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Journal of Virology, July 2000, p. 6570-6580, Vol. 74, No. 14
0022-538X/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.

Formation of the Poliovirus Replication Complex Requires Coupled Viral Translation, Vesicle Production, and Viral RNA Synthesis

Denise Egger,1 Natalya Teterina,2 Ellie Ehrenfeld,2 and Kurt Bienz1,*

Institute for Medical Microbiology, University of Basel, Basel, Switzerland,1 and Laboratory of Viral Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, Maryland2

Received 23 December 1999/Accepted 17 April 2000


    ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Poliovirus (PV) infection induces the rearrangement of intracellular membranes into characteristic vesicles which assemble into an RNA replication complex. To investigate this transformation, endoplasmic reticulum (ER) membranes in HeLa cells were modified by the expression of different cellular or viral membrane-binding proteins. The membrane-binding proteins induced two types of membrane alterations, i.e., extended membrane sheets and vesicles similar to those found during a PV infection. Cells expressing membrane-binding proteins were superinfected with PV and then analyzed for virus replication, location of membranes, viral protein, and RNA by immunofluorescence and fluorescent in situ hybridization. Cultures expressing cellular or viral membrane-binding proteins, but not those expressing soluble proteins, showed a markedly reduced ability to support PV replication as a consequence of the modification of ER membranes. The altered membranes, regardless of their morphology, were not used for the formation of viral replication complexes during a subsequent PV infection. Specifically, membrane sheets were not substrates for PV-induced vesicle formation, and, surprisingly, vesicles induced by and carrying one or all of the PV replication proteins did not contribute to replication complexes formed by the superinfecting PV. The formation of replication complexes required active viral RNA replication. The extensive alterations induced by membrane-binding proteins in the ER resulted in reduced viral protein synthesis, thus affecting the number of cells supporting PV multiplication. Our data suggest that a functional replication complex is formed in cis, in a coupled process involving viral translation, membrane modification and vesicle budding, and viral RNA synthesis.


    INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Infection of cells with poliovirus (PV) induces a number of biochemical and morphological modifications to cellular proteins and structures that are required to promote efficient replication of the virus. Previous studies have described the extensive rearrangement of membranes from intracellular organelles that generates masses of membranous vesicles that support viral RNA synthesis (11, 14, 17) and possibly encapsidation (35, 44, 48). All viral proteins required for RNA replication, as well as newly synthesized viral RNA, are associated with the surfaces of these virus-induced vesicles. Upon isolation from infected cells, they form a rosette-like cluster surrounding and enclosing the replication complex (12, 15). These isolated crude replication complexes are active in viral RNA synthesis in vitro (12, 28, 58). The membrane structure appears to be essential for the initiation steps of the RNA synthesis reaction but is not required for RNA chain elongation (27). The association of replication complexes with membranous structures appears to be a general feature of plus-strand RNA viruses (references 32, 41, 47, 50, and 55 and references therein).

Formation of the PV replication complex probably provides the structural basis for rapid and efficient RNA replication; it also causes a severe structural reorganization of the cell, leading ultimately to cytopathology and cell death. Thus, cytopathology is related to virus replication. Under restricted viral growth conditions, apoptosis serves as a second mechanism for virus-induced cell death (2, 62). Apoptosis is generally believed to be a defense mechanism of the host that limits virus replication. Both types of cell death, however, may cause clinical symptoms in vivo (31).

The generation of the vesicles associated with the PV replication complex depends on the production of viral nonstructural proteins (8, 13, 39). Expression of individual viral nonstructural gene products in mammalian cells revealed that many of these proteins have multiple functions (references 6, 7, 30, 36, 45, 49, 66, and 69 and references therein). Proteins containing 2B, 2C, or 3A sequences have membrane-binding properties and associate with cellular membranes in the absence of other viral proteins (4, 9, 18, 21, 22, 24, 25, 54, 63, 66). Proteins 2B and 2BC thereby change the permeability of the plasma membrane and cause Ca2+ ions to redistribute from the endoplasmic reticulum (ER) during infection (3, 5, 33, 66, 67). Proteins 2B and 3A interfere with membrane traffic through the Golgi complex (23, 54), and proteins 2BC and 2C reorganize the structure of the intracellular membranes (18). Expression of protein 2BC induces vesicles quite similar in appearance to those formed during PV infection (18).

Characterization of the PV-induced vesicles with immunological probes demonstrated the presence of cellular markers of the ER, Golgi complex, and lysosomes, in addition to the viral P2/P3 proteins (27, 56). Apparently, ER membranes are used to form vesicles initially (11), while markers for the Golgi complex colocalize with virus-induced vesicles later in infection (16). Nuclear or cell surface membranes do not appear to contribute to the formation of vesicles. Rather, the membranes participating in vesicle formation apparently all originate from organelles involved in the secretory pathway in uninfected cells.

Exocytotic intracellular membrane traffic starts by formation of transport vesicles, which are induced by a layer of complexed proteins (coat protein complex II). This set of proteins is responsible for deforming the ER membrane and causing the vesicles to bud off from the ER (reviewed in references 51 and 57). Budding of virus-induced vesicles from the rough ER (rER) might, in principle, be based on this cellular process, adapted, and possibly enhanced by the virus. The exact mechanism of viral vesicle formation, however, is not known.

In contrast, the formation of the vesicle-containing replication complex for RNA-dependent RNA synthesis results in an intricate virus-specific organelle with no structural or functional counterpart in an uninfected cell. For the formation of the replication complex, certain functions of the protein secretory pathway, notably brefeldin A-sensitive steps, appear to play an important role. This was inferred from the finding that brefeldin A, an inhibitor of ADP-ribosylation factor 1 (ARF1) function, is an inhibitor of PV RNA synthesis in vivo (34, 42) and in a cell-free system (20).

In this study, we investigated whether modified membrane structures induced by expression of viral or cellular membrane-binding proteins might act as substrates for vesicle formation during a subsequent PV infection. We also looked for the ability of vesicles induced by PV protein 2BC or the entire set of P2 and P3 proteins to participate in the formation of replication complexes for the replication of PV.

We found that ER-derived, smooth-surfaced membranes of different morphology and protein content are not utilized to form vesicles. Vesicles induced by the expression of PV proteins and closely resembling vesicles found in PV-infected cells do not participate in the formation of replication complexes and do not associate with replication complexes engaged in RNA synthesis. Our findings suggest that a functional replication complex is formed in a coupled process involving viral translation, membrane modification and budding, and viral RNA synthesis.


    MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Plasmids. For the transient expression of viral and cellular proteins, pTM plasmids containing an encephalomyocarditis virus (EMCV) internal ribosome entry site (IRES) were used (29) (Fig. 1). The construction of plasmids pTM-2BC, pTM-2C, and pTM-2C(K135S) is described in reference 18, that of plasmid pTM-2C(1-274) is described in reference 59, and that of pTM-Fg-3AB, pTM-Fg-3AB DII 3E, pTM-Fg-Cyt b5, and pTM-Fg-beta -globin is described in reference 63. The Flag sequence (Kodak, Rochester, N.Y.) was introduced at the N terminus of the 2BC protein by PCR amplification of the PV cDNA (nucleotides [nt] 3833 to 4653) with the sense primer 5'-AACAACAACATATGGGAGACTACAAGGACGACGATGACAAAGGCCTCACCAATTACATAG-3', containing a NdeI site (underlined) and the Flag sequence (bold), and the antisense primer 5'-TCGTCCATAATCACCACTCC-3'. The resulting fragment was cut with NdeI and SpeI and inserted back into the plasmid to obtain pTM-Fg-2BC. To add the Flag sequence to wild-type (wt) 2C, 2C(1-274), and 2C(K135S), the sense 5'-TATGGGTGACTACAAGGACGACGATGACAAGGGTGACAGCTGGTTGAAGAAGTTTACTGAAGCATG-3' and antisense 5'-CTTCAGTAAACTTCT TCAACCAGCTGTCACCCTTGTCATCGTCGTCCTTGTAGTCACCCA-3' oligonucleotides (NdeI and SphI sites are underlined; the Flag sequence is in bold) were annealed and ligated into the corresponding plasmids, cut with NdeI and SphI.


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  		FIG. 1.   Schematic representation of the constructs used in this study. PV and pPVDelta P1 have authentic PV 5' NTRs and are replication competent. pE5PVDelta P1 and the remainder of the constructs contain the EMCV IRES instead and are not replication competent. The construction of the plasmids is described in Materials and Methods. *, mutations; Fg, Flag sequence; An, poly(A)tail.

The construction of plasmids pE5PVDelta P1 and pPVDelta P1 is described elsewhere (unpublished data). Plasmid pE5PVDelta P1 contains PV type 1 cDNA from plasmid pT7-PV1(SalI) (10) starting from nt 3371 of PV and extending to the end of the poly(A) sequence, placed downstream of the EMCV IRES in plasmid pTM-1. Plasmid pPVDelta P1 is a derivative of plasmid pT7-PV1(SalI) with a deletion of the entire P1 coding region.

Infection and transfection of HeLa cells and expression of proteins. To express plasmid-encoded proteins, HeLa cells were infected with vaccinia virus vTF7-3 and transfected with 10 µg of DNA per 3.5-cm2 plate using Lipofectin (Life Technologies, Gaithersburg, Md.) as described previously (59). Supertransfection of already transfected cells was done as described above for the first transfection but omitting vTF7-3 infection. The efficiency of the second transfection was generally lower than that of the first. To infect cells with PV, 30 PFU of Mahoney type 1 virus per cell was adsorbed at 36°C for 30 min and the infection was left to proceed for 5 h. Superinfection of already transfected cells with PV was identical.

Time course series were used to determine the optimal expression time of membrane-binding proteins before superinfection or supertransfection. This was found to be 6 to 7 h (see Table 1 and Results).

Ab and IF. For indirect immunofluorescence (IF), cells were grown and infected or transfected on glass coverslips, fixed with paraformaldehyde, and permeabilized as described previously (16). For the simultaneous detection of two antigens, the cell preparations were incubated with an appropriate mixture of primary antibodies (Ab) from two different animal species, washed, and incubated with a cocktail of corresponding antispecies antibodies, each of which was labeled with a different fluorochrome. After another wash, coverslips were mounted in Tris-glycerol (pH 8.5) containing 2.5% 1,4-diazabicyclo(2.2.2)octane (Sigma, Buchs, Switzerland) (65) or 1% n-propyl gallate (Sigma) (40).

The following Ab and dilutions were used: anti-Flag M2 monoclonal Ab (MAb) (Sigma), diluted 1:200; anti-PV 2B 1D3.B1 MAb (27), diluted 1:3; anti-PV 2C rabbit antiserum (18), diluted 1:100; anti-PV VP1 MAb B3/H.2 (48), diluted 1:4; anti-PV VPg rabbit antiserum raised against a synthetic peptide comprising the entire 22 amino acids of VPg (a gift of L. Pasamontes), diluted 1:100; anti-CD 155 (NeoMarker, Fremont, Calif.), diluted 1:100; goat anti-mouse Ab coupled to Texas red (Molecular Probes, Eugene, Oreg.), diluted 1:200; and goat anti-rabbit Ab coupled to fluorescein isothiocyanate (FITC) (Sigma), diluted 1:80.

PV-infected cells were quantitated by counting VPg- or 2C-positive cells within the subpopulation of productively transfected Flag-positive cells (see Table 1).

For the quantification of PV infection in pE5PVDelta P1-transfected cells, cultures in petri dishes containing several coverslips were transfected with the pE5PVDelta P1 DNA and incubated for 6.3 h. Before superinfection with PV, a coverslip was removed from each dish and further incubated separately for the determination of the efficiency of expression by appropriate IF. In the superinfected culture incubated in parallel, the number of PV infected cells was counted by IF using the anti-VP1 MAb.

RNA probes and FISH. The single-stranded RNA (ssRNA) probe of minus polarity, complementary to nt 6012 to 6736, was prepared and labeled with FITC-UTP (Roche Molecular Biochemicals, Mannheim, Germany) during in vitro transcription with T7 RNA polymerase from a DNA template (26). The probe was hydrolyzed and purified as described previously (16, 26).

This probe, nt 6012 to 6736, is complementary to a part of the 3D genomic region. There was no cross-reactivity with P2 sequences, as shown by applying the probe to cells which were transfected with DNA encoding protein 2BC. The fluorescent in situ hybridization (FISH) protocol to detect RNA of plus polarity has been described in detail previously (16, 26).

For double-labeling ISH-IF, IF was performed in an indirect assay after completion of FISH. To preserve the viral antigen, DNase digestion of the transfected DNA and thermal denaturation of the specimen were omitted from the FISH protocol. The specificity of this modified procedure was verified in HeLa cells transfected with pE5PVDelta P1 DNA but not coinfected with vTF7-3, to avoid transcription of the plasmid DNA into RNA. Plasmid DNA was detectable only after thermal denaturation. The labeling pattern, however, was found to be different from the RNA pattern observed when RNA was transcribed (data not shown). Without a denaturation step prior to FISH, the probe complementary to nt 6012 to 6736 did not detect any double-stranded DNA (dsDNA). Likewise, the probe did not produce a signal if the specimen was treated with DNase (1 U/µl at 37°C for 60 min) prior to denaturation and FISH.

Electron microscopy and confocal laser-scanning microscopy (CLSM). For electron microscopy, cell cultures were trypsinized, fixed with 2.5% glutaraldehyde and 2% OsO4, and embedded in Epon 812 by standard procedures. Sections were viewed in a Philips CM 100 electron microscope. For confocal laser-scanning microscopy (CLSM), a Leica TCS4D microscope was used with the photomultiplier settings adjusted to avoid bleeding from one channel into the other. Raw images were adjusted for contrast and background staining with Adobe Photoshop software.


    RESULTS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Morphological changes of cytoplasmic membranes by the expression of viral and cellular proteins. Expression of cellular or viral membrane-binding proteins, mediated by recombinant vaccinia viruses producing T7 RNA polymerase, was used to change the architecture of the membranes of the ER. During a subsequent PV infection, we investigated whether the altered membranes, made immunologically traceable with an N-terminal Flag epitope, could be transformed into vesicles and whether such membranes could contribute to the formation of a viral replication complex.

Figure 2 shows that the aspects of the altered membranes range from myelin-like threads and whirls to smooth vesicles. The cellular protein cytochrome b5 (Cyt b5) induces concentric myelin-like membrane sheets (Fig. 2a). PV membrane-binding proteins 2C (and certain domains thereof [18, 59]) and 3AB transform rER into membrane configurations not found in PV-infected cells (Fig. 2b and c). In contrast, protein 2BC, the K135S mutant of 2C, and peptides consisting of the first 274 amino acids of wt or mutated 2C induce vesicles closely resembling those arising during a PV infection (18, 59) (Fig. 2d to f). Thus, the membrane-binding proteins generated smooth-surfaced (i.e., ribosome-free) membranes and, concomitantly, the rER was drastically reduced. Proteins that do not bind to membranes, such as a 3AB mutant (3AB DII 3E [63]) or beta -globin, did not induce membrane alterations (data not shown).


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  		FIG. 2.   Electron micrographs of membrane alterations in HeLa cells. Cell cultures were transfected with plasmids expressing the following membrane-binding proteins: Cyt b5 (a) and PV proteins 3AB (b), 2C (c), 2BC (d), 2C(K135S) (e), and 2C(1-274) (f). Arrowheads indicate the characteristic alterations for each protein: concentric, myelin-like membranes (a and b); vesicles surrounding lipid droplets (b); rigid, extended membrane sheets (c); and vesicles similar to those found during a PV infection (d to f). The micrographs were obtained 9 to 14 h posttransfection. Bars, 500 nm.

Cells with membranes altered by membrane-binding proteins show reduced permissiveness for PV replication. To test whether PV can replicate in cells containing membranes altered by the expression of viral or cellular proteins, HeLa cell monolayers were transfected with DNA encoding 2C, 2C(K135S), 2C(1-274), 2BC, 3AB, 3AB DII 3E, or Cyt b5 to express the corresponding protein. All proteins carried the Flag epitope at their N termini (Fig. 1). The cultures were infected with PV 6 to 7 h after transfection and fixed for IF analysis after another 5 h. The number of cells expressing one of the above proteins and supporting PV infection was determined by IF. Cells expressing a membrane-binding protein were identified by IF with a mouse MAb detecting the Flag epitope. PV-infected cells were monitored simultaneously in the same preparation by measurement of their content either of 2C (after P3 protein expression) or of VPg (after P2 protein or Cyt b5 expression), using IF with polyclonal rabbit antisera (Table 1). This is visualized in Fig. 3B for cells transfected with 2BC. The percentage of cells expressing a membrane-binding protein varied between 20 and 90%, depending on the plasmid used for induction of membrane alterations. The time interval between transfection and superinfection with PV was optimized. At 6 to 7 h after transfection, the expressed proteins and the corresponding membrane alterations became detectable. Shorter intervals led to little or no membrane alterations due to a low expression of membrane-binding proteins as a consequence of PV-induced shutoff of vaccinia virus-mediated T7 RNA polymerase production. Longer times resulted in a more extensive inhibition of PV replication (see below), rendering the in situ analysis difficult. Figure 3A shows that the percentage of cells permissive for PV was 95 to 98% in vaccinia virus-infected, mock-transfected control cultures. In contrast, the cells productively transfected with plasmids encoding membrane-binding proteins showed a reduced susceptibility to PV. The percentage of PV-susceptible cells in the expressing subpopulation varied between 12 and 55% (Fig. 3A).

                              
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  		TABLE 1.   Combinations of Ab and RNA probes for in situ detection experiments



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  		FIG. 3.   Expression of membrane-binding proteins reduces the susceptibility of cells to a PV infection. (A) Cells were transfected with one of the constructs indicated and superinfected with PV 6 to 7 h later. At 5 h p.i., the cells were subjected to IF with MAb against the Flag epitope of the transfected protein and with an Ab detecting PV (see Table 1). PV-infected cells are indicated as the percentage of the subpopulation of cells which express the indicated protein. (B) The upper and lower panels show the identical area of micrographs of cells transfected with pTM-Fg-2BC and superinfected with PV. (Top) IF performed with MAb against Flag to detect Fg-2BC. (Bottom) IF performed with anti-VPg Ab to detect PV. Most cells productively infected with PV were not expressing 2BC. Solid arrowheads indicate cells infected with PV and not expressing 2BC. Open arrowheads indicate cells expressing 2BC and not infected with PV. The asterisk indicates a cell expressing 2BC and infected with PV. Magnification, ×320.

As judged by fluorescence intensity, cells exhibiting both expression of a membrane-binding protein and infection with PV show a distinctly lower level of PV proteins compared with that of cells positive for PV only. This points to a direct effect of the expression of membrane-binding proteins on PV replication and not to a nonspecific inhibition of PV due to the transfection procedure per se. This is further substantiated by the observation that expression of the PV protein 3AB DII 3E, which does not bind to membranes, inhibits PV replication only in approximately 5% of the transfected cells (Fig. 3A). Thus, the mutated, soluble protein 3AB DII 3E did not interfere with PV replication whereas the membrane-binding proteins reduced susceptibility to PV by 40 to 90%.

It should be noted that the reduced susceptibility of cells expressing membrane-binding proteins to PV infection was detected only using in situ methods. Previous biochemical analyses of RNA accumulation performed on whole-cell populations did not demonstrate significant differences in PV infection permissiveness in cell populations transfected with plasmids encoding protein 2BC (60). To resolve this apparent discrepancy, we performed experiments quantitating PV replication by slot blot hybridization of progeny viral RNA in parallel with in situ experiments. In cultures exhibiting a 60% transfection efficiency, the number of cells supporting PV replication was typically around 55%, i.e., 40% nontransfected and thus PV-susceptible cells plus 15% transfected and PV-susceptible cells (compare with Fig. 3). Thus, more than half of the amount of viral RNA is likely to be produced in such a culture compared to a nontransfected PV-infected culture. This reduction could not be found reproducibly by the slot blot method employed.

To test whether the reduced susceptibility of transfected cells was not due to a loss of the PV receptor as a consequence of the expression of membrane-binding proteins, the presence of the receptor was monitored by IF with anti-CD 155 Ab on the surface of living, nonpermeabilized cells. 2BC-transfected HeLa cells clearly were positive for the receptor (data not shown). Untreated HeLa cells and mutagenized HT1080 fibrosarcoma cells were used as positive and negative controls, respectively.

Membranes altered by membrane-binding proteins are not converted to PV-specific vesicles. To determine whether the altered membranes became converted to PV-specific vesicles in cells in which PV infection was not inhibited, the population of cells expressing the membrane-binding protein and still allowing PV replication was analyzed for colocalization of transfection-derived membrane-binding protein and infection-derived protein 2C or VPg. Note that both 2C and VPg can be used as markers for virus-induced vesicles. This was established in PV-infected cells, where the vesicle-associated P2 protein sequences 2C and 2B (11, 27) were found by IF to be associated with structures identical to those associated with VPg (data not shown).

Figure 4 shows that the expressed membrane-binding proteins, whether of PV or cellular origin, did not colocalize in vesicles with 2C or VPg protein produced after infection. This was found regardless of whether the transfection-induced altered membranes exhibited structures not found during PV infection, such as those observed after 2C (18) (data not shown) or Cyt b5 (Fig. 4a to c) expression, or whether the altered membranes consisted of 2C(1-274)- or 2BC-induced ER-derived vesicles that resemble those in PV-infected cells (Fig. 4d to i).


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  		FIG. 4.   Localization of membrane-binding proteins and viral products in transfected, infected cells. HeLa cells expressing membrane-binding proteins Fg-Cyt b5 (a to c), Fg-PV 2C(1-274) (d to f), and Fg-PV 2BC (g to m) were superinfected with PV at 6 to 7 h posttransfection. Cells that were both transfected and infected were selected for analysis by CLSM. (a, d, g, and k) IF with anti-Flag MAb and Texas red-labeled secondary Ab to detect the expressed membrane-binding proteins. (b, e, and h) PV replication complex visualized at 5 h p.i. with anti-VPg Ab and FITC-coupled secondary Ab. (l) PV plus-strand RNA detected by ISH with FITC-labeled riboprobe. (c, f, i, and m) Overlay of the corresponding individual reactions: PV replication complex-associated proteins and RNA stay separate from membrane-binding proteins. Each image area is 28 by 35 µm.

To show that the PV-induced vesicular clusters contain viral RNA and thus represent replication complexes (11), FISH was performed to detect PV plus-strand RNA. Simultaneously, the expressed protein 2BC was detected by IF with anti-Flag MAb. Figure 4k to m show that the viral RNA does not colocalize to the preexisting 2BC-induced vesicles.

The findings indicate that PV infection does not convert transfection-induced nonvesiculated smooth membranes, although ER derived, into virus-induced vesicles. Furthermore, the vesicular rosettes harboring the viral replication complex during PV replication (15) do not incorporate altered membranes, even if the membranes are vesicles closely resembling those found in a PV replication complex.

Vesicles induced by expression of all viral nonstructural proteins are not used to form replication complexes. In the above experiments, vesicles induced by protein 2BC and, consequently, carrying only this viral protein were shown not to contribute to viral replication complexes formed after subsequent virus infection. To produce vesicles which might contain a complete set of viral replication proteins, cells were transfected with a plasmid, pE5PVDelta P1, encoding the entire PV P2 and P3 regions, fused to the EMCV IRES (Fig. 1), downstream of a T7 promoter. This plasmid encodes all of the viral proteins required for viral RNA replication and induces the formation of vesicles closely resembling in size and array those found in PV-infected cells. However, the transcripts do not serve as templates for RNA replication because they lack the PV 5' terminal RNA sequences and structures required for template recognition by the replication complex (unpublished data). For comparison, a control plasmid, pPVDelta P1, which encodes the same P2 and P3 regions fused to the authentic PV 5' untranslated region, was constructed. pPVDelta P1 produces transcripts which can be replicated by RNA-dependent RNA synthesis, whereas pE5PVDelta P1 produces transcripts only by T7-mediated DNA-dependent RNA synthesis, and these transcripts are unable to replicate on their own. Since the association of P2 proteins with viral RNA and virus-induced membranes is indicative of the formation of a PV replication complex, we tested whether the biochemical differences in RNA synthesis properties can be visualized by an in situ analysis demonstrating the intracellular location of plasmid-specific protein and RNA. pE5PVDelta P1-transfected cells accumulate viral protein and plus-strand RNA clearly in distinct, nonoverlapping regions (Fig. 5a to c). pPVDelta P1 induces the formation of structures in which protein and RNA largely colocalize and which resemble virus-induced replication complexes early in infection (16) (Fig. 5d to f). Some free plus-strand RNA, thought to consist of RNA produced by T7-mediated DNA-dependent RNA synthesis, can also be seen.


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  		FIG. 5.   Localization of viral proteins and RNA in transfected HeLa cells. (a to f) Cells transfected with pE5PVDelta P1 (a to c) or the replicon pPVDelta P1 (d to f) were treated with actinomycin D at 2.5 h posttransfection to stop T7-mediated transcription. 2B and 2BC were visualized by IF and CLSM with anti-2B MAb and Texas red-labeled secondary Ab (a and d). RNA of the transfected construct was localized by ISH with FITC-labeled riboprobe (b and e) at 7 h posttransfection. (c and f) Overlay. Replication complexes, similar to those found in a PV-infected cell, can be formed only with pPVDelta P1-derived replicating RNA (f). (g to i) Cells dually transfected with pTM-Fg-2BC for 7 h and supertransfected with pE5PVDelta P1 for another 7 h. 2BC-induced vesicles were visualized by IF with anti-Flag MAb and Texas red-labeled secondary Ab (g), and pE5PVDelta P1-induced vesicles were visualized with anti-VPg Ab and FITC-labeled secondary Ab (h). (i) Overlay. 2BC- and pE5PVDelta P1-induced vesicles, not engaged in RNA-dependent RNA synthesis, mix and associate with each other. Each image area is 26 by 33 µm.

To determine whether the vesicles which are induced by expression of all of the viral nonstructural proteins and which are not associated with viral RNA could be used subsequently in the formation of replication complexes by superinfecting virus, HeLa cells were transfected with pE5PVDelta P1 and 6.3 h later infected with PV. PV replication was monitored by IF at 5 h postinfection (p.i.) using MAb against capsid protein VP1, a protein from the genomic region deleted in pE5PVDelta P1 (Fig. 1). In a mock-transfected, vaccinia virus-infected culture, which was PV infected 6.3 h after the mock transfection, 90% of the cells tested positive for PV VP1 at 5 h p.i. In a pE5PVDelta P1-transfected cell culture that was not PV infected, 95% of the cells were found positive for pE5PVDelta P1 as measured by IF with MAb against protein 2B (Fig. 6A and B, upper panel). In a parallel experiment with pE5PVDelta P1-transfected cells which were superinfected with PV at 6.3 h posttransfection, the percentage of PV-permissive cells was reduced by 56%, suggesting that the preexisting vesicles were not readily used in the formation of replication complexes with infecting virus (Fig. 6B, lower panel).


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  		FIG. 6.   HeLa cells were transfected with pE5PVDelta P1 and superinfected with PV at 6.3 h posttransfection. (A) The number of PV-susceptible cells was found to be reduced compared to a mock-transfected culture. Since pE5PVDelta P1 proteins cannot be discriminated from PV proteins, the conclusion that pE5PVDelta P1 inhibits PV replication was drawn from counting infected or transfected cells in parallel cultures which were mock transfected and PV infected, transfected with pE5PVDelta P1 only, or transfected with pE5PVDelta P1 and PV superinfected. n.a., not applicable. (B) The upper picture shows cells transfected with pE5PVDelta P1. IF performed with anti-2B MAb showed that 95% of the cells were efficiently transfected. The lower picture shows a parallel culture transfected with pE5PVDelta P1 and superinfected with PV. The number of PV-infected cells was determined by IF with anti-VP1 MAb at 5 h p.i. Magnification, ×100.

These results argue that vesicles induced in the presence of all of the viral replication proteins are not recruited to support RNA synthesis of superinfecting PV and apparently are not readily formed into new replication complex vesicles. The vesicle clusters surrounding viral replication complexes appear to be assembled from cellular membrane structures by a mechanism coupled to viral RNA synthesis. This implies that preformed modified vesicles do not assemble with viral RNA to form a functional replication complex.

However, in cells transfected with 2BC and supertransfected with pE5PVDelta P1, the discrimination between vesicle populations is not observed. Figure 5g to i show that pE5PVDelta P1-induced vesicles, which are not associated with RNA or engaged in RNA synthesis, can readily mix and associate with 2BC-induced preexisting vesicles [compare to the complementary experiment in Fig. 4d to i, where PV-induced vesicles in a replication complex did not colocalize to preformed 2BC- or 2C(1-274)-induced vesicles].

Together, these findings indicate that RNA synthesis is necessary for the formation of the replication complex and that the resulting replication complex does not readily exchange components with other membranous structures.

Translation of viral proteins is reduced in cells with previously altered membranes. RNA synthesis in PV-infected cells is dependent on translation of viral RNA, so that sufficient amounts of viral proteins necessary for RNA replication can be created. In pE5PVDelta P1-transfected cells, however, RNA synthesis is mediated by T7 RNA polymerase and thus is independent of translation of plasmid-derived RNA. Therefore, pE5PVDelta P1 can be used to study modifications in translational or transcriptional activity separately.

To test whether induction of membranes altered by the expression of membrane-binding protein exerts an influence on translation of viral RNA, cell cultures were transfected with 2BC and then supertransfected with pE5PVDelta P1. By combined IF and FISH, we determined the number of 2BC-expressing cells that transcribed pE5PVDelta P1 RNA and expressed the corresponding proteins. For comparison, parallel cultures of 2BC-transfected cells were superinfected with PV, where viral RNA synthesis can occur only in cells that synthesize protein. As expected, in 2BC-transfected cells superinfected with PV, all of the viral RNA-replicating cells also synthesized viral protein (data not shown). In 2BC- and pE5PVDelta P1-transfected cells, however, 52% of the 2BC-positive cells produced pE5PVDelta P1 RNA but only 16% synthesized protein from that RNA (Table 2). These data suggest that translation of viral RNA is inhibited in cells whose internal membranes have been rearranged to form the 2BC-induced vesicles.

                              
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  		TABLE 2.   Expression of a membrane-binding protein inhibits translation of pE5PVDelta P1 RNAa

To ensure that the inhibition of translation was not due to competition for components needed to utilize the EMCV IRES, which was present in both the 2BC and pE5PVDelta P1 constructs, control cultures were transfected with constructs expressing proteins that do not bind to and modify membranes but are still translated from an EMCV IRES. The control plasmids encoded either the 3AB DII 3E mutant or beta -globin. In both cases, approximately the same percentage of productively transfected cells subsequently produced both pE5PVDelta P1 RNA and protein (Table 2).


    DISCUSSION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Modified intracellular membranes are not transformed into replication complexes. Infection of cells with PV induces an extensive rearrangement of intracellular membranes into characteristic vesicles which assemble into a higher-order rosette structure (12, 15) surrounding and sequestering the site of RNA synthesis. It is not known what characteristics qualify a membrane for integration into replication complexes. In this study, we altered ER membranes in HeLa cells by expressing different cellular or viral membrane-binding proteins. Although the morphologies of the altered membrane structures varied depending on the protein used to induce the alterations, some of the membrane alterations induced by membrane-binding proteins, particularly by PV protein 2BC or the entire P2-P3 region, produced vesicles similar to those induced during a PV infection. Such vesicles were associated with one or multiple PV proteins. The altered membranes, which were immunocytochemically traceable, were challenged for their utilization during a subsequent PV infection.

Unexpectedly, none of the altered ER membranes contributed to PV-induced replication complexes. If the altered membranes were myelin-like threads and whirls (Fig. 2), they were not transformed into PV-induced vesicles. If the membrane alterations produced vesicles, such as those induced by PV protein 2BC or 2C(1-274), they were not incorporated into rosettes of replication complexes built up during a PV infection.

Cells with altered intracellular membranes do not support PV replication. The expression of membrane-binding proteins led to a marked reduction of permissiveness of the cells for PV replication. No loss of PV receptor CD 155 was detected due to the expression of membrane-altering proteins.

Other studies have reported that a functional Golgi complex might be essential for PV replication (54). Since long-term expression (12 to 14 h) of some of the membrane-binding proteins induces disruption of the Golgi stacks (59), we considered whether the inhibition of PV replication in cells expressing membrane-binding proteins might be due to an absence of Golgi complexes in the transfected cells. However, disintegration of the Golgi complex is not likely to cause failure to support virus growth, since IF analysis with a Golgi-specific MAb showed that in 85 to 90% of the cells at the time of superinfection with PV (6 to 7 h after transfection), the Golgi is morphologically intact (data not shown).

The reduction of PV replication in cells expressing a membrane-binding protein might be due to a reduced translational activity of these cells, as shown in Table 2. The alteration in intracellular membrane morphology induced by expression of membrane-binding proteins (Fig. 2) might interfere with rER-associated protein synthesis by reducing the amount of available ER. This reduced translational activity might inhibit PV replication if PV RNA were translated on membrane-bound ribosomes (52, 53). The site of PV translation is currently being investigated.

The amount of vesiculated membranes available to form a replication complex for viral transcription might become limiting in cells expressing membrane-binding proteins. This shortage of utilizable membranes seems more likely to occur in cells expressing proteins, such as Cyt b5, wt 2C, or 3AB, which induce membranes that are not suitable for transformation into vesicles by superinfecting PV. However, in cells expressing proteins, such as 2BC, 2C(1-274) or pE5PVDelta P1, which induce vesicles, PV replication is still largely suppressed. This indicates that preformed vesicles, even if they carry the entire set of P2 and P3 proteins, cannot be used to form replication complexes and therefore cannot compensate for a possible limiting amount of PV-induced vesicles.

Formation of a replication complex requires replicating RNA. Simultaneous intracellular localization of preformed vesicles and the replication complexes containing replicating RNA of superinfecting PV demonstrated that preformed vesicles were excluded from PV replication complexes, and therefore they were not used for replication of the superinfecting virus (Fig. 4). In contrast, 2BC-induced preformed vesicles associated freely with vesicles induced by supertransfection of pE5PVDelta P1, whose RNA transcript was not competent to undergo replication (Fig. 5). This suggests a role for RNA synthesis in forming and maintaining a replication complex.

The influence of RNA synthesis on replication complex formation was also demonstrated with the pE5PVDelta P1 construct. pE5PVDelta P1 is transcribed by T7 RNA polymerase and expresses proteins that induce vesicles but no detectable membrane-bound replication complexes (Fig. 5). Surprisingly, the vesicles induced by pE5PVDelta P1 remained clearly separated from pE5PVDelta P1 RNA (Fig. 5). The striking separation of RNA and vesicles formed by proteins translated from the nonreplicating RNA results from the absence of RNA replication and not from the absence of PV-specific signals or sequences in the EMCV 5' nontranslated region (NTR). This is inferred from similar observations made on cells transfected with a nonreplicating pPVDelta P1 construct that has an authentic PV 5' NTR and a replication defect due to a mutation in the 3Dpol region (unpublished data).

The view that the formation and integrity of a replication complex depend on viral RNA synthesis is supported by the in vitro finding that an isolated vesicular rosette surrounding a functional replication complex can be dissociated into single vesicles in the cold, under conditions where RNA synthesis is stopped. After raising the temperature and allowing resumption of RNA synthesis, the intact rosette is re-formed (27).

Formation of a replication complex requires viral proteins, vesicle formation, and RNA replication in cis. We interpret our data to mean that during the onset of the rapid replication of PV as occurs in a living cell, translation of viral RNA into proteins, induction of vesicles, and their association into a functional replication complex are coupled processes. This is in agreement with reports that viral RNA must first be translated in order to replicate (43) and also with the suggestion that all components of the replication complex are delivered en bloc directly following translation (19, 64). Furthermore, it was shown that PV replication occurs only after uninterrupted translation of the P2 and P3 coding sequences (46). Our observations indicate that preformed vesicles carrying only one PV protein (2BC) cannot accept RNA and other PV replication proteins, nor can vesicles carrying all P2 and P3 replication proteins (pE5PVDelta P1) accept RNA and become organized into a replication complex. Thus, we propose that vesicles to be utilized in a replication complex require that they be formed and provided with the relevant proteins and RNA in cis.

Mutations in most of the noncapsid proteins are unable to be complemented in trans, or if they are, they represent only certain functions of a multifunctional protein (60, 61, 68). This may result from the formation of the replication complex in cis and from its compact architecture, which sequesters its components and prevents their physical association or exchange with complementing counterparts. In contrast, genetic recombination, which occurs by strand switching during minus-strand synthesis (38), is reported to take place with rather high frequency (reviewed in references 1, 37, and 68). The observations presented here and summarized in Fig. 7 indicate that there is little exchange of components between different replication complexes. Taken together with the finding that the replication complex is rather tightly sealed (12, 28), the high frequency of recombination events appears difficult to reconcile.


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  		FIG. 7.   Schematic diagram illustrating that the PV replication complex is formed in cis. (A) ER membranes are altered by the expression of membrane-binding proteins, e.g., Cyt b5 or PV protein 2C, or all of the PV nonstructural proteins encoded in the plasmid pE5PVDelta P1. The altered membranes are not utilized (crossed arrows) in trans for PV replication complex formation during a subsequent PV infection. (B) Replication complex formation as it occurs during PV replication. Membrane-bound translation of viral RNA into protein (step 1) triggers the formation of vesicles on the ER (step 2). Vesicles carrying PV nonstructural proteins and previously translated RNA with initiated minus strand (step 3) form a viral replication complex (vesicular rosette) with replicating RNA in the replicative intermediate (RI) configuration (step 4). Hatched lines, RNA; stippled symbols, viral proteins.

We propose the following order of events during early steps of PV RNA replication. After translation of viral plus-strand RNA, presumably on the rER, viral proteins induce vesicles and, concomitantly, initiate transcription of the RNA into a minus strand (Fig. 7, steps 1 to 3). Several ER-derived vesicular clusters, each emerging from one translated RNA, are thought to combine in spherical structures, previously found by FISH to contain plus- and minus-strand RNA (16). These structures are considered to be the site of continued minus-strand RNA synthesis, thereby enabling recombination. After completion of minus-strand RNA, multiple initiations of plus-strand RNA create the replicative intermediate contained in membrane-bound replication complexes (Fig. 7, step 4) which eventually aggregate into the large juxtanuclear area of vesicles, characteristic of PV-infected cells.


    ACKNOWLEDGMENTS

This work was supported by grant 31-055397.98 from the Swiss National Science Foundation and grant 10348 from INTAS-RFBR and by the U.S. National Institutes of Health.

We thank B. L. Semler and J. S. Towner, University of California, Irvine, Calif., for plasmids pTM-Fg-3AB, pTM-Fg-3AB DII 3E, pTM- Fg-beta -globin, and pTM-Fg-Cyt b5; V. Boyko for introducing the Flag sequence into some of the constructs; L. Pasamontes, Roche, for providing the anti-VPg antibody; and A. Wyss for supplying the mutagenized HT1080 fibrosarcoma cells. We are grateful to L. Landmann, Department of Anatomy, University of Basel, for his help and hospitality at the CLSM unit. A. Glaser-Ruhm provided excellent technical assistance, and laboratory members helped with stimulating discussions.


    FOOTNOTES

* Corresponding author. Mailing address: Institute for Medical Microbiology, University of Basel, Petersplatz 10, CH-4003 Basel, Switzerland. Phone: 41 61 267 3290. Fax: 41 61 267 3298. E-mail: Kurt.Bienz{at}unibas.ch.


    REFERENCES
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

1. Agol, V. I. 1997. Recombination and other genomic rearrangements in picornaviruses. Semin. Virol. 8:1-9.
2. Agol, V. I., G. A. Belov, K. Bienz, D. Egger, M. S. Kolesnikova, N. T. Raikhlin, L. I. Romanova, E. A. Smirnova, and E. A. Tolskaya. 1998. Two types of death of poliovirus-infected cells: caspase involvement in the apoptosis but not cytopathic effect. Virology 252:343-353[CrossRef][Medline].
3. Aldabe, R., A. Barco, and L. Carrasco. 1996. Membrane permeabilization by poliovirus proteins 2B and 2BC. J. Biol. Chem. 271:23134-23137[Abstract/Free Full Text].
4. Aldabe, R., and L. Carrasco. 1995. Induction of membrane proliferation by poliovirus proteins 2C and 2BC. Biochem. Biophys. Res. Commun. 206:64-76[CrossRef][Medline].
5. Aldabe, R., A. Irurzun, and L. Carrasco. 1997. Poliovirus protein 2BC increases cytosolic free calcium concentrations. J. Virol. 71:6214-6217[Abstract].
6. Andino, R., G. E. Rieckhof, P. L. Achacoso, and D. Baltimore. 1993. Poliovirus RNA synthesis utilizes an RNP complex formed around the 5'-end of viral RNA. EMBO J. 12:3587-3598[Medline].
7. Andino, R., G. E. Rieckhof, and D. Baltimore. 1990. A functional ribonucleoprotein complex forms around the 5' end of poliovirus RNA. Cell 63:369-380[CrossRef][Medline].
8. Bablanian, R. 1972. Depression of macromolecular synthesis in cells infected with guanidine-dependent poliovirus under restrictive conditions. Virology 47:255-259[CrossRef][Medline].
9. Barco, A., and L. Carrasco. 1995. A human virus protein, poliovirus protein 2BC, induces membrane proliferation and blocks the exocytic pathway in the yeast Saccharomyces cerevisiae. EMBO J. 14:3349-3364[Medline].
10. Bell, Y. C., B. L. Semler, and E. Ehrenfeld. 1999. Requirements for RNA replication of a poliovirus replicon by coxsackievirus B3 RNA polymerase. J. Virol. 73:9413-9421[Abstract/Free Full Text].
11. Bienz, K., D. Egger, and L. Pasamontes. 1987. Association of polioviral proteins of the P2 genomic region with the viral replication complex and virus induced membrane synthesis as visualized by electron microscopic immunocytochemistry and autoradiography. Virology 160:220-226[CrossRef][Medline].
12. Bienz, K., D. Egger, T. Pfister, and M. Troxler. 1992. Structural and functional characterization of the poliovirus replication complex. J. Virol. 66:2740-2747[Abstract/Free Full Text].
13. Bienz, K., D. Egger, Y. Rasser, and W. Bossart. 1983. Intracellular distribution of poliovirus proteins and the induction of virus specific cytoplasmic structures. Virology 131:39-48[CrossRef][Medline].
14. Bienz, K., D. Egger, Y. Rasser, and W. Bossart. 1980. Kinetics and location of poliovirus macromolecular synthesis in correlation to virus induced cytopathology. Virology 100:390-399[CrossRef][Medline].
15. Bienz, K., D. Egger, M. Troxler, and L. Pasamontes. 1990. Structural organization of poliovirus RNA replication is mediated by viral proteins of the P2 genomic region. J. Virol. 64:1156-1163[Abstract/Free Full Text].
16. Bolten, R., D. Egger, R. Gosert, G. Schaub, L. Landmann, and K. Bienz. 1998. Intracellular localization of poliovirus plus- and minus-strand RNA visualized by strand-specific fluorescent in situ hybridization. J. Virol. 72:8578-8585[Abstract/Free Full Text].
17. Caliguiri, L. A., and I. Tamm. 1970. The role of cytoplasmic membranes in poliovirus biosynthesis. Virology 42:100-111[CrossRef][Medline].
18. Cho, M. W., N. Teterina, D. Egger, K. Bienz, and E. Ehrenfeld. 1994. Membrane rearrangement and vesicle induction by recombinant poliovirus 2C and 2BC in human cells. Virology 202:129-145[CrossRef][Medline].
19. Collis, P. S., B. J. O'Donnell, D. J. Barton, J. A. Rogers, and J. B. Flanegan. 1992. Replication of poliovirus RNA and subgenomic RNA transcripts in transfected cells. J. Virol. 66:6480-6488[Abstract/Free Full Text].
20. Cuconati, A., A. Molla, and E. Wimmer. 1998. Brefeldin A inhibits cell-free, de novo synthesis of poliovirus. J. Virol. 72:6456-6464[Abstract/Free Full Text].
21. Datta, U., and A. Dasgupta. 1994. Expression and subcellular localization of poliovirus VPg-precursor protein 3AB in eukaryotic cells: evidence for glycosylation in vitro. J. Virol. 68:4468-4477[Abstract/Free Full Text].
22. Doedens, J. R., T. H. Giddings, and K. Kirkegaard. 1997. Inhibition of endoplasmic reticulum-to-Golgi traffic by poliovirus protein 3A: genetic and ultrastructural analysis. J. Virol. 71:9054-9064[Abstract].
23. Doedens, J. R., and K. Kirkegaard. 1995. Inhibition of cellular protein secretion by poliovirus proteins 2B and 3A. EMBO J. 14:894-907[Medline].
24. Echeverri, A., R. Banerjee, and A. Dasgupta. 1998. Amino-terminal region of poliovirus 2C protein is sufficient for membrane binding. Virus Res. 54:217-223[CrossRef][Medline].
25. Echeverri, A. C., and A. Dasgupta. 1995. Amino terminal regions of poliovirus 2C protein mediate membrane binding. Virology 208:540-553[CrossRef][Medline].
26. Egger, D., R. Bolten, C. Rahner, and K. Bienz. 1999. Fluorochrome-labeled RNA as a sensitive, strand-specific probe for direct fluorescence in situ hybridization. Histochem. Cell Biol. 111:319-324[CrossRef][Medline].
27. Egger, D., L. Pasamontes, R. Bolten, V. Boyko, and K. Bienz. 1996. Reversible dissociation of the poliovirus replication complex: functions and interactions of its components in viral RNA synthesis. J. Virol. 70:8675-8683[Abstract].
28. Etchison, D., and E. Ehrenfeld. 1981. Comparison of replication complexes synthesizing poliovirus RNA. Virology 111:33-46[CrossRef][Medline].
29. Fuerst, T. R., E. G. Niles, F. W. Studier, and B. Moss. 1986. Eukaryotic transient-expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. USA 83:8122-8126[Abstract/Free Full Text].
30. Gamarnik, A. V., and R. Andino. 1997. Two functional complexes formed by KH domain containing proteins with the 5' noncoding region of poliovirus RNA. RNA 3:882-892[Abstract].
31. Girard, S., T. Couderc, J. Destombes, D. Thiesson, F. Delpeyroux, and B. Blondel. 1999. Poliovirus induces apoptosis in the mouse central nervous system. J. Virol. 73:6066-6072[Abstract/Free Full Text].
32. Grun, J. B., and M. A. Brinton. 1988. Separation of functional West Nile virus replication complexes from intracellular membrane fragments. J. Gen. Virol. 69:3121-3127[Abstract/Free Full Text].
33. Irurzun, A., J. Arroyo, A. Alvarez, and L. Carrasco. 1995. Enhanced intracellular calcium concentration during poliovirus infection. J. Virol. 69:5142-5146[Abstract].
34. Irurzun, A., L. Perez, and L. Carrasco. 1992. Involvement of membrane traffic in the replication of poliovirus genomes: effects of brefeldin A. Virology 191:166-175[CrossRef][Medline].
35. Jia, X.-Y., M. Van Eden, M. G. Busch, E. Ehrenfeld, and D. F. Summers. 1998. trans-Encapsidation of a poliovirus replicon by different picornavirus capsid proteins. J. Virol. 72:7972-7977[Abstract/Free Full Text].
36. Jore, J., B. De Geus, R. J. Jackson, P. H. Pouwels, and B. E. Enger-Valk. 1988. Poliovirus protein 3CD is the active protease for processing of the precursor protein P1 in vitro. J. Gen. Virol. 69:1627-1636[Abstract/Free Full Text].
37. King, A. M. Q. 1988. Genetic recombination in positive strand RNA viruses, p. 149-165. In E. Domingo, J. J. Holland, and P. Ahlquist (ed.), RNA genetics, vol. 2. CRC Press, Inc., Boca Raton, Fla.
38. Kirkegaard, K., and D. Baltimore. 1986. The mechanism of RNA recombination in poliovirus. Cell 47:433-443[CrossRef][Medline].
39. Levine, R. A., and D. A. Wolff. 1979. Bovine enterovirus CPE at different multiplicities of infection in the absence of viral RNA synthesis. Intervirology 11:255-260[Medline].
40. Longin, A., C. Souchier, M. Ffrench, and P. A. Bryon. 1993. Comparison of anti-fading agents used in fluorescence microscopy: image analysis and laser confocal microscopy study. J. Histochem. Cytochem. 41:1833-1840[Abstract].
41. Mackenzie, J. M., M. K. Jones, and E. G. Westaway. 1999. Markers for trans-Golgi membranes and the intermediate compartment localize to induced membranes with distinct replication functions in flavivirus-infected cells. J. Virol. 73:9555-9567[Abstract/Free Full Text].
42. Maynell, L. A., K. Kirkegaard, and M. W. Klymkowsky. 1992. Inhibition of poliovirus RNA synthesis by brefeldin A. J. Virol. 66:1985-1994[Abstract/Free Full Text].
43. Novak, J. E., and K. Kirkegaard. 1994. Coupling between genome translation and replication in an RNA virus. Genes Dev. 8:1726-1737[Abstract/Free Full Text].
44. Nugent, C. I., K. L. Johnson, P. Sarnow, and K. Kirkegaard. 1999. Functional coupling between replication and packaging of poliovirus replicon RNA. J. Virol. 73:427-435[Abstract/Free Full Text].
45. Parsley, T. B., J. S. Towner, L. B. Blyn, E. Ehrenfeld, and B. L. Semler. 1997. Poly(rC) binding protein 2 forms a ternary complex with the 5'-terminal sequences of poliovirus RNA and the viral 3CD proteinase. RNA 3:1124-1134[Abstract].
46. Paul, A. V., J. Mugavero, A. Molla, and E. Wimmer. 1998. Internal ribosomal entry site scanning of the poliovirus polyprotein: implications for proteolytic processing. Virology 250:241-253[CrossRef][Medline].
47. Pedersen, K. W., Y. van der Meer, N. Roos, and E. J. Snijder. 1999. Open reading frame 1a-encoded subunits of the arterivirus replicase induce endoplasmic reticulum-derived double-membrane vesicles which carry the viral replication complex. J. Virol. 73:2016-2026[Abstract/Free Full Text].
48. Pfister, T., D. Egger, and K. Bienz. 1995. Poliovirus subviral particles associated with progeny RNA in the replication complex. J. Gen. Virol. 76:63-71[Abstract/Free Full Text].
49. Pfister, T., and E. Wimmer. 1999. Characterization of the nucleoside triphosphatase activity of poliovirus protein 2C reveals a mechanism by which guanidine inhibits poliovirus replication. J. Biol. Chem. 274:6992-7001[Abstract/Free Full Text].
50. Restrepo-Hartwig, M. A., and P. Ahlquist. 1996. Brome mosaic virus helicase- and polymerase-like proteins colocalize on the endoplasmic reticulum at sites of viral RNA synthesis. J. Virol. 70:8908-8916[Abstract].
51. Roth, M. G. 1999. Snapshots of ARF1: implications for mechanisms of activation and inactivation. Cell 97:149-152[CrossRef][Medline].
52. Roumiantzeff, M., J. V. Maizel, Jr., and D. F. Summers. 1971. Comparison of polysomal structures of uninfected and poliovirus infected HeLa cells. Virology 44:239-248[CrossRef][Medline].
53. Roumiantzeff, M., D. F. Summers, and J. V. Maizel, Jr. 1971. In vitro protein synthetic activity of membrane-bound poliovirus polyribosomes. Virology 44:249-258[CrossRef][Medline].
54. Sandoval, I. V., and L. Carrasco. 1997. Poliovirus infection and expression of the poliovirus protein 2B provoke the disassembly of the Golgi complex, the organelle target for the antipoliovirus drug Ro-090179. J. Virol. 71:4679-4693[Abstract].
55. Schaad, M. C., P. E. Jensen, and J. C. Carrington. 1997. Formation of plant RNA virus replication complexes on membranes: role of an endoplasmic reticulum-targeted viral protein. EMBO J. 16:4049-4059[CrossRef][Medline].
56. Schlegel, A., T. H. Giddings, Jr., M. S. Ladinsky, and K. Kirkegaard. 1996. Cellular origin and ultrastructure of membranes induced during poliovirus infection. J. Virol. 70:6576-6588[Abstract/Free Full Text].
57. Springer, S., A. Spang, and R. Schekman. 1999. A primer on vesicle budding. Cell 97:145-148[CrossRef][Medline].
58. Takegami, T., B. L. Semler, C. W. Anderson, and E. Wimmer. 1983. Membrane fractions active in poliovirus RNA replication contain VPg precursor polypeptides. Virology 128:33-47[CrossRef][Medline].
59. Teterina, N. L., A. E. Gorbalenya, D. Egger, K. Bienz, and E. Ehrenfeld. 1997. Poliovirus 2C protein determinants of membrane binding and rearrangements in mammalian cells. J. Virol. 71:8962-8972[Abstract].
60. Teterina, N. L., W. D. Zhou, M. W. Cho, and E. Ehrenfeld. 1995. Inefficient complementation activity of poliovirus 2C and 3D proteins for rescue of lethal mutations. J. Virol. 69:4245-4254[Abstract].
61. Tolskaya, E. A., L. I. Romanova, M. S. Kolesnikova, A. P. Gmyl, A. E. Gorbalenya, and V. I. Agol. 1994. Genetic studies on the poliovirus 2C protein, an NTPase. A plausible mechanism of guanidine effect on the 2C function and evidence for the importance of 2C oligomerization. J. Mol. Biol. 236:1310-1323[CrossRef][Medline].
62. Tolskaya, E. A., L. I. Romanova, M. S. Kolesnikova, T. A. Ivannikova, E. A. Smirnova, N. T. Raikhlin, and V. I. Agol. 1995. Apoptosis-inducing and apoptosis-preventing functions of poliovirus. J. Virol. 69:1181-1189[Abstract].
63. Towner, J. S., T. V. Ho, and B. L. Semler. 1996. Determinants of membrane association for poliovirus protein 3AB. J. Biol. Chem. 271:26810-26818[Abstract/Free Full Text].
64. Towner, J. S., M. M. Mazanet, and B. L. Semler. 1998. Rescue of defective poliovirus RNA replication by 3AB-containing precursor polyproteins. J. Virol. 72:7191-7200[Abstract/Free Full Text].
65. Valnes, K., and P. Brandtzaeg. 1985. Retardation of immunofluorescence fading during microscopy. J. Histochem. Cytochem. 33:755-761[Abstract].
66. Van Kuppeveld, F. J., W. J. Melchers, K. Kirkegaard, and J. R. Doedens. 1997. Structure-function analysis of coxsackie B3 virus protein 2B. Virology 227:111-118[CrossRef][Medline].
67. Van Kuppeveld, F. J. M., J. G. J. Hoenderop, R. L. L. Smeets, P. H. G. M. Willems, H. B. P. M. Dijkman, J. M. D. Galama, and W. J. G. Melchers. 1997. Coxsackievirus protein 2B modifies endoplasmic reticulum membrane and plasma membrane permeability and facilitates virus release. EMBO J. 16:3519-3532[CrossRef][Medline].
68. Wimmer, E., C. U. T. Hellen, and X. Cao. 1993. Genetics of poliovirus. Annu. Rev. Genet. 27:353-436[CrossRef][Medline].
69. Ypma-Wong, M. F., P. G. Dewalt, V. H. Johnson, J. G. Lamb, and B. L. Semler. 1988. Protein 3CD is the major poliovirus proteinase responsible for cleavage of the P1 capsid precursor. Virology 166:265-270[CrossRef][Medline].

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