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Journal of Virology, January 2007, p. 872-883, Vol. 81, No. 2
0022-538X/07/$08.00+0     doi:10.1128/JVI.01785-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Role of the Amphipathic Peptide of Semliki Forest Virus Replicase Protein nsP1 in Membrane Association and Virus Replication{triangledown}

Pirjo Spuul,1 Anne Salonen,1 Andres Merits,3 Eija Jokitalo,2 Leevi Kääriäinen,1 and Tero Ahola1*

Program in Cellular Biotechnology,1 Electron Microscopy Unit, Institute of Biotechnology, University of Helsinki, Helsinki, Finland,2 Estonian Biocentre and Institute of Molecular and Cellular Biology, Tartu, Estonia3

Received 17 August 2006/ Accepted 26 October 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Semliki Forest virus RNA replication takes place in association with specific cytoplasmic vacuoles, derived from the endosomal apparatus. Of the four virus-encoded replicase proteins, nsP1 serves as the membrane anchor of the replication complex. An amphipathic peptide segment, G245STLYTESRKLLRSWHLPSV264, has been implicated in the membrane binding of nsP1. nsP1 variants with changes within the peptide were studied after protein expression and in the context of virus infection. Proteins with mutations R253E and W259A accumulated in the cytoplasm and were very poorly palmitoylated. The same mutations also drastically affected the localization of the precursor polyprotein P123, and they were lethal when introduced into the virus genome. Mutations R253A and L255A+L256A partially changed the localization of nsP1, and the respective viruses acquired compensatory changes. L255A+L256A only yielded virus encoding L255A+L256V, indicating the importance of a hydrophobic residue in the central 256 position. When fused to green fluorescent protein, the peptide was required in at least two tandem copies to effect a change in localization, but even then the fusion protein was associated with membranes in a nonspecific manner. Thus, the amphipathic peptide is a crucial element for the membrane association of nsP1 and the replication complex. It provides essential affinity for membranes, and other regions of nsP1 also appear to contribute to the localization of the protein.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Upon gaining entry to the host cell cytoplasm, the genome of positive-strand RNA viruses is immediately translated to yield the virus-specific subunits of the RNA-dependent RNA polymerase complex, which then recognizes the genome, synthesizes a complementary minus-strand copy/copies of it, and uses this as a template to make multiple new positive-strand RNAs. This cycle can be rapidly repeated so that an animal cell infected with an efficiently replicating virus can within hours contain thousands of replication complexes and hundreds of thousands of new positive-strand RNAs. Surprisingly, it has turned out that the RNA replication complexes of all positive-strand RNA viruses infecting eukaryotic cells are associated with cytoplasmic membranes (reviewed in reference 38). One or several of the subunits of the replication complex mediate its attachment to membranes. Different virus families use the cytoplasmic surfaces of different membrane compartments as replication platforms, e.g., endoplasmic reticulum (ER), Golgi apparatus, endo/lysosomes, mitochondria, and peroxisomes (10, 15, 20, 30). In most cases the replication complexes also induce morphological alterations of the target membranes, which can interfere with their normal functions.

Alphaviruses, particularly Semliki Forest virus (SFV), are one of the better-understood models for membrane-associated RNA replication. Alphavirus replication utilizes the membranes of the endosomal apparatus, inducing large cytopathic vacuoles (CPVs). Their limiting membrane contains pear-shaped invaginations, or spherules (diameter, ~50 nm), which appear to be the actual sites of RNA synthesis (12, 20). The alphavirus-encoded replication proteins nsP1 to -4 are synthesized as a large nonstructural polyprotein precursor, P1234, which is sequentially processed to its components by a specific protease activity present within the polyprotein. A crucial intermediate is P123, which together with properly cleaved nsP4 is responsible for the synthesis of a complementary minus-strand copy of the 42S genomic RNA (19, 25, 42, 50; reviewed in reference 17). The polyprotein stage is essential for the proper formation and membrane association of the replication complex. The nsPs cannot form a complex if they are simultaneously expressed in the same cell as individual proteins, whereas expression derived from polyprotein P1234 or P123 leads to the membrane association and complex formation (39). Thus, the regulated slow processing of the replicase polyprotein enables both minus-strand RNA synthesis and the correct membrane targeting of the complex. The rate of P1234 proteolysis and the order of cleavages are finely tuned (49) to yield first the negative-strand polymerase complex and then the stable positive-strand polymerase which consists of fully processed nsP1 to -4.

nsP1 (537 amino acids [aa]) is the sole membrane anchor of the replication complex, since the other replication proteins have no independent membrane affinity in cells (39). nsP1 also catalyzes the methyltransferase and guanylyltransferase reactions needed in the capping of viral mRNAs (1, 23, 29), and the membrane binding activates nsP1 as an enzyme (3). The mode of membrane association of nsP1 has been studied in some detail. In infected cells as well as when expressed alone, SFV nsP1 is covalently palmitoylated to three consecutive cysteine residues (418 to 420), which renders the protein highly hydrophobic, resembling integral membrane proteins (21, 32). Related Sindbis virus nsP1 contains only one cysteine residue, 420, in the same region, and it also is modified by palmitoylation (2). Introduction of mutations to the infectious RNAs of the two viruses, which changed these cysteine residues to alanines and prevented palmitoylation, resulted in viable viruses which gave rise to CPVs similar to those seen in wild-type virus-infected cells (2). The membrane association of nsP1 expressed alone also takes place when the protein is not palmitoylated (21), and this weaker peripheral binding to membranes is therefore sufficient for the formation of normal membranous replication complexes. However, removal of the palmitoylation sites reduces the pathogenicity of SFV for mice (2). Only the palmitoylated form of nsP1 induces numerous filopodia-like structures on the cell surface, the significance of which remains unknown (22).

Extensive analysis of truncated nsP1 variants revealed a specific segment in the middle of the protein, G245STLYTESRKLLRSWHLPSV264, whose deletion abolished palmitoylation-independent binding to membranes when nsP1 was expressed in Escherichia coli, which cannot catalyze protein palmitoylation. The corresponding synthetic peptide competed with the binding of nsP1 to liposomes in vitro, although high concentrations of the peptide were required (3). The structure of the synthetic peptide in solution, determined by nuclear magnetic resonance, showed an amphipathic {alpha}-helix. In vitro studies further indicated that both hydrophobic and polar interactions were important in the binding of the peptide to liposomes (24). Due to its affinity for liposomes, this peptide was termed the membrane binding peptide (BP). The current model for the membrane binding of nsP1, and thereby the entire replication complex, proposes that the primary mechanism of membrane association would be binding through the amphipathic peptide, which would be followed by palmitoylation to tighten membrane association (38).

All the previous studies of the BP have been conducted either in vitro with the synthetic peptide or with nsP1 expressed in E. coli cells or produced by in vitro translation. Here we have studied the fate and properties of nsP1 variants with mutations in the BP region in mammalian cells to understand the functions and significance of the peptide in an authentic environment. Our results show that the BP is essential for the membrane binding of nsP1 and for the replication of SFV.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cells and viruses. BHK-21 (baby hamster kidney) cells were used for SFV infection and viral RNA transfection. Cells were cultured in Dulbecco's modified Eagle's medium supplemented with 7.5% inactivated fetal calf serum, 2% tryptose phosphate broth, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. To express SFV nsP1 and its derivatives, HeLa cells were grown in Dulbecco's modified Eagle's medium with 10% fetal calf serum, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin.

Wild-type SFV and mutant viruses were obtained by transfecting capped in vitro RNA transcripts of the infectious cDNA clone of SFV, pSP6-SFV4 (26), and its derivatives into BHK cells (see below). Virus titers were determined by plaque assay (18). Modified recombinant vaccinia virus Ankara (MVA) expressing T7 RNA polymerase, kindly provided by B. Moss (NIH, Bethesda, MD), was propagated in BHK cells (8).

Plasmid constructs. Some of the point mutations in plasmid pTSF1, a derivative of the pGEM3 vector (Promega) encoding SFV nsP1 under the T7 promoter, have been described (3). New mutants were made with the QuikChange XL site-directed mutagenesis kit (Stratagene). Polyprotein construct P12CA3 expressing a protease-defective mutant precursor from the T7 promoter has been described (49). Mutations R253E and W259A were cloned into the precursor in EcoRV-DraIII fragments. Full-length SFV cDNA clones carrying mutations Y249A, R253A, R253E, K254E, L255A+L256A, R257E, and W259A were obtained by substituting a PstI-StuI fragment (nucleotides [nt] 477 to 1464) of the wild-type infectious cDNA with the corresponding fragments from pTSF1 plasmids carrying the mutations. Mutation L255A+L256V was derived from a revertant virus stock by reverse transcription-PCR (RT-PCR). A PCR fragment containing SFV nucleotides 414 to 998 was cloned in the pBluescript II SK(+) vector (Stratagene) and sequenced, and the mutation was transferred to the pTSF1 and pSP6-SFV4 plasmids. The putative compensatory mutations for R253A were derived by RT-PCR from plaque-purified virus stocks. The PCR product (SFV nt 303 to 1759) was cloned into pGEM-T (Promega), and the PstI-DraIII fragment (containing the D202Y mutation), DraIII-StuI fragment (D437G), and PstI-StuI fragment (D202Y+D437G) were transferred to the infectious cDNA clone containing the R253A mutation. The genotypes of all the mutant clones were verified by sequencing.

Plasmid pEGFP-N1 (Clontech) was used to create C-terminal green fluorescent protein (GFP) fusions of the nsP1 binding peptide. Synthetic annealed oligo pairs containing the appropriate cohesive ends, and encoding nsP1 aa 245 to 264, were cloned into pEGFP-N1 as ApaI-AgeI (first copy; oligos CGCGGGCCACCATGGGATCTACATTGTACACTGAGAGCAGAAAGCTACTGAGGAGCTGGCACTTACCCTCCGTA and CCGGTACGGAGGGTAAGTGCCAGCTCCTCAGTAGCTTTCTGCTCTCAGTGTACAATGTAGATCCCATGGTG GCC) or AgeI-AgeI (second and third copies; oligos CACTTACCCTCCGTACCGGGAGCAGGATCTACATTGTACACTGAGAGCAGAAAGCTACTGAGGAGCTGGCACTTACCCTCCGTA and CCGGTACGGAGGGTAAGTGCCAGCTCCTCAGTAGCTTTCTGCTCTCAGTGTACAATGTAGATCCTG CTC) fragments to obtain one to three copies of the peptide fused to GFP. Subsequently, a spacer encoding the amino acid sequence GGSGSAG was added between the peptide and GFP, and the methionine codon in the beginning of GFP was changed to a leucine codon to prevent inappropriate translation initiation yielding GFP not fused to the peptide(s). These changes were introduced in a single step with inverse PCR using primers TCCGCGGGTCTGGTGAGCAAGGGCGAGGAGCTG and TCCACTCCCGCCTACGGAGGGTAAGTGCCAGCTCCTC and Phusion high-fidelity polymerase (Finnzymes). The construct containing three copies of the peptide coding sequence was used as the template. Among the product clones, variants containing one, two, or three peptide sequences with the spacer sequence could be recovered. The clones were verified by sequencing.

RNA transcription, transfection, and infectious center assay. Plasmid pSP6-SFV4 and its mutant derivatives were linearized with SpeI, purified with a PCR purification kit (Genomed), and transcribed in vitro with the SP6 RNA polymerase (Promega). The reaction mixture contained 1 mM each of ATP, CTP, and UTP, 0.5 mM GTP, and 1 mM cap analog m7G(5')ppp(5')G (Amersham Pharmacia Biotech). Usually, 1 µg of RNA was used without purification to transfect BHK cells derived from a nearly confluent 10-cm plate (~107 cells). Control transfections were performed without any nucleic acid. The cells were detached with trypsin and washed once with phosphate-buffered saline (PBS), and electroporation was performed in 800 µl of PBS with two consecutive pulses of 850 V and 25 µF in a Bio-Rad Gene Pulser apparatus in a 0.4-cm cuvette (26). A 720-µl aliquot of the electroporation mixture was used to generate the primary virus stock. These cells were immediately diluted with 10 ml of complete BHK medium and seeded onto a 10-cm dish. The medium was harvested when cytopathic effect (CPE) appeared (at 24 h for the wild-type virus) or after 72 h if no CPE became visible. These primary virus stocks were used in all of the experiments without additional passaging.

The remaining 1/10 (80 µl) of the electroporation mixture was used in an infectious center assay. Tenfold dilutions of the mixture were immediately prepared in minimal essential medium (MEM) containing 0.2% bovine serum albumin (BSA). The dilutions were seeded onto duplicate wells of confluent BHK cells grown on six-well plates, and the transfected cells were allowed to attach for 2 h. Subsequently, the inoculum was replaced with nutrient agarose in complete BHK medium. After incubation at 37°C for 48 to 72 h, plaques were visualized by staining with neutral red. To obtain the relative infectivity values for the mutants R253A and L255A+L256A, 10 µg of these RNAs was used in the electroporation.

To study the putative revertant stock generated after R253A mutant RNA transfection, viruses were plaque purified. Diluted primary stock was used to infect fresh BHK cells, which were overlaid with nutrient agarose and incubated for 3 days at 37°C. Viruses from individual plaques were eluted in 1 ml of MEM-0.2% BSA and used to infect confluent BHK cells on 10-cm dishes. The media were collected at 24 h (CPE was observed at this point) and used for RNA isolation.

DNA transfection, cell fractionation, and isotope labeling. Mutant and wild-type nsP1 and P12CA3 proteins were transiently expressed using the MVA-T7 system. HeLa cells were infected with MVA encoding the T7 RNA polymerase using 20 PFU/cell for 1 h, followed by transfection with plasmid derivatives. Lipofectamine 2000 (Invitrogen) was used as a transfection reagent according to the manufacturer's instructions. For transfections of GFP fusion constructs (expressed from the human cytomegalovirus IE1/2 promoter), ExGen500 reagent (MBI Fermentas) was used.

Lipofectamine 2000 transfection mixture was removed from the cells at 2.5 h posttransfection, and 60 µCi of [9,10(n)-3H]palmitic acid (52 Ci/mmol; Amersham) was added into a 3.5-cm dish in 1.5 ml of MEM-0.2% BSA. After 6 h incubation, cells were washed with PBS and lysed in 100 µl of 1% sodium dodecyl sulfate (SDS). Part of the sample was immunoprecipitated with anti-nsP1 antiserum (20) as described previously (33). Samples were analyzed by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) in 10% gels, which were treated after electrophoresis with Amplify (Amersham Biosciences) according to the manufacturer's instructions, dried, and exposed to preflashed X-ray film. Western blotting was performed as described elsewhere (23).

For cell fractionation, the cells were swollen in a hypotonic buffer (10 mM Tris-HCl, pH 8.0, 10 mM NaCl) for 15 min on ice, followed by disruption in a Dounce homogenizer. KCl (100 mM) was added, and the nuclei were pelleted at 500 x g for 5 min. The postnuclear supernatant was centrifuged at 15,000 x g for 20 min to obtain the P15 and S15 fractions. Membrane flotation was performed in discontinuous sucrose gradients consisting of 10%, 50%, 60% (original sample loading layer), and 67% (wt/wt) layers (3), using the postnuclear supernatant as a starting material.

RNA purification and RT-PCR. Viral RNA was purified from 500 µl of virus stock with RNeasy Mini kit (QIAGEN). Purified and heat-denatured RNA was first reverse transcribed using Moloney murine leukemia virus reverse transcriptase (RNase H minus mutant; Promega) and random hexamer primers (Promega) and then treated with RNase H. The resulting cDNAs were purified with a PCR purification kit (Genomed). PCR was performed with appropriate primers to amplify the entire nsP1 region and with Dynazyme EXT DNA polymerase (Finnzymes).

Immunofluorescence and confocal microscopy. Indirect immunofluorescence microscopy of transfected cells was carried out essentially as described previously (39). Briefly, paraformaldehyde-fixed cells were treated with Alexa488- or Alexa568-conjugated concanavalin A (Sigma) to stain the plasma membrane. Cells were then permeabilized and stained by using anti-nsP1 rabbit antiserum, followed by anti-rabbit antibody conjugated with Alexa568 or Alexa488 (Molecular Probes). Labeled cells were analyzed with the Leica TCS SP2 AOBS confocal microscopy system.

Electron microscopy. For ultrastructural detection of CPVs, SFV-infected BHK cells grown on glass coverslips were fixed with 2.5% glutaraldehyde (Fluka), 0.1 M Na-cacodylate buffer, pH 7.4, for 1 h at room temperature, postfixed with 1% reduced osmium tetroxide for 1 h at room temperature and 1% uranyl acetate, 0.3 M sucrose for 1 h at 4°C, and processed for epon embedding as described previously (41). For immunolabeling, transfected HeLa cells growing on coverslips were fixed with PLP fixative (2% formaldehyde, 0.01 M periodate, 0.075 M lysine-HCl in 0.075 M phosphate buffer, pH 7.4) for 2 h at room temperature, starting at 16 h posttransfection. Cells were permeabilized with 0.01% saponin (Sigma) and immunolabeled using monoclonal anti-GFP (mixture of two mouse monoclonal antibodies, clones 7.1 and 13.1; Roche Diagnostics Corp.) and 1.4-nm gold particle-conjugated Fab' fragments against mouse immunoglobulin G (IgG; Nanoprobes). Nanogold was silver enhanced using an HQ silver kit (Nanoprobes) for 1 to 4 min and gold toned with 0.05% gold chloride (5). Cells were processed for epon embedding as above, except omitting the uranyl acetate en bloc staining. Sections were cut parallel to the coverslip, picked up on single-slot copper grids, poststained with uranyl acetate and lead citrate, and examined using a Tecnai 12 transmission electron microscope (FEI, The Netherlands) at 80 kV.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
According to our previous studies, a synthetic peptide derived from the central part of SFV nsP1 forms an amphipathic {alpha}-helix (Fig. 1A). The hydrophobic surface, consisting of a tryptophan, a valine, and several leucine residues, can interact, in monotopic fashion, with the lipid acyl chains of one leaflet of the lipid bilayer in liposomes, while the positively charged residues (R253, R257, and K254) as well as Y249 would be well placed to interact with the polar head groups of the phospholipids. The peptide laterally sinks to the membrane, so that W259 is on the average located at the depth of 9 to 10 carbons of the acyl chains, as assessed by fluorescence quenching measurements with the synthetic peptide (24). To study the functions of the peptide as part of nsP1 in animal cells, we generated seven mutants of the BP within nsP1, altering positively charged residues in the central part of the BP to neutral alanine or to negatively charged glutamate or altering hydrophobic residues to much less hydrophobic alanine (Fig. 1B).


Figure 1
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  		FIG. 1. Membrane binding peptide of SFV nsP1 and its mutant derivatives. (A) Solution structure of the BP as determined by nuclear magnetic resonance (24). The hydrophobic side of the peptide points downwards. Several protruding amino acid side chains are indicated. (B) Amino acid sequences of the wild-type BP and its mutant derivatives; the mutated residues are underlined. The nucleotide changes made in constructing the mutated amino acid codons are given on the right.

 
Palmitoylation of nsP1 BP mutants. To study the membrane association of nsP1 variants, we expressed the nsP1 proteins in HeLa cells by the aid of vaccinia virus (MVA) encoding T7 RNA polymerase. First, we wanted to know whether the different nsP1 variants would be palmitoylated, since this would be expected to have a significant effect on their behavior. Cells infected with MVA for 1 h were transfected with plasmids encoding nsP1 variants. The cells were labeled with [3H]palmitate for 6 h starting at 2.5 h posttransfection. Whole-cell lysates were collected, and a portion of the samples was subjected to immunoprecipitation with antiserum against nsP1, followed by separation by SDS-PAGE and fluorography (Fig. 2A). Western blotting was used to verify that the variant proteins were expressed at similar levels (Fig. 2B). Mutant proteins Y249A, R253A, K254E, L255A+L256A, and R257E showed palmitate labeling similar to wild-type nsP1 (Fig. 2A, lanes 1, 3, 4, and 6 to 8). A negative control mutant, Pa– (C418 to 420A), in which the palmitoylation site is destroyed, was not labeled, as expected (lane 2). Mutants R253E and W259A were palmitoylated at very low levels (lanes 5 and 9). Quantitation of incorporated [3H]palmitate by densitometry showed that W259A and R253E had less than 10% of palmitate label compared to wild-type nsP1.


Figure 2
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  		FIG. 2. Palmitoylation and membrane association of nsP1 binding peptide mutants. (A) HeLa cells transfected to express nsP1 and its indicated derivatives were labeled with [9,10(n)-3H]palmitic acid and collected into 1% SDS. A fraction of the cell lysates was immunoprecipitated with polyclonal rabbit anti-nsP1 antiserum, and the immunoprecipitates were subjected to SDS-PAGE and fluorography to visualize the labeled proteins. Molecular mass markers are shown on the left in; nsP1 migrates at its expected size of 64 kDa. (B) Portions of the lysates were also analyzed by Western blotting using the anti-nsP1 antiserum to detect the relative amounts of nsP1 variants expressed. (C) HeLa cells expressing the indicated proteins were lysed, the nuclei were removed, and the cytoplasmic extract was fractionated to yield cytoplasmic membranes sedimenting at 15,000 x g and the remaining supernatant. Equal amounts of the pellet (P15) and supernatant (S15) fractions were analyzed by SDS-PAGE and Western blotting with anti-nsP1 antiserum.

 
It has been reported previously that wild-type nsP1 is associated almost exclusively with the cytoplasmic membrane fraction sedimenting at 15,000 x g (P15) (21). Proteins were expressed by transfection as described above in HeLa cells for 10 h, followed by the isolation of P15 and the respective supernatant (S15) fractions. After SDS-PAGE separation, the proteins were detected by immunoblotting with anti-nsP1 antiserum (Fig. 2C). Wild-type nsP1 and nonpalmitoylated mutant Pa– served as controls. Wild-type nsP1 was found quantitatively in the P15 fraction, whereas Pa– was rather equally distributed between the P15 and S15 fractions (Fig. 2C). In this fractionation experiment, Y249A, R253A, K254E, L255A+L256A, and R257E were predominantly found in the P15, in accordance with their palmitoylation, whereas R253E and W259A behaved similarly to the palmitoylation-defective Pa–.

Intracellular localization of BP mutant proteins. Expression of the wild-type nsP1 alone results in the accumulation of the protein at the cytoplasmic side of the plasma membrane. The palmitoylated protein induces also filopodia-like extensions, which stain intensively with anti-nsP1 antibodies according to indirect immunofluorescence microscopy and immuno-electron microscopy (22, 32). To study the subcellular localization of the BP mutants, the constructs encoding different nsP1 mutants were expressed in HeLa cells by the MVA system as above. Samples were fixed at 6 h posttransfection and immunostained with anti-nsP1 antibodies. The cells were also stained with concanavalin A (ConA) before the permeabilization to show the contours of the plasma membrane.

Confocal fluorescence images of the cells expressing the wild-type nsP1 and the mutant variants are shown in Fig. 3. The typical staining of the plasma membrane and of numerous filopodia was seen in the cells expressing wild-type nsP1 (Fig. 3A). The palmitoylation-defective Pa– was also often found on the plasma membrane, but not in filopodia-like structures (Fig. 3B). Staining of both the plasma membrane and filopodia was seen in cells expressing Y249A, R253A, K254E, L255A+L256A, and R257E (Fig. 3). In contrast, cells expressing R253E as well as W259A (Fig. 3E and I) showed no costaining at the plasma membrane, which was visualized by ConA labeling. Rather, there was a diffuse staining throughout the cytoplasm, without any apparent specificity for certain structures, although the proteins were excluded from the nucleus (best visible in the section shown in Fig. 3I). When comparing the other mutants with wild-type nsP1, it was apparent that K254E and R257E very closely resembled the wild type, in that virtually all the nsP1 signal is on the plasma membrane with very little internal staining (Fig. 3A, F, and H). In contrast, while a fraction of nsP1 costained with ConA on the plasma membrane in mutants Y249A, R253A, and L255A+L256A, there was also significant disperse staining in the cytoplasm (Fig. 3C, D, and G). Immunoelectron microscopy of the cells expressing either wild-type nsP1 or W259A was also used to assess the altered localization. Wild-type nsP1 showed heavy staining beneath the plasma membrane and in filopodia (Fig. 4A), whereas W259 appeared rather homogenously distributed in the cytoplasm (Fig. 4B). No specific organelle staining was revealed by the preembedding immunogold labeling technique, and there was no specific association of W259A with the plasma membrane, in accordance with the confocal fluorescence images.


Figure 3
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  		FIG. 3. Confocal immunofluorescence localization of wild-type and mutant nsP1 proteins. HeLa cells infected with vaccinia virus MVA followed by transfection with plasmid encoding the indicated nsP1 variants were fixed and stained with Alexa488-ConA (green) to visualize the plasma membrane and then permeabilized for Alexa568-anti-nsP1 staining (red). Colocalization of these two is seen in yellow. Each image represents one confocal microscopy section (0.24 µm in thickness), selected to give a representative staining of both the plasma membrane and internal structures. The sections are close to the bottom of the cell, typically ~1 µm from the glass surface.

 

Figure 4
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  		FIG. 4. Electron microscopic localization of wild-type and W259A nsP1. HeLa cells expressing wild-type nsP1 (A) or W259A (B) proteins were processed for immuno-electron microscopy detection of nsP1, as described in Materials and Methods. Bar, 400 nm.

 
It was also of interest to study the localization of the precursor polyprotein P123, since it is involved in the formation of the replication complexes during infection. To prevent the self-cleavage of the polyprotein, we used a protease active site mutant P12CA3 (49). Previous studies showed that P12CA3 targets differently from nsP1: the precursor was not found on the plasma membrane but instead resided in numerous punctuate structures in the cytoplasm, some of which were endo/lysosomal vesicles (39). The wild-type P12CA3 was again localized in punctuate structures (Fig. 5A), but when the mutation W259A was introduced to the precursor, the protein was found diffusely all over the cytoplasm but not in the nucleus (Fig. 5B). The precursor with the mutation R253E was rather poorly expressed, but in all the positive cells it was also diffusely localized (Fig. 5C). These results suggest that the BP is also important for the localization and membrane binding of the polyprotein precursor.


Figure 5
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  		FIG. 5. Confocal immunofluorescence localization of the polyprotein precursor P12CA3 wild type (A) and mutants W259A (B) and R253E (C). HeLa cells infected with vaccinia virus MVA followed by transfection with plasmids encoding the indicated variants were fixed and stained with anti-nsP3 antibodies (red). Staining with anti-nsP1 antibodies (not shown) was identical, indicating the intact state of the polyprotein.

 
Localization of binding peptide-GFP fusion proteins. To study the targeting properties of the BP in a heterologous context, the BP sequence was joined to the expression plasmid pEGFP-N1 to produce GFP fusion proteins with a single, duplicate, or triplicate copy of the BP at the amino terminus of GFP (Fig. 6A). After 16 to 24 h of expression in HeLa cells from the cytomegalovirus promoter, the fusion proteins were easily detected by immunoblotting with GFP antibodies. They showed the expected increase in molecular weight conferred by the added moieties (Fig. 6B). No additional intermediates or degradation products that might confound the localization studies were seen in the immunoblots. The earlier time points (16 to 18 h) were selected for microscopic examination to avoid overexpression. According to simple fractionation of the cytoplasm by centrifugation, GFP and the 1xBP fusion were soluble at 15,000 x g, but the 3xBP fusion was almost quantitatively and 2xBP fusion partially found in the P15 fraction (Fig. 6C). When the postnuclear supernatant was subjected to flotation in discontinuous sucrose gradients, the GFP and 1xBP fusion did not float, but the 2xBP and 3xBP fusions partially floated with membranes (Fig. 6D), indicating that they had acquired membrane binding properties. There was essentially no difference in the flotation behavior of the 2xBP and 3xBP fusions; relatively weakly bound proteins may become partly dissociated from membranes during the flotation procedure. Wild-type nsP1 was used as a positive control in this experiment, and it showed almost quantitative flotation with membranes (Fig. 6D).


Figure 6
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  		FIG. 6. Expression and characterization of BP-GFP fusion proteins in HeLa cells. (A) The BP was fused in frame with GFP, in one, two, and three copies with short spacer sequences between the copies and a longer spacer to separate the peptides from GFP. The original initiating methionine of GFP was mutated to leucine to prevent the production of free GFP from the fusion protein constructs through the use of a downstream initiation site. (B) Extracts of cells expressing the indicated proteins were prepared, and expression of the GFP constructs was detected by SDS-PAGE and Western blotting with GFP antibodies. Molecular mass markers are given on the left. (C) Cytoplasmic extracts from cells expressing the GFP constructs were fractionated by centrifugation at 15,000 x g to yield the supernatant (S15) and pellet (P15) fractions, which were analyzed by SDS-PAGE and Western blotting. (D) Cytoplasmic extracts were analyzed by flotation in discontinuous sucrose gradients as indicated in Materials and Methods. The sample was originally loaded near the bottom, from where membranes floated during centrifugation. Fractions were collected from the top and subjected to immunoprecipitation with the relevant antibodies, and the precipitates were analyzed by Western blotting. Wild-type nsP1 (bottom panel) was used as a positive control (totally membrane associated), and GFP (top panel) was used as a negative control (soluble protein). The heavy chain of immunoglobulin molecules (labeled as IgG on the right) is also visualized by blotting and acts as a molecular mass marker. The protein expressed in each panel is indicated on the right at its position of migration. The top fractions of the gradient are located on the left, and bottom fractions are on the right. (E to G) HeLa cells expressing the BP-GFP fusion constructs were fixed 18 h after transfection, and the localization of GFP was detected by using GFP fluorescence. (E) GFP without fusion; (F) 2x BP fusion; (G) 3x BP fusion. The plasma membrane was visualized with Alexa568-ConA (red).

 
Fixed cell samples expressing these GFP-containing proteins were subjected to fluorescence confocal microscopy. GFP expressed alone was localized to the nucleus and the cytoplasm (Fig. 6E). Similar GFP localization was seen when the fusion protein construct contained only one copy of the binding peptide (not shown). However, a different distribution started to appear when a duplicate copy of BP was joined to the N terminus of GFP. In most cells, GFP was not so predominantly seen in the nucleus, and the cytoplasmic staining was partially punctuate, although diffuse staining was also observed (Fig. 6F). The new features were further increased with three copies of the BP fused to GFP. This construct showed in most cells punctuate staining in the cytoplasm (Fig. 6G). The 3xBP fusion was also localized by immunoelectron microscopy, in which it was found associated with various membranous organelles in the cytoplasm. It was most often visualized on mitochondrial membranes, but also on lysosomes and sometimes on the ER (data not shown). Thus, when placed in the amino terminus of GFP, the BP was unable to direct GFP to a specific intracellular destination but in two or three copies caused the protein to associate with several types of membranous organelles.

Effects of BP mutations on SFV replication. We incorporated the BP mutations into the infectious cDNA of SFV to study their effects on virus replication. In vitro-transcribed, capped 42S RNAs were transfected into BHK cells by electroporation. In wild-type SFV-infected cells, CPE manifested within 24 h, and already at 12 h postinfection peak virus titers of the order of 109 PFU/ml were found in the medium (Fig. 7A). In the same experiment the relative infectivities of the RNAs were measured by using infectious center assay. Dilutions of the electroporated cells were allowed to attach onto plates containing fresh confluent BHK cells and overlaid with nutrient agar for 2 to 3 days to permit the formation of plaques, each arising from a productively infected single cell upon electroporation. Wild-type SFV yielded 106 plaques/µg RNA (Table 1), which is similar to the value originally reported (26). Using the appearance of infectious virus and CPE as well as the relative infectivity as criteria, mutants K254E and R257E behaved similarly to the wild-type SFV (Table 1). Sequencing was used to verify that these virus stocks still contained the expected mutations. When the primary virus stocks obtained after electroporation were used to infect BHK cells at 10 PFU/cell, no significant differences in the growth of wild type, K254E, and R257E were observed (Fig. 7B).


Figure 7
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  		FIG. 7. Growth curves of SFV containing mutations. (A) Growth of viruses with mutations in the BP after RNA transfection. Virus growth was studied in BHK cells after electroporation of 1 µg of capped RNA transcript containing the indicated mutations. Virus titers in aliquots withdrawn at different times were determined by plaque assay and are expressed in PFU/ml. (B) Growth of stock viruses. The virus stock collected at 24 h or 72 h post-RNA transfection was titrated and used to infect BHK cells at 10 PFU/cell. Virus appearance in the cell culture supernatant was followed by taking aliquots at 2-h intervals, which were subjected to plaque titration. (C) Growth curves of SFV containing compensatory mutations. Virus growth was studied after electroporation of 1 µg of capped RNA transcript containing the indicated mutations into BHK cells. Virus titers in aliquots withdrawn at each time point were determined by plaque assay.

 

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  		TABLE 1. Properties of viruses and nsP1 proteins containing mutations in the BP

 
Cells transfected with Y249A, W259A, and R253E mutant RNAs consistently failed to exhibit CPE even at 72 h, and the supernatants were negative at 24 h, 48 h, and 72 h after transfection when monitored for virus production. Cells transfected with these mutants also failed to show any evidence of structural protein production in immunofluorescence analysis. Furthermore, in the infectious center assay, no plaques were observed. Thus, we conclude that each of these three mutations is individually lethal for SFV replication.

Mutants R253A and L255A+L256A behaved in an interesting intermediate fashion. They slowly gave rise to CPE, the initial signs of which were visible at 48 h. The titers of the viruses in the medium also rose slowly, and viruses were first observed only at 24 h in low amounts (Fig. 7A). The relative infectivities of the RNAs were very low (Table 1), and these numbers could only be obtained when 10-fold more RNA than normal was used in the infectious center assay. The few plaques observed in the assay were usually smaller than those of the wild-type virus. Thus, either these two viruses replicated extremely poorly or suppressor mutations were generated during the incubation. This experiment was repeated three times for each mutant, with closely similar results. When one selected primary stock derived from each mutant, collected at 72 h, was used to infect fresh BHK cells at 10 PFU/cell, the virus growth rates were only very slightly delayed compared to the wild type (Fig. 7B). This indicates that the initial growth impairment observed after electroporation was no longer present in the stock viruses.

For the mutant L255A+L256A, five independent virus stocks were generated by separate transcription and electroporation reactions, collected at 72 h, and subjected to sequencing. The initial sequence coding for the dipeptide Ala-Ala at this position of the infectious cDNA was GCTGCG, whereas each of the stocks contained the sequence GCTGTG, which encodes Ala-Val. In the case of the stock selected for growth studies (Fig. 7B), five independent PCR clones were sequenced. Again, they all gave the sequence GCTGTG, demonstrating that the stock was homogenous for this mutation. We conclude that the mutant cDNA L255A+L256A reproducibly gave rise only to a revertant virus which encoded 255A and 256V. To further confirm this result, we constructed an infectious cDNA containing the combined mutation L255A and L256V. The relative infectivity of the RNA derived from this was 1.1 x 106 PFU/µg (Table 1), and the virus appeared rapidly after electroporation, at a rate and yield similar to the wild type (Fig. 7C). Confocal and electron microscopy were used to compare BHK cells infected with L255A+L256V to wild-type virus-infected cells. The latter method showed the presence of typical CPVs containing the spherule invaginations (Fig. 8A). In confocal microscopy, vacuoles costaining with the replication proteins nsP1 and nsP3 were observed (Fig. 8B), again indistinguishable from wild-type virus-infected cells. Thus, mutant L255A+L256A was quasi-infectious (34) and was only able to grow when A256 mutated to V, increasing the hydrophobicity of the BP and giving a wild-type replication phenotype. When expressed alone, the nsP1 protein containing the mutation L255A+L256V showed a wild-type-like pattern of plasma membrane and filopodia staining (Fig. 8C).


Figure 8
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  		FIG. 8. Morphology of cells infected with the L255A+L256V mutant virus or expressing the L255A+L256V nsP1. (A) Electron micrograph of an infected BHK cell (5 h postinfection) showing the presence of cytopathic vacuoles containing the spherule invaginations. In a few spherules (arrows), the connecting neck is visible in the plain of this section. Bar, 400 nm. (B) Confocal fluorescence image of an infected cell at the same time point, showing the colocalization of nsP1 and nsP3 in the replication complex containing vacuoles (yellow). Additionally, nsP1 (green) is found on the plasma membrane and nsP3 (red) is in other cytoplasmic structures, likely representing protein aggregates (39). (C) Confocal fluorescence image of nsP1 L255A+L256V expressed in HeLa cells by the MVA-T7 system. The expression conditions were as for Fig. 3. nsP1 staining is shown in red, the plasma membrane marker ConA is in green, and colocalization is shown in yellow.

 
Three independent stocks derived from electroporation of mutant R253A were also sequenced. In this case, all of the stocks showed the presence of the original alanine codon, GCA. Considering the possible heterogeneity of the virus stocks, one of the stocks was subjected to plaque cloning, and virus genomes derived from 10 independent plaques were sequenced at this position. Again all gave the original sequence, indicating that the mutation was maintained. However, given that the original infectivity was very low but the resulting virus stock showed no impairment in growth, we had strong reasons to suspect that genetic alterations had taken place, as they had in the mutant L255A+L256A. Further sequencing revealed, in each of the virus clones, mutations in nsP1, some of which may be involved in suppressing the original mutation. These mutations were found in several locations within nsP1 in different clones, but none were inside or in the immediate vicinity of the BP. One well-growing virus clone was selected for initial analysis. It contained two single-nucleotide substitutions in nsP1, encoding changes D202Y and D437G. These changes, both separately and combined, were examined in the R253A virus background. While both changes independently restored the infectivity of R253A, it remained 15 to 50 times lower than wild type (Table 1). The growth of these viruses was also significantly delayed compared to the wild type (Fig. 7C). Only the combination of these changes fully restored the wild-type level of infectivity and growth to the R253A mutant (Table 1; Fig. 7C).


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The genome replication of all positive-strand RNA viruses infecting eukaryotic cells is associated with cytoplasmic membranes. Alphaviruses are a favorable system to analyze this phenomenon, since only one of their replication proteins, nsP1, has affinity for membranes. We show here that the membrane binding property of nsP1 is essential for SFV replication. For the other major groups of animal viruses that have been studied with regard to membrane association of the replicase, the mechanisms for membrane attachment appear to be more complex and involve several viral proteins. In hepatitis C virus and other members of Flaviviridae, several replicase proteins contain transmembrane and/or amphipathic helices, which direct the proteins into the endoplasmic reticulum membrane (10, 14, 27, 40). In poliovirus and other picornaviruses, three of the replicase proteins have hydrophobic or amphipathic sequences, which play a role in their membrane association (9, 11, 31, 44, 45, 47). Likewise, the nidoviruses (arteri- and coronaviruses) possess several replicase proteins with hydrophobic properties (43, 48).

In alphaviruses, the mRNA capping enzyme nsP1 plays a decisive role in the membrane binding of the replication complex, since the other three replicase proteins cannot directly associate with membranes (39). nsP1 is not a classical integral membrane protein with a hydrophobic transmembrane sequence(s). Instead, its membrane association was proposed to be mediated by a binding peptide, which forms an amphipathic {alpha}-helix and which is located in the middle of the protein sequence (3, 24) (Fig. 1). Secondly, covalent palmitoylation of nsP1 can tighten its membrane binding, but palmitoylation is not essential for the proper formation or targeting of the viral replication complexes or for virus viability (2). Our present results strongly support the primary role of the amphipathic helix in mediating nsP1 membrane association. Two of the initial seven BP mutant proteins used in this study could no longer become palmitoylated, whereas the lack of palmitoylation does not prevent membrane binding through the BP (3). These two mutations, R253E and W259A, caused a total redistribution of nsP1 from the plasma membrane into the cytoplasm (Fig. 3). These mutations also caused the polyprotein P123 to become diffusely localized. When transferred to the genome of the virus, the mutations were lethal, and no virus replication could be observed even in prolonged incubations. Based on our results, lethality and lack of palmitoylation are two consequences of the primary failure of nsP1 to bind to membranes through the BP. The current results show that loss of membrane binding is lethal. Secondly, nsP1 can only become palmitoylated after associating to membranes via the BP. Palmitoylation is not essential, but it is important in vivo, as loss of palmitoylation greatly reduces SFV pathogenicity in mice (2). The functions of nsP1 palmitoylation are not adequately understood, and they could be more significant in particular cell types.

Three other mutations (Y249A, R253A, and L255A+L256A) caused an intermediate phenotype in nsP1 localization. A fraction of the protein was found at the plasma membrane, but significant staining of the cytoplasm was also observed. These proteins could become palmitoylated, but this was not sufficient to rescue a fully infectious virus phenotype. The mutation Y249A was lethal, and the other two mutants acquired, at a low frequency, compensatory mutations, which permitted their replication at levels resembling wild type. Thus, the original mutations were so debilitating that no progeny viruses with the original genotypes were recovered. The mutant L255A+L256A always reverted to Val at position 256. The replication complexes of the virus L255A+L256V resembled the wild type at both light and electron microscopic levels (Fig. 8). This increase in hydrophobicity was achieved by a single nucleotide change (GCG to GTG), which was repeatedly observed in independent experiments. Reversion to the original Leu or to any other hydrophobic residue besides Val would have required a change of two nucleotides. The same holds true for position 255, but no alterations arose there, indicating the importance a hydrophobic residue at position 256. According to the structure of the isolated BP, the crucial residues revealed by the mutagenic analysis (Y249, R253, L256, and W259) form a continuous region on the BP, extending diagonally across the peptide from the more hydrophilic side (Y249 and R253) to the hydrophobic W259. L256 occupies a central position in this critical patch. These results support the idea that BP also as part of nsP1 would form an amphipathic {alpha}-helix.

According to fluorescence studies with the synthetic BP, W259 sinks deeply to the hydrophobic layer of the membrane, and the average depth of the tryptophan is at the level of 9 to 10 carbon atoms of the acyl chains (24). Based on the structure of the peptide, L256 is also expected to contribute to the hydrophobic interactions of membrane binding. R253 extends to the opposite direction from W259 and could interact with negatively charged phospholipid head groups, as the peptide only binds to vesicles containing negatively charged lipids. While Y249 could also be involved in direct lipid interactions, we alternatively speculate that it could be part of the link between the BP and the rest of nsP1 and perhaps even with the active site of the protein. The mutation Y249A was strictly lethal for the virus, yet it did not reduce the membrane localization of nsP1 as drastically as did the mutation of W259 or R253 (Fig. 3). Sequence comparisons in alphaviruses (24) show that all four of the crucial residues identified here are strictly conserved. However, as the general sequence identity among alphavirus nsPs exceeds 60%, conservation is not very informative. When comparisons are made between alphavirus nsP1 and the related proteins from the alphavirus superfamily (mostly plant viruses), the sequence identity becomes extremely low (generally below 15%). As aligned by Rozanov and coworkers (36), the BP region is among the more conserved segments of the protein, and the four residues are either maintained (Y249) or display mostly conservative substitutions (R253 being most variable). However, there is as yet no evidence linking this region to membrane binding in these very distantly related viruses.

The mutant R253A did not revert at the same site or in its vicinity, but instead acquired changes elsewhere in nsP1. These again were single nucleotide alterations compared to the original sequence. Notably, the Ala codon could not have reverted to positively charged Arg or Lys by a single nucleotide change. Work is in progress to fully characterize the set of the compensatory mutations, and their mode of action. One initially studied virus clone contained two mutations, D202Y and D437G, which probably arose sequentially during the growth of the virus. Both restored the replication partially, and in combination they effected full compensation. Both of these mutations remove a negative charge. Further studies are required to examine whether these regions of nsP1 directly contribute to membrane binding or stabilize the BP by intramolecular interactions.

During infection and if expressed alone, nsP1 is found at the inner surface of the plasma membrane (Fig. 3) (20, 21), suggesting that it is specifically targeted there. We have hypothesized that the binding peptide might serve as a simple targeting signal for nsP1 (20), since the BP has special affinity to anionic phospholipids. The best binding of the synthetic BP to liposomes was obtained when they contained 20 to 30% phosphatidylserine (24). As the cytoplasmic side of the plasma membrane is rich in phosphatidylserine (4), it was tempting to assume that the BP would be involved in targeting nsP1 there. The participation of the BP in this process is evident, since several mutations of the BP either abolished or reduced the plasma membrane localization of nsP1. However, the BP does not appear to constitute an independent and specific protein-targeting element. We found that one copy of the BP, when added to the N terminus of GFP, was insufficient to change GFP localization in cells. The peptide by itself interacts with liposomes in vitro (24), but the affinity for membranes may be insufficient to target the fusion protein or, alternatively, the peptide might not attain its proper conformation as part of the heterologous fusion. Therefore, the fusion protein results should be treated with some caution. A short amphipathic helix has a relatively weak affinity for membranes, and therefore several helices or additional mechanisms such as lipid modification or positively charged segments are often involved in membrane association (16). Two and especially three tandem copies of the BP caused unspecific GFP association with several membranous organelles. Thus, the BP provides at least a propensity for membrane localization.

It is increasingly clear that many proteins are associated with membranes by nonclassical means, either through amphipathic {alpha}-helices or through yet other mechanisms (reviewed in reference 16). There are a few examples where amphipathic helices affecting membrane binding and protein localization have been well characterized. Hepatitis C virus nonstructural protein 5A possesses an amphipathic helix of 30 aa at its N terminus, which can direct the protein to the ER membranes. Remarkably, this sequence is also capable of targeting a heterologous protein (GFP) to the ER (6, 40). Several cellular proteins also possess amphipathic helices which are used for membrane binding and targeting purposes. The N-terminal helices of regulator of G protein signaling 2 and Rho guanine nucleotide dissociation inhibitor 3 can direct these proteins to the plasma membrane and Golgi complex, respectively (7, 13). A C-terminal helix mediates the plasma membrane localization of G protein-coupled receptor kinase 5 (46). All these helices are also able to mediate the targeting of GFP, although the relocalization can be only partial.

In contrast, the BP of SFV nsP1 is located centrally in the protein sequence, and it is likely to be in intimate interaction with the rest of the protein. It is not surprising that when removed from its natural context, it is not a totally independent functional unit. The palmitoylation site of nsP1 is also unusual in laying in the central part of the protein, although distant in primary sequence from the amphipathic helix. In most cases, protein palmitoylation takes place close to the N or C terminus of a protein (which often is additionally modified by myristoylation or prenylation, respectively) or adjacent to transmembrane helices (35). The crystal structures of five monotopic membrane proteins have been solved (28, 37). They interact with membranes through amphipathic segments which can be located centrally in the protein. These proteins are all enzymes interacting with lipophilic substrates; the membrane association areas are often close to the substrate binding pocket, and their membrane-targeting properties have not been studied in isolation. As an RNA capping enzyme, nsP1 interacts with hydrophilic nucleoside-containing substrates.

The targeting determinant of nsP1 appears to be complex, with contributions from regions outside the BP. Some of these regions may be pinpointed through genetic interactions, such as those being analyzed for the R253A substitution. The targeting of the alphavirus replication complex currently appears to contain three layers: (i) the BP mediates membrane binding; (ii) further determinants within nsP1 target the protein to the plasma membrane; and (iii) signals in the other, soluble, domains of the precursor P123 are required for nsP1 and the replication complex to reach their final destination at the endo/lysosomal membrane, and at least the nsP3 portion appears to participate in this process (39).


   ACKNOWLEDGMENTS
 
We thank Mervi Lindman and Merja Joensuu for technical assistance in electron microscopy.

This work was supported by Academy of Finland grant 211121 and by the Sigrid Juselius Foundation, as well as by the European Union 5th Framework Program project SFVECTORS. E. Jokitalo is supported by Academy of Finland (201441) and University of Helsinki research funds.


   FOOTNOTES
 
* Corresponding author. Mailing address: Institute of Biotechnology, P.O. Box 56 (Viikinkaari 9), University of Helsinki, 00014 Helsinki, Finland. Phone: 358-9-19159403. Fax: 358-9-19159560. E-mail: tero.ahola{at}helsinki.fi. Back

{triangledown} Published ahead of print on 8 November 2006. Back


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 Introduction
 Materials and Methods
 Results
 Discussion
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Journal of Virology, January 2007, p. 872-883, Vol. 81, No. 2
0022-538X/07/$08.00+0     doi:10.1128/JVI.01785-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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