of the enzyme complex faces the in vitro medium and probably the cytosol in vivo" (52).

While the association of the RF with VP has been suggested for both DENV- and KUNV-infected cells using anti-dsRNA antibody in cryo-immunoelectron microscopy (15, 21), the low resolution of the technique did not permit deciphering the exact orientation of the RF. Association of vRNA with the SMS has also been determined using electron microscopic in situ hybridization of DENV-infected cells (53). Although the precise location of vRNA was again difficult to assign, they were often found to be present on the surface of the SMS. Results from our biochemical studies not only extend these observations but also offer conclusive proof for the RF being present within the VP/SMS while the single-stranded vRNA is extruded out as depicted in our model (Fig. 6, inset). The exact mechanism adopted for the extrusion process is yet to be delineated.

The differential susceptibility to solubilization by detergents of the outer and inner membranes of the structures harboring the RC revealed by our study would suggest that they are derived from different host cell organelles. On the other hand, alterations in membrane properties can also be brought about by incorporation of viral and/or associated host proteins into these membranes (54). Furthermore, detergent resistance can be conferred by a high proportion of lipids like cholesterol or glycosphingolipid in these membranes (55) as also by specific posttranslational modification of proteins such as acylation and glycosylphosphatidylinositol (GPI) anchoring that are known to render the membranes resistant to nonionic detergents (55, 56). The biogenesis of GPI-anchored proteins that give rise to "liquid-ordered domains" is believed to initiate in the Golgi apparatus (56). Interestingly, the membranes of VP that contain flaviviral RC were shown to be derived from the Golgi (22). In keeping with these inferences, a recent report (57) showed the association of caveolin-2, a lipid-raft-associated intracellular membrane protein with the nonionic detergent-resistant membranes housing the RC from the closely related hepatitis C virus.

The presence of double-layered membrane vesicles that harbor the replication machinery is a common feature shared by poliovirus, coronavirus, and flaviviruses. However, critical differences also exist between these viruses in the architecture of the RNA in the RC. The plus strand polio viral RNA as well as the 3D polymerase have been shown to be "superficially associated" with the RC (45). The "core" in poliovirus, which is equivalent to RF, was accessible to nuclease after DOC treatment, whereas the single-stranded viral RNAs as well as the nascent plus strands were nuclease-sensitive even in the absence of prior detergent treatment (45). In the porcine transmissible gastroenteritis coronavirus also, the bulk of plus strand RNA was accessible to nuclease in the absence of detergents (58), leading to the conclusion that viral RNA was "surface-adherent." In contrast, the nascent viral RNA in flaviviruses was present between the two membrane layers as a result of which similar susceptibility was manifested only following nonionic detergent treatment. The different patterns of detergent-induced nuclease susceptibility of the RNA of different positive strand RNA viruses could also be due to the use of different host organellar membranes to house replication complexes referred to earlier. Whereas the BMV RNA3 was resistant to nucleases in the absence of detergents as we observed for flaviviral RNAs in intact P16 membranes (Fig. 2A, lanes 2 and 3), nonionic detergents rendered it completely susceptible to nucleases as expected for ER-derived spherules that harbor the BMV RC (59). In the transmissible gastroenteritis coronavirus, on the other hand, a sizeable part of all of the viral RNAs were destroyed by nuclease even in total absence of detergents, although a distinct proportion of both positive and negative strand viral RNAs were protected from nuclease action following treatment with the ionic detergent DOC (58), which was attributed by these workers to the presence of a membrane barrier(s).

Inclusion of a trypsin digestion step at critical points during our manipulations suggested the involvement of protein(s) in protecting the RF from nuclease action even after solubilization of all of the membranes with DOC. In KUNV, all of the viral NS proteins with the exception of NS2b could be co-immunoprecipitated using anti-dsRNA antibodies (15). Therefore, it is very likely that these viral NS proteins that constitute the RC interact with RF and consequently afford protection against nucleases in the case of WNV, JEV, and also DENV. The number of replication forks present on one RI molecule is 6–7 for DENV (60), resulting in the simultaneous presence of 6–7 RC on the template, which could potentially protect the RF from degradation. Since the number of replication forks vary among flaviviruses (61), it is difficult to predict the same for WNV and JEV, viruses for which this information is presently not available. In addition to viral proteins, it is also possible that unknown host protein(s) interact with RF. Our use of the milder agent sodium citrate to solubilize the bounding rough endoplasmic reticulum followed by sequential treatment with trypsin and MNase revealed that vRNA too was protected by protein(s), albeit in a relatively loose manner since these proteins could be removed by detergents. Although the role of proteins in conferring nuclease resistance was not investigated in polio and BMV, the exposure of transmissible gastroenteritis conavirus-negative strands to nuclease was reported to be unaffected by protease treatment in the absence of detergents (59). The concerted/sequential action of detergents and proteases, which in our studies revealed with clarity the relative roles of proteins and membranes in protecting viral RNAs from nucleases, are yet to be investigated for other positive-strand RNA viruses.

The differential orientation of the two flaviviral RNAs RF and vRNA, reflects the function they perform. The vRNA is the template for both translation as well as negative-strand synthesis and has to be packaged to form virus particles. It has been proposed that in the postlatent phase, the translation of viral RNA predominantly takes place in the heavy membrane fractions (62). The presence of the viral protease complex in the CM (15) revealed this to be the site for polyprotein processing within the same heavy membranes. The close association of VP through the rough endoplasmic reticulum connections with the CM/PC (Fig. 6) (15) reveals an additional level of organization adopted by flaviviruses that would enhance the efficiency of protein synthesis using vRNA synthesized within VP as the template followed by subsequent processing of the polyprotein. However, as noted earlier (22), clarity is wanting in our understanding of the crucial step of release of vRNA into the cytosol for the purpose of translation as well as packaging (Fig. 6). The organization of the flaviviral RC revealed by our studies could in fact help to concentrate precursors vital for RNA synthesis provided efficient transporters are present and thereby increase the efficiency of replication. In this regard, the recent identification of poliovirus 2B protein as a viroporin that allows passage of solutes (63) as well as the reported increase in permeability of bacterial membranes upon expression of small hydrophobic JEV proteins (54) suggests strategies adopted by viruses to facilitate communication between the host cytosol and the membranous compartments containing the viral RC.

The intricate mechanism adopted by flaviviruses to encase the dsRF behind two membranes emphasizes the need for the virus to prevent or reduce the exposure to dsRNA-mediated host defenses such as protein kinase R and RNase L as well as RNA interference. Such a placement of RF therefore points to the vital function it plays as the template, which needs to be protected and sequestered from the deleterious effects of the host defense mechanism. Additionally, this retention of RF inside the VP not only allows the reuse of RF, aptly called the recycling template (23), but also helps in maintaining template specificity, making the whole process of replication highly efficient. In conclusion, our study on the organization of flaviviral RNA in the RC provides valuable insights that would impact on design of potential therapeutics and inhibitory agents aimed at targeting the most critical component of the viral life cycle, namely replication.


    FOOTNOTES
 
* This work was supported by a grant (SP/SO/D-76/97) from the Department of Science and Technology, Government of India. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} Recipient of a senior research fellowship from the Council of Scientific and Industrial Research. Back

§ To whom correspondence should be addressed: Dept. of Microbiology and Cell Biology, Indian Institute of Science, Bangalore 560012, India. Tel./Fax: 91-80-2932685; E-mail: vijaya{at}mcbl.iisc.ernet.in.

1 The abbreviations used are: NS, nonstructural; ds, double-stranded; RF, replicative form; RI, replicative intermediates; RdRp, RNA polymerase; RC, replication complex; KUNV, Kunjin virus; DENV, dengue virus; CM, convoluted membranes; PC, paracrystalline arrays; VP, vesicle packets; JEV, Japanese encephalitis virus; WNV, West Nile virus; p.i., postinfection; MNase, micrococcal nuclease; TX100, Triton X-100; ER, endoplasmic reticulum; DOC, sodium deoxycholate; SMS, smooth membrane vesicle-like structures; BMV, brome mosaic virus. Back


    ACKNOWLEDGMENTS
 
We thank Dr. Priti Kumar for constant help and valuable discussions throughout the course of this investigation. We also acknowledge the help extended by electron microscope facility of the Department of Microbiology and Cell Biology for ultrastructural analysis. Mridula Nandan, Bilwa Dasarathi, and K. S. Ananda are acknowledged for excellent technical assistance.



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