Architecture of the flaviviral replication complex: protease, nuclease and detergents reveal

encasement within double-layered membrane compartments

Pradeep Devappa Uchil¶ and Vijaya Satchidanandam*†

Department of Microbiology and Cell Biology, Indian Institute of Science

Bangalore 560012, INDIA

* Corresponding author:

Vijaya Satchidanandam

Department of Microbiology and Cell Biology

Indian Institute of Science

Bangalore-560012

INDIA

Tel: 91-80-3942685

Fax: 91-80-3942685

E-mail: vijaya@mcbl.iisc.ernet.in

Running title: Viral RNA organization in flaviviral replication complexes

Copyright 2003 by The American Society for Biochemistry and Molecular Biology, Inc.

JBC Papers in Press. Published on April 16, 2003 as Manuscript M301717200 by guest on January 24, 2020

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Summary

Flavivirus infection causes extensive proliferation and reorganization of host cell membranes to form specialized

structures called convoluted membranes/paracrystalline arrays (CM/PC) and vesicle packets (VP), the latter of

which are believed to harbor flaviviral replication complexes (RC). Using detergents, 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 non-ionic 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 (RI)

RNA. The RF co-sedimented at high speed from non-ionic 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 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 non-ionic detergent-resistant vesicles.

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INTRODUCTION

While 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- and posttranslationally by the host signalase and a virus-encoded proteinase to give three structural

(capsid [C], premembrane/membrane [prM/M] and envelope [E]), and seven nonstructural (NS) proteins (NS1 to

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 the viral proteins, functions as the RNA

dependent RNA polymerase (RdRp; 5-7) and a methyl transferase (8), the latter implicating its role in capping of

viral genomic RNA. The multifunctional protein NS3 manifests three activities; the viral protease along with the

cofactor NS2b critical for proper processing of the viral polyprotein (9-11), a helicase required most probably for

unwinding dsRF (12), and an NTPase activity (13); presumably 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’ UTR 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).

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The RNA-synthesizing machinery of virtually all 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). Cryoimmunoelectron microscopy (CIEM) 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 rough endoplasmic reticulum (RER; 22). The CM/PC originate from membranes derived from

intermediate compartments (IC) 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- 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 RNA polymerase (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, the first being the presence of a bounding membrane that protects the enzymatically active replicase

from protease action and the second, the requirement for only catalytic amounts of replicase proteins. In the present

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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

Viruses and Cells—WNV strain E101, JEV strain P20778 (Genbank Accession number AF080251), and

DEN-2 virus strain TR 1751 [National Institute of Virology (NIV), Pune, India] were propagated in the Aedes

albopictus cell line, C6/36 [National Centre for Cell Science (NCCS), Pune, India] in Minimum Essential Medium

(MEM, Gibco BRL) supplemented with 5% fetal bovine serum (FBS), 0.3% tryptose phosphate broth (DIFCO

Laboratories), 0.22% NaHCO3 and 2 mM HEPES (pH 7.3). Confluent monolayers of C6/36 were infected with virus

at a multiplicity of infection (m.o.i.) 0.1 for routine expansion and medium-containing virus was harvested at 5-½ d

post infection (p.i.), aliquoted and stored at –80ºC till further use. The porcine kidney cell line PS (NCCS)

maintained at 37ºC in MEM with 10% FBS, 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

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 × g at 18-22 h p.i. and used to obtain heavy membrane fractions sedimenting at

16,000 × g (P16) as 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% polyacrylamide gel electrophoresis (urea-

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

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RNA species generated was estimated by scanning the gels on a Fuji BAS1000 phosphorimager and analyzed using

the Fuji MacBAS V2.4 software.

Metabolic labeling of proteins—Subconfluent monolayers of PS cells were mock or flavivirus-infected at

an m.o.i. of 10. At 16 h p.i., the cells were labeled with 50 µCi per ml of [35S] methionine-cysteine (EXPRE35S35S,

NEN, 1175.0 Ci/mmol) as described previously (29). Protein samples after appropriate treatments were analysed on

SDS-10% PAGE. The gels were processed for fluorography using AMPLIFYTM (Amersham Pharmacia biotech)

according to manufacturers 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 (NEN) for 1 h in presence of 3 µg/ml actinomycin D

(AMD). 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 of micrococcal nuclease (MNase,

MBI Fermentas) and 20 units/ml of DNase I (Roche) in presence of 1 mM CaCl2 at 30ºC for 30 min. The treatments

were terminated by adding ethylene glycol-bis(beta-aminoethyl ether)-N, N, N’, N’-tetraacetic acid (EGTA) pH 8.0

to 5 mM and holding on ice for 30 min. Trypsin (Promega Corporation, sequencing grade) treatment was carried out

on ice for 15 min at the concentrations mentioned and terminated using soybean trypsin inhibitor (GIBCO BRL) and

phenylmethylsulfonyl fluoride (PMSF, 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 hour. The non-ionic 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 4ºC for 30 min (34). The protein concentration in the homogenates and P16 fractions during all treatments was

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maintained at 2 mg/ml. All the detergents used were nuclease-free molecular biology grade obtained from Sigma-

Aldrich.

Floatation analysis— The P16 fraction after in vitro RdRp assay from mock- and flavivirus-infected cells

were subjected to 1% TX100 or 1.5% DOC at 4ºC for 1 h, followed by sedimentation at 16,000 × g for 15 min to

obtain S16 fractions which were used for floatation analysis. Briefly, 0.5 ml (2 106 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) and 1 ml of 5% (wt/wt) sucrose in TNMg buffer. 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, RNA extracted and analyzed as mentioned above.

Electron microscopy of TX100-resistant membrane structures—Detergent treated S16 fraction obtained as

above were subjected to ultracentrifugation at 35,000 rpm (150,000 × g) for 5 h. The pellet (P150) obtained

(detergent-resistant membrane fraction) was resuspended in ice-cold phosphate buffered saline (PBS) and deposited

on formvar coated copper grids (Ted Pella Incorporated) 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 Incorporated) according to manufacturer’s instructions after fixing the samples

with 3.7% paraformaldehyde (TAAB Laboratory Equipment) and 0.01% glutaraldehyde (Sigma, EM grade) in PBS.

Ultra thin sections were then incubated at room temperature as follows: 2 h in PBG [PBS containing 0.1% (w/v)

BSA, 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 10 min in PBG; 2 h in anti-rabbit IgG (H + L)

antibodies coupled to either 15 or 10 nm gold (Ted Pella Incorporated) diluted 1:100 in PBG; 5 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.

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RESULTS

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 2-3 for WNV and 5-6 for DENV) despite the near complete

destruction of the metabolically labeled major replicase proteins NS5 and NS3 (Fig. 1B, lanes 1-2 for WNV and 5-6

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, which was similar to the properties of JEV RC previously demonstrated by us (29). The suggested

existence of a membrane barrier was tested using the non-ionic 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 4-6 for DENV) over and above the 30% reduction in activity observed due

to the detergent treatment alone (Fig. 1C, lanes 1-2 for WNV and lanes 3-4 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 due to the complete loss of

activity suffered by JEV RC following detergent treatment alone (Fig. 1C, lanes 5 and 6; 29). This could be due

either to the greater inherent inhibition of JEV RC compared to WNV and DENV by TX100 or selective loss of one

or more factors from JEV RC, possibilities that are under investigation. While DOC treatment did not adversely

affect RdRp activity (Fig. 1D, lanes 7, 10 and 13), it however led to complete loss of activity when followed by

trypsin in all 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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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 non-specific nuclease such as micrococcal nuclease (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 towards the lumenal 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 (compare

lanes 5 and 6 in Fig. 2A and lanes 1 and 2 in Fig. 2B) 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 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, since

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 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 in order 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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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 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 AMD-treated and 35S-

methionine labeled proteins from virus-infected cells as shown in figure 1B, lane 2. Similar nuclease-resistance of

all three JEV and DENV RNA species was also observed (data not shown). Thus our results suggested that all 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.

Non-ionic 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, which

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 non-ionic 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 non-ionic detergent and MNase (Fig. 3A, lane 2), compared to samples treated with detergent alone (Fig. 3A,

lane 1). The residual label in the wells following MNase treatment represents insoluble and non-specific aggregates

since labeled RI RNA free of membranes and proteins is fully susceptible to MNase as seen in figure 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 non-ionic 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 render 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 non-ionic detergents as well

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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.

The non-ionic 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

non-ionic 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 non-ionic 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 atleast 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

figure 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 of trypsin with complete loss of full-length intact RF achieved at the highest concentration of trypsin used

(Fig. 4B, lanes1-6). This was in contrast to the inability of trypsin to facilitate access to the RF for MNase following

non-ionic 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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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 × g (24,25). The differential solubility of the outer ER-like membranes alone to non-ionic

detergent should as a result release all the inner vesicles, which being smaller would no longer be expected to

sediment at 16,000 × g. Indeed, treatment of P16 fractions with 1% TX100 followed by sedimentation at 16,000 × g

showed that approximately 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 non-ionic detergent-resistant membrane structures. Successful co-sedimentation of

the major replicase proteins NS3 and NS5 in all three flaviviruses studied with RF at 150,000 × 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-solubilised RNA from P16 fractions of WNV-infected

cells did not sediment at 150,000 × g, proving that this detergent completely solubilised membranes housing the RC

(Fig. 4C, lanes 4-6). In contrast, DOC-solubilised RNA from P16 fractions of WNV-infected cells did not sediment

at 150,000 × g, proving that this detergent completely solubilised 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 non-ionic detergents (35). We were however unable to verify this difference in our

laboratory under similar conditions since 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,

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as monitored by the presence of radiolabeled RF in these fractions (Fig. 5A, middle panel). More than 70% of

radiolabeled RF from TX100 extracts was however 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 due to

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 owing to its increased

sensitivity. Similar results were obtained for TX100 extracts of P16 membranes from JEV-infected cells (data not

shown).

Electron microscopic analysis revealed vesicular structures measuring 75-100 nm (Fig. 5, B) in ultra-

sedimented fractions of TX100 extracts from WNV-infected cells, a size similar to that previously reported for

structures enclosed within bounding RER 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, which

suggested its solubilization by non-ionic 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 × 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 ultra-sedimented 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 harbour viral RNA have been in fact observed previously in closely related togaviruses,

mouse hepatitis and poliovirus (37-39). The mechanisms by which these vesicles interact with their host

environment for obtaining precursors for and releasing products of RNA synthesis remains 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

cryoimmunoelectron microscopy inside KUNV- and DENV-infected cells (15,21).

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DISCUSSION

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 (46-51). 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

till date. This study thus 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 since trypsin treatment, even in the absence of detergents, destroyed most of the major replicase proteins

NS5 and NS3 and other small non-structural 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

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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 CIEM (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 into

them of viral and/or associated host proteins (54). Furthermore, detergent-resistance can be conferred by a high

proportion of lipids like cholesterol or glyco-sphingolipid in these membranes (55) as also by specific post-

translational modification of proteins such as acylation and glycosyl-phosphatidylinositol-anchoring (GPI) which are

known to render the membranes resistant to non-ionic 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 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

(57).

The presence of double-layered membrane vesicles that harbour 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,

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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 (TGEV) 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. While 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 harbour the BMV RC (59). In the coronavirus TGEV on the other hand, a sizeable part of all 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), that 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 the membranes with DOC. In

KUNV, all the viral NS proteins except NS2b could be co-immunoprecipitated using anti-dsRNA antibodies (15). It

is therefore 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 DENV also. 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 RER 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. While the role of proteins in conferring nuclease resistance was not investigated in

polio and BMV, the exposure of TGEV 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

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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 post-latent 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

RER connections, with the CM/PC (15; Fig. 6) 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 PKR and RNase L as

well as RNA interference (RNAi). 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.

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FOOTNOTES

This work was supported by a grant (SP/SO/D-76/97) from the Department of Science and Technology,

Government of India.

¶ PDU was a recipient of a senior research fellowship from the Council of Scientific and Industrial Research

† To whom the correspondence should be addressed: Department of Microbiology and Cell Biology, Indian Institute

of Science, Bangalore-560012, INDIA. Tel: 91-80-3942685; Fax: 91-80-3942685; E-mail: vijaya@mcbl.iisc.ernet.in

ACKNOWLEDGEMENTS

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

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biology for ultrastructural analysis. Mridula Nandan, Bilwa Dasarathi and K. S. Ananda are acknowledged for

excellent technical assistance.

FIGURE LEGENDS

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 to 1 mg/ml) as depicted in the flow chart before carrying out the RNA dependent RNA polymerase (RdRp) assays

using [ -32P]GTP. The labeled RNA products generated were resolved on a partially denaturing 7 M urea 3%

polyacrylamide gel electrophoresis (urea-PAGE). (B) Effect of in vitro trypsin treatment on metabolically labeled

flaviviral proteins. P16 fractions metabolically labeled with 35S-methionine-cysteine from AMD-treated WNV

(lanes 1-4) and DENV (lanes 5-8) infected cells were subjected to trypsin (1mg/ml) without (lanes 2 and 6) or with

prior treatment with 1% sodium deoxycholate (DOC; lanes 4 and 8) or 1% triton X-100 (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 weight size markers are mentioned on the right. (C) Effect of TX100 on in vitro flaviviral RdRp activity.

P16 fractions from WNV (lanes 1-2), DENV (lanes 3-4) or JEV (lanes 5-6) were treated (T; lanes 2, 4 and 6) or not

treated (N; lanes 1, 3 and 5) with 1% TX100 for one hour on ice followed by in vitro RdRp assay using [ -

32P]GTP. The labeled RNA products after extraction were analyzed using urea-PAGE. (D) P16 fractions from

flavivirus infected cells were processed as depicted in the flow chart and RdRp assays 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 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.

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

micrococcal nuclease (MNase) prior to carrying out in vitro RNA dependent RNA polymerase (RdRp) assay using

[ -32P]GTP. The labeled RNA products were resolved using urea-PAGE. Lanes 1-3 are ethidium bromide-stained

gel photograph of the same gel whose autoradiogram is shown in lanes in 4-6. Lanes 7 and 8 show the MNase

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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 [ -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 hrs for WNV and JEV and 24

hours for DENV.

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 [ -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 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 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 weight

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 flowchart. The RNA samples after extraction were

analyzed using urea-PAGE. The arrowheads in A, B and D denote the positions of RI, vRNA and RF.

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FIG. 4. (A) P16 fractions treated with 1.5% sodium deoxycholate (DOC) were sedimented at 16,000 × 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 urea-PAGE. (B) DOC treated P16 fractions were subjected to increasing concentrations of

trypsin following [ -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 urea-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 [ -32P]GTP (T; lane 1) were sedimented at 16,000 × 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 × 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 which were analyzed using urea-PAGE. (D) The metabolically labeled proteins released into the

supernatant fractions from TX100 treated heavy membranes from WNV, DENV and JEV were subjected to ultra

centrifugation at 150,000 g to obtain proteins associated with detergent resistant vesicles (UP; pellet) and those

which were completely solubilised (US; supernatant) by the detergent. Equivalent amounts of pellet and supernatant

fractions were analyzed and the proteins visualized by autoradiography. The dots on the right represent the major

replicase proteins NS3 and NS5. The positions of standard molecular weight size markers are shown on the left.

FIG. 5. Characterization of TX100-resistant membranous structures. (A) P16 fraction (lower panel; P16) and its

TX100 (top panel; S16TX100) and DOC (middle panel; S16DOC) extracts after RdRp assay with [ -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 urea-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

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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.

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.

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0

DENV

1.00.51.00.50Trypsin

WNV

-

-

-

M

+

-

+---+---1% DOC

+

+

+

+

-

-

WNV

+-++Trypsin

----1% TX100

DENVVirus

NS5

NS3

ENVNS1

NS4bprM

NS2a/4a

CNS2b

97.4

66

46

21

kDa

14

A. B.

1 2 3 4 5 6

RF

RI

vRNA

1 2 3 4 5 6 7 8 9P16

fractionTrypsin

treatmentRdRp

assay

Inactivate

trypsin

TN

JEV

TNTNTX100

DENVWNVVirus

0.5 1.0001.50.001.50.01.0.5 001.0.5 00Trypsin

---++TX100

+++--DOC

JEVDENVWNVDENVWNVVirus

C. D.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 151.0 0.52 0.48 1.0 0.48 0.40 1 0 0 1 0 0 1 0 0

vRNA

RF

RI

RdRp

assay

Trypsin

treatmentInactivate

trypsin

P16

fractionTX100/DOC

treatment

RI

vRNA

RF

1 2 3 4 5 6 1.0 0.70 1.0 0.75 1.0 0

FIG. 1

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A. B.

−+−−+−−+−Assay3

++−++−++−MNase2

+−++−++−+Assay1

DENVJEVWNVVirus

M

MNase

IIIII

+−−+−+−-

MI

32P labelEtBr

RI

RI

vRNA

vRNA

RF

RF

1 2 3 4 5 6 7 8 91 2 3 4 5 6 7 8

C. D.

Trypsin (mg/ml)WNV JEV DENV Mock

MNase - + - + - +

0.0 0.5 1.0MNase − + − + − + − +

1 2 3 4 5 6 7 8

RI

1 2 3 4 5 6

RI

RdRp Assay

Trypsin treatment

Inactivate trypsin

± MNase

vRNAvRNA

RFRF

FIG. 2

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A. B.

RI

vRNA

RF

++−MNase

+−−Trypsin

(1mg/ml;Inactivated)

+++1% TX100

RI

vRNA

RF

1.3 83.71.83.41.3.2RI

+-+-+-MNase

1.00.5-Trypsin

1% TX100

RdRp assay

Inactivate

trypsin

± Trypsin

treatment

± MNase

Detergent

treatment

1 2 3

654321Lanes

7.6 15.77.65.27.5.6RF

C. D.

1% Sodium

Citrate

+−+−+−MNase

1.00.5−Trypsin

UntreatedSource

RI

vRNA

RF

1 2 3 4 5 6

Inactivate

trypsin

Trypsin

treatment

± MNase

Citrate

treatment

RdRp

Assay

NS5

NS3

ENS1

NS4bprMNS2a/NS4a

NS2bC

97.4

66

21

kDa

14

TX100 T P S

46

1 2 3

FIG. 3

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A. B.

RI

vRNA

RF

1 2 3 4 5 6

PelletSupernatant

−+−−+−3) Assay

++−++−2) MNase

+−++−+1) Assay

DOC treated P16

Inactivate

trypsin

RdRp

AssayTrypsin

treatment

DOC

treatment± MNase

32P label

EtBrRF

RF

+−+−+−MNase

1.00.50.0Trypsin

1 2 3 4 5 6

C.

WNV DENV JEV

UP US UP US UP US

NS5

NS3

D.

97.4

66

46

21

kDa

14

TX100

16K

T P S

DOC TX100

Ultra Ultra

S16 S P S16 S P

RI

vRNA

RF

4 5 6 7 8 91 2 3 1 2 3 4 5 6

FIG. 4

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A.B. C. D.

1 2 3 4 5 6 7 8 9 10 Fraction

S16TX100

RI

RF

RI

G. H.

E. F.

S16DOC

RF

P16

RI

vRNA

RF

Top Bottom

FIG. 5

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CM/PC

Polyprotein

processing

Release of

vRNA into

cytosol???

Protein

synthesis

ER connection

between VP

and CM

Vesicle packets

(non-ionic

detergent resistant)

Nascent single-

stranded RNA

Packaging

Outer TX100-sensitive

membrane

Flaviviral RC

DsRF bound

by proteins

FIG. 6

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Pradeep Devappa Uchil and Vijaya Satchidanandamreveal encasement within double-layered membrane compartments

Architecture of the flaviviral replication complex: protease, nuclease and detergents

published online April 16, 2003J. Biol. Chem. 

  10.1074/jbc.M301717200Access the most updated version of this article at doi:

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