The first hydrophobic domain of the hepatitis C virus E1 protein is important for interaction with the capsid protein

Hsin-Chieh Ma1, Cheng-Hung Ke1, Tsai-Yuan Hsieh4 and Shih-Yen Lo1,2,3

Institute of Medical Research1 and Department of Medical Technology2, Tzu Chi University, 701, Section 3, Chung-Yang Road, Hualien, Taiwan 970, Republic of China
Department of Medical Technology, Buddhist Tzu Chi General Hospital, Hualien, Taiwan, Republic of China3
Department of Internal Medicine, Tri-Service General Hospital, National Defense Medical Center, Taipei, Taiwan, Republic of China4

Author for correspondence: Shih-Yen Lo at Department of Medical Technology. Fax +886 3 8571917. e-mail losylo{at}mail.tcu.edu.tw


   Abstract
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Abstract
Introduction
Methods
Results
Discussion
References
 
The interaction between the hepatitis C virus capsid protein and the envelope protein E1 has been demonstrated previously in vivo. To determine the binding region of the E1 protein with the capsid protein, this interaction was characterized in vitro. This study shows that the interaction between these proteins should occur in the endoplasmic reticulum membrane rather than in the cytosol and that the first hydrophobic domain of the E1 protein (aa 261–291) is important for the interaction with the capsid protein.


   Introduction
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Abstract
Introduction
Methods
Results
Discussion
References
 
Hepatitis C virus (HCV) infection is associated with the development of hepatitis, cirrhosis and hepatocellular carcinoma (HCC) (Houghton, 1996 ). HCV is a member of the Flaviviridae family (Trepo et al., 1997 ) and is an enveloped, positive-stranded RNA virus with a genome of 9–10 kb. The HCV genome encodes a polyprotein with a length of over 3000 aa. This polyprotein is cleaved by cellular and viral proteases to generate at least 10 viral gene products (Grakoui et al., 1993 ), which are arranged in the order NH2–C (21 kDa)–E1(31 kDa)–E2(70 kDa)–p7–NS2(23 kDa)–NS3(70 kDa)–NS4A(8 kDa)–NS4B(27 kDa)–NS5A(58 kDa)–NS5B(68 kDa)–COOH. The capsid (C) protein and the envelope proteins E1 and E2 are structural proteins, while NS2–NS5B are non-structural proteins. The processing of the capsid protein with the downstream E1 protein is mediated by a cellular signal peptidase to generate a capsid protein of either 191 and/or 173 aa in length (Santolini et al., 1994 ; Yasui et al., 1998 ). In addition to its structural properties, the HCV capsid protein could also have multiple regulatory functions (Lai & Ware, 2000 ). The HCV capsid protein could regulate various viral and cellular gene promoters (Ray et al., 1995 ; Shrivastava et al., 1998 ), interact with several cellular signalling proteins (Heim et al., 1999 ; Hsieh et al., 1998 ; Lu et al., 1999 ) and even induce HCC in transgenic mice (Moriya et al., 1998 ).

Morphogenic studies of HCV have been hampered by the lack of a cell culture system for the efficient propagation of this virus. The interaction of the HCV capsid protein with positive-sense RNA has been characterized previously (Shimoike et al., 1999 ). The interactions between the structural proteins of HCV are also important for the morphogenesis of this virus. There are extensive interactions between these structural proteins: the capsid protein can interact both with itself (Lo & Ou, 1998 ; Matsumoto et al., 1996 ) and with E1 (Baumert et al., 1998 ; Lo et al., 1996 ) and E2 (Baumert et al., 1998 ); the E1 protein can also interact with the E2 protein (Dubuisson et al., 1994 ; Yi et al., 1997 ).

It has been demonstrated that the C-terminal sequences of both capsid and E1 proteins are important for their interaction in vivo (Lo et al., 1996 ). In order to determine the binding region of the E1 protein with the capsid protein, we have performed proteinase K protection assays to study the topology of the E1 protein, and glutathione S-transferase (GST) pull-down assays to study the interaction between the capsid protein and various E1 mutant proteins in vitro.


   Methods
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Abstract
Introduction
Methods
Results
Discussion
References
 
{blacksquare} Construction of expression plasmids.
Expression plasmids of GST–capsid fusion proteins and serially truncated E1 proteins with signal peptides (to aa 380, 370, 360, 350 and 340) have been described previously (Hsieh et al., 1998 ; Lo et al., 1996 ).

To clone the complete E1-coding sequence (aa 192–383), primers 192-S (5' GGAATTCCATATGTACCAAGTGCGCAATTC 3') and 383-AS (5' GAAGATCTTTACGCGTCGACGCC 3') were used (underlined nucleotides indicate restriction sites; bold nucleotides indicate a stop codon). After amplification, the DNA fragment was digested with the restriction enzymes NdeI/BglII. This DNA insert was cloned into the pET3a vector (Novagen), linearized previously with NdeI/BamHI, to generate plasmid C383.

To clone the E1-coding sequence without the second hydrophobic domain (H2), primers 192-S and 328-AS (5' CGGGATCCTTAAGGGGACCAGTTCAT 3') were used. After amplification, the DNA fragment was digested with NdeI/BamHI. This DNA insert was cloned into pET3a, linearized previously with NdeI/BamHI, to generate plasmid C328.

To clone the E1-coding sequence without the first hydrophobic domain (H1), primers 192-S and 260-AS (5' GAGAAGGTACGTCGAAGCTGCGT 3') were used to amplify the gene fragment from aa 192 to 260, while primers 292-S (5' TCGACGTACCTTCTCTCCCAGG 3') and 383-AS were used to amplify the gene fragment from aa 292 to 383 (nucleotides in bold indicate the codon for aa 260; underlined nucleotides indicate the codon for aa 292). These two DNA fragments were linked and amplified using primers 192-S and 383-AS. After that, the DNA fragment was digested with NdeI/BglII. This DNA insert was cloned into pET3a, linearized previously with NdeI/BamHI, to generate plasmid C383D.

To clone the E1-coding sequence without either the H1 or the H2 domain, primers 192-S and 260-AS were used to amplify the gene fragment from aa 192 to 260, while primers 292-S and 328-AS were used to amplify the gene fragment from aa 292 to 328. These two DNA fragments were linked and amplified using primers 192-S and 328-AS. After that, the DNA fragment was digested with NdeI/BamHI and cloned into pET3a to generate plasmid C328D.

To clone the entire C–E1-coding sequence (HCV nt 321–1517), primers 321-S (5' CGGAATTCAGGTCTCGTAGACCG 3') and 1517-AS (5' GCTCTAGATTAGGCACTTCCCCCGGT 3') were used. After amplification, the DNA fragment was digested with EcoRI/XbaI. This DNA insert was then cloned into pcDNA3 (Invitrogen), linearized previously with EcoRI/XbaI, to generate plasmid pcDNA3-CCE1.

To clone the entire C–E1-coding sequence without the H1 domain, primers 321-S and 260-AS were used to amplify the gene fragment from nt 321 to 1121 (aa 260), while primers 292-S and 1517-AS were used to amplify the gene fragment from nt 1215 (aa 292) to 1517. These two DNA fragments were linked and amplified using primers 321-S and 1517-AS. After that, the DNA fragment was digested with EcoRI/XbaI. This DNA insert was cloned into pcDNA3, linearized previously with EcoRI/XbaI, to generate plasmid pcDNA3-CCE1D.

To clone the entire C–E1-coding sequence, but deleting aa 119–152, primers 321-S and 118-AS (5' GACGCCATGATTGCGCGACCTACG 3') were used to amplify the gene fragment from nt 321 to 695 (aa 118), while primers 153-S (5' TCGCGCAATCATGGCGTCCGGGTT 3') and 1517-AS were used to amplify the gene fragment from nt 798 (aa 153) to 1517 (underlined nucleotides indicate the codon for aa 118; bold nucleotides indicate the codon for aa 153). These two DNA fragments were linked and amplified using primers 321-S and 1517-AS. After that, the DNA fragment was digested with EcoRI/XbaI. This DNA insert was then cloned into pcDNA3, linearized previously with EcoRI/XbaI, to generate pcDNA3-CCDE1.

To clone the H1-coding sequence (aa 261–291), primers S (5' CGGGATCCCATATCGATCTGCTT 3') and AS (5' GGAATTCTTAAAACAGTTGACCAAC 3') were used. After amplification, the DNA fragment was digested with BamHI/EcoRI. This DNA insert was cloned into pGEX2T (Pharmacia), linearized previously with BamHI/EcoRI, to generate plasmid pGEX2T-E1H1.

To clone the H2-coding sequence (aa 329–383), primers S2 (5' GAAGATCTACGGCAGCGTTGGTG 3') and AS2 (5' GGAATTCTTACGCGTCGACGCCGGC 3') were used. After amplification, the DNA fragment was digested with BglII/EcoRI. This DNA insert was cloned into pGEX2T, linearized previously with BamHI/EcoRI, to generate plasmid pGEX2T-E1H2.

All expression plasmids derived from PCR were verified by sequencing.

{blacksquare} Proteins for making polyclonal antibodies.
The HCV capsid protein (aa 1–191) of HCV strain RH (Lo et al., 1995 ), a truncated capsid protein (aa 1–115) of an HCV strain isolated from Taiwan and a partial E1 protein (aa 192–328) of HCV strain RH were expressed separately in Escherichia coli. After expression, the capsid and E1 proteins were partially purified by SDS–PAGE on a 13% polyacrylamide gel. Proteins were eluted for immunization in rabbits.

{blacksquare} GST pull-down assay.
The GST pull-down assay was conducted using the Bulk GST Purification kit (Pharmacia Biotech), following the manufacturer's procedure. A sample of 10 µl of 35S-labelled, in vitro-translated protein was incubated with 2 µg purified GST or GST–capsid fusion protein in 500 µl PBS containing 4 mM PMSF and 0·5% Triton X-100 at room temperature. At 2 h after incubation, glutathione–Sepharose 4B gel slurry was added to the reaction mixture and incubation was continued for another 2 h. After washing, the precipitated products were analysed by SDS–PAGE on a 13% polyacrylamide gel. A sample of 1 µl of 35S-labelled, in vitro-translated protein was loaded as the input control.

{blacksquare} Proteinase K protection assay.
pcDNA3-CCE1 was used to express the HCV capsid and E1 proteins in vitro using the TNT Transcription·Translation system in the presence of canine pancreatic microsomal membranes (Promega). A sample of 0·5 µl of 35S-labelled, in vitro-translated protein was incubated with proteinase K (final concentration 30 µg/ml) on ice for 30 min. The incubation was stopped by adding 1 µl 200 mM PMSF. The protein samples were then analysed by SDS–PAGE on a 13% polyacrylamide gel. For the disruption of the membrane, the in vitro-translated protein was treated with 1% Triton X-100.

{blacksquare} Radioimmunoprecipitation.
Confluent HuH-7 (human hepatoma) cells maintained in Dulbecco’s modified Eagle’s medium containing 10% foetal calf serum, 100 µg/ml penicillin/streptomycin and 100 µg/ml non-essential amino acids (Gibco BRL) were infected with a recombinant vaccinia virus (vTf7-3) carrying the T7 phage RNA polymerase gene (Fuerst et al., 1986 ). At 2 h after infection, cells were transfected with 1 µg plasmid DNA using the Effectene Transfection reagent (Qiagen). At 18 h after transfection, cells were incubated in methionine-free medium for 2–3 h and were subsequently radiolabelled with [35S]methionine in the same medium (160 µCi/ml; 5·92 MBq/ml) for 1–2 h. Cells were lysed with 1 ml RIPA buffer (50 mM Tris–HCl, pH 7·5, 300 mM NaCl, 4 mM EDTA, 0·5% Triton X-100, 0·1% SDS and 0·5% sodium deoxycholate) and immunoprecipitated with either 2·5 µl rabbit anti-capsid and/or anti-E1 antibody. Protein samples were then analysed by SDS–PAGE on a 13% polyacrylamide gel.

{blacksquare} Immunofluorescence.
HuH-7 cells infected with the recombinant vaccinia virus and transfected with the expression plasmids were fixed with -20 °C acetone for 2 min. The rabbit anti-capsid or anti-E1 antibody, diluted 1:200 in PBS containing 0·05% NaN3, 0·02% saponin and 1% BSA, was used as the primary antibody. The secondary antibody used was FITC-conjugated goat anti-rabbit antibody.


   Results
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Abstract
Introduction
Methods
Results
Discussion
References
 
Glycosylated E1 protein can interact with capsid proteins of different length in vitro
Glycosylated E1 proteins were translated in vitro (with the signal sequence located in the C terminus of the capsid protein) in the presence of microsomal membranes. After translation, the membrane was disrupted with Triton X-100. The interaction of glycosylated E1 and capsid protein was analysed using a GST pull-down assay. As shown in Fig. 1(a), glycosylated E1 protein was not only pulled down by the 191 aa capsid protein (Fig. 1a, lane 4) but also by the 153 aa capsid protein (Fig. 1a, lane 3). The capsid protein binding affinities with E1 between these GST–capsid fusion proteins are similar. The GST–capsid fusion protein with 153 aa instead of 173 aa was used for the in vitro-binding assay due to the low level of expression of the 173 aa GST–capsid fusion protein in E. coli (data not shown).



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Fig. 1. Interaction between the HCV capsid and E1 proteins in vitro. (a) Glycosylated E1 protein can interact with different length capsid proteins. Lanes: 1, in vitro-translated E1 (with microsomal membranes); 2, pulled down by GST alone; 3, pulled down by GST-C153; 4, pulled down by GST-C191. (b) Non-glycosylated E1 protein can interact with different length capsid proteins. Lanes: 1, in vitro-translated E1 (to aa 383); 2, pulled down by GST alone; 3, pulled down by GST-C115; 4, pulled down by GST-C153. (c) Truncated E1 (to aa 340) can still interact with different length capsid proteins. Lanes: 1, in vitro-translated E1 (to aa 340); 2, pulled down by GST alone; 3, pulled down by GST-C115; 4, pulled down by GST-C153.

 
Non-glycosylated E1 protein can interact with capsid proteins of different length in vitro
Non-glycosylated E1 (aa 192–383) was translated in vitro in the absence of microsomal membranes. The interaction of non-glycosylated E1 and capsid protein was analysed using a GST pull-down assay. As shown in Fig. 1(b), non-glycosylated E1 protein was pulled down by the 153 aa capsid protein (Fig. 1b, lane 4). These data indicate that glycosylation does not affect the interaction between HCV capsid and E1 proteins in vitro. Serially truncated E1 proteins (to aa 380, 370, 360, 350 and 340) were also pulled down by the 153 aa capsid protein (data not shown, see also Fig. 1c, lane 4) with at least the same efficiency as that of full-length E1 protein. This observation suggests that the C terminus of E1 is not essential for binding to the capsid protein in vitro.

Possible E1 topology: proteinase K protection assay
Based on the hydrophobicity of the E1 protein, there are two hydrophobic domains (H1 and H2) (Takamizawa et al., 1991 ). H1 spans from aa 261 to 291, while H2 spans from aa 329 to 383. Based on amino acid hydrophobicity, there are two most likely models for the topology of the E1 protein (Fig. 2): the first is that a short peptide (aa 364–368) in the H2 domain is exposed in the cytosolic phase (model a) and the second is that the entire H2 domain is located in the membrane (model b). To study the topology of the E1 protein, we have performed previously trypsin-digestion protection assays in cells (Lo et al., 1996 ). These data indicated that the E1 protein resides mostly, if not entirely, in the endoplasmic reticulum (ER) membrane and/or lumen. To study whether there is a short peptide (aa 364–368) in the H2 domain exposed in the cytosolic phase, we have performed a proteinase K protection assay using in vitro-translated proteins. As shown in Fig. 3, the intact, glycosylated E1 protein was protected by microsomal membranes when treated with proteinase K (Fig. 3, lane 3), while the capsid protein was not. No obvious reduction in size of the E1 protein (e.g. a removal of 20 aa from the C terminus of E1) was observed. This result is similar to that of a previous report (Hijikata et al., 1991 ), suggesting that the topology of the E1 protein was likely to be that of model B. This implies that the E1 protein does not interact with the capsid protein in the cytosolic phase.



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Fig. 2. Two models for HCV E1 topology. One short peptide (aa 364–368) in the H2 domain is exposed in the cytosolic phase (model a) or the entire H2 domain is located in the ER membrane (model b).

 


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Fig. 3. Proteinase K protection assay. Lanes: 1, in vitro-translated capsid and E1 proteins in the presence of microsomal membranes; 2, treated with PBS; 3, treated with proteinase K; 4, treated with Triton X-100 and proteinase K.

 
The H1 domain of E1 plays a more important role than the H2 domain for capsid protein interaction in vitro
In order to determine which hydrophobic domain of the E1 protein is important for capsid protein interaction, various E1 protein mutants were constructed (Fig. 4a). The binding efficiencies of these E1 mutants were also analysed using a GST pull-down assay. As shown in Fig. 4(b), the binding efficiencies of various E1 mutants with the capsid protein were C328>C383>C328D>C383D. These experiments were done three times and the binding efficiencies of these E1 mutants were plotted in Fig. 4(c). The H1 domain of the E1 protein plays a more important role than the H2 domain in interacting with the capsid protein in vitro.



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Fig. 4. The H1 domain of the E1 protein is more important than the H2 domain in binding the capsid protein in vitro. (a) Construction of various E1 mutants. (b) Interaction of various E1 mutants with capsid protein. Lanes: 1, in vitro-translated C328; 2, C328 pulled down by GST alone; 3, C328 pulled down by GST-C153; 4, in vitro-translated C328D; 5, C328D pulled down by GST alone; 6, C328D pulled down by GST-C153; 7, in vitro-translated C383; 8, C383 pulled down by GST alone; 9, C383 pulled down by GST-C153; 10, in vitro-translated C383D; 11, C383D pulled down by GST alone; 12, C383D pulled down by GST-C153. (c) Binding efficiencies of various E1 mutants with capsid protein: C328>C383>C328D>C383D. The binding efficiency of each E1 mutant was calculated using the following formula: (signal pulled down by GST-C153)-(signal pulled down by GST)/signal pulled down by GST. Comparison of binding efficiencies of these E1 mutants was done using C383D as one basic unit in each individual experiment. Data are from three different experiments.

 
Neither the H1 nor the H2 domain of E1 is stable when expressed as a GST fusion protein in E. coli
In order to determine the binding affinity between the H1 domain (or the H2 domain) of the E1 and capsid proteins directly, the H1 domain (or H2 domain) of the E1 protein has been expressed as a GST fusion protein in E. coli. However, neither the H1 nor the H2 domain of E1 is stable (data not shown).

The E1 protein without the H1 domain could not interact well with the capsid protein in vivo
We have shown previously that the E1 protein without the H2 domain could not interact with the capsid protein in cells (Lo et al., 1996 ). This may be due to the fact that the H2 domain is the retention signal of the E1 protein in the ER (Cocquerel et al., 1999 ). To determine the importance of the H1 domain in binding in vivo, the capsid protein and the E1 protein without the H1 domain were expressed together in HuH-7 cells. Similar to the subcellular localization of the E1 protein and the E1 protein without the H2 domain (Fig. 5a, b), the E1 protein without the H1 domain stains as a perinuclear protein (Fig. 5c). In the immunoprecipitation experiment, unlike the complete E1 protein (Fig. 6b, lane 2), the E1 protein lacking the H1 domain could not be co-immunoprecipitated with anti-capsid antibodies when in the presence of the capsid protein (Fig. 6a, b, lanes 2 and 3, respectively).



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Fig. 5. Subcellular localization of various HCV E1 and capsid mutants. HuH-7 cells were transfected with pcDNA3-CCE1 (a), pcDNA3-E1 350 (b), pcDNA3-CCE1D (c) or pcDNA3-CCDE1 (d). After fixation, the cells were stained with rabbit anti-E1 (a–c) or anti-capsid (d) antibodies and FITC-conjugated goat anti-rabbit secondary antibody.

 


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Fig. 6. E1 protein with but not without the H1 domain could be co-immunoprecipitated by anti-capsid antibody in the presence of the capsid protein. (a) HuH-7 cells were transfected with pcDNA3 (lanes 1 and 3) or pcDNA3-CCE1D (lanes 2 and 4). After labelling, proteins were precipitated with rabbit anti-capsid antibody (lanes 1 and 2) or with both anti-capsid and anti-E1 antibodies. (b) HuH-7 cells were transfected with pcDNA3 (lane 1), pcDNA3-CCE1 (lane 2) or pcDNA3-CCE1D (lane 3). After labelling, proteins were precipitated with rabbit anti-capsid antibody.

 
The capsid protein without the hydrophobic domain is labile and located in the nucleus
Due to the topology of the E1 protein, it is reasonable to assume that the hydrophobic domain of the capsid protein (aa 119–152) is involved in interaction with the E1 protein. To prove this assumption, the capsid protein without the hydrophobic domain was expressed with the downstream E1 protein. However, unlike the intact capsid protein (Fig. 7, lane 2), the capsid protein lacking the hydrophobic domain is almost undetectable by Western blot analysis (Fig. 7, lane 3), while both expression plasmids with the same expression level of the downstream E1 protein were detectable. Therefore, in comparison to the intact capsid protein, the capsid protein without the hydrophobic domain is labile. On the other hand, in some instances, the majority of the capsid protein without the hydrophobic domain could be detected in the nucleus by immunofluorescence (Fig. 5d).



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Fig. 7. Protein analysis by Western blotting. HuH-7 cells were transfected with pcDNA3 (lane 1), pcDNA3-CCE1 (lane 2) or pcDNA3-CCDE1 (lane 3). After transfection, total proteins were extracted, separated by SDS–PAGE on a 13% polyacrylamide gel and analysed by Western blotting. Rabbit anti-capsid and anti-E1 antibodies were used.

 
We have also performed the in vitro GST pull-down assay to verify the importance of the capsid protein hydrophobic domain in binding with serial E1 proteins (Fig. 1b, c, lanes 3). The capsid protein without the hydrophobic domain (aa 1–115) could not interact with the E1 protein as well as the complete capsid protein with the hydrophobic domain could (Fig. 1b, c).


   Discussion
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Abstract
Introduction
Methods
Results
Discussion
References
 
Because HCV is an enveloped virus, its maturation probably requires the interaction between its capsid and envelope proteins. The interaction between HCV capsid and E1 envelope proteins has been demonstrated in CV-1 cells (Lo et al., 1996 ) and in insect cells (Baumert et al., 1998 ). Results from the proteinase K protection assay (Fig. 3) indicate that the entire H2 domain is located in the membrane, which is in agreement with the model proposed by others (Op De Beeck et al., 2001 ). It also implies that the E1 protein should interact with the capsid protein in the ER membrane.

There are two hydrophobic domains in the E1 protein based on its hydrophobicity (Takamizawa et al., 1991 ): H1 (aa 261–291) and H2 (aa 329–383). Based on the topology of E1 (Fig. 2b), the E1 protein without these two hydrophobic domains would not interact with capsid protein in cells. The interaction between the capsid and the E1 protein without these two hydrophobic domains (C328D) in vitro, which would not likely occur in cells, is due to the involvement of the E1 protein sequence exposed in the ER lumen (Fig. 4b). Therefore, binding between the capsid and the E1 proteins in vitro would not be the same as that in vivo. However, the comparison of the binding efficiencies of various E1 mutant proteins with the capsid protein in vitro could still identify sequences of the E1 protein that are important for the interaction with capsid protein.

In the in vitro GST pull-down assay (Fig. 4b, c), the E1 protein lacking the H2 domain is better than that with the H2 domain (C328>C383 and C328D>C383D) in binding the capsid protein. This suggests that the H2 domain may interfere with the interaction between the capsid and the E1 proteins. This is supported by the observation that serially truncated E1 proteins (to aa 370, 360, 340 and 328) could still interact with the capsid protein with at least the same affinity as that of full-length E1 in vitro (Figs 1 and 4). On the other hand, the E1 protein with the H1 domain is better than that without H1 domain (C383>C383D and C328>C328D) in binding the capsid protein. Therefore, the H1 domain of the E1 protein is more important than the H2 domain in binding the capsid protein in vitro. These data do not conflict with our previous finding that the H2 domain is essential for the interaction of the E1 protein with the capsid protein in cells (Lo et al., 1996 ), because the H2 domain is the retention signal for the E1 protein in the ER membrane (Cocquerel et al., 1999 ). To verify that the H1 domain of the E1 protein is indeed important for binding the capsid protein, the HCV capsid protein was co-expressed with downstream E1 protein without the H1 domain (E1D) in cells. The immunoprecipitation assay (Fig. 6) showed that without the H1 domain, the E1 protein could not bind well with the capsid protein.

If the interaction between the HCV capsid and the E1 proteins occurs in the ER membrane and the H1 domain of the E1 protein is important for this interaction, it is reasonable to assume that the hydrophobic domain (aa 119–152) of the capsid protein is involved in this interaction as well. It is difficult to prove this assumption because the capsid protein without this hydrophobic domain is relatively labile (Fig. 7). Furthermore, in some instances, the majority of the capsid proteins lacking the hydrophobic domain stain as nuclear proteins (Fig. 5d). This argues for the importance of this hydrophobic domain (aa 119–152) in the association of capsid protein with the ER membrane. The capsid protein without this hydrophobic domain (C115) could not interact with serial E1 deletion proteins in vitro to the same efficiency as capsid protein with this hydrophobic domain (C153) (Fig. 1b, c). This argues for the importance of this hydrophobic domain in binding with E1 protein.

In other flaviviruses, the hydrophobic C-termini of both prM and E proteins are highly conserved and could be involved in envelope–nucleocapsid interactions (Rice, 1996 ). However, the H1 domain of the HCV E1 protein is not the most conserved region in this protein (Bukh et al., 1993 ). Nor is the hydrophobic domain of the capsid protein the most conserved region in this protein (Bukh et al., 1994 ). It may imply that the interaction between the HCV capsid and the E1 proteins depends on hydrophobic interactions but not sequence-specific interactions.

It has been demonstrated that the correct folding of the HCV E1 protein depends on the presence of the E2 protein (Deleersnyder et al., 1997 ). Therefore, the interaction between the E1 and the capsid proteins is not dependent on conformation. In addition to being the signals for ER retention, the C-terminal hydrophobic domains of E1 and E2 are also responsible for E1/E2 heterodimerization (Charloteaux et al., 2002 ; Op De Beeck et al., 2001 ). Our data showed that the H1 domain of the E1 protein is important in binding with the capsid protein in vitro and that the H2 domain may interfere with this interaction (Fig. 4). During the assembly of HCV particles, it is possible that the E2 protein helps the correct folding of the E1 protein by interacting with the H2 domain and subsequently facilitating the H1 domain to interact with the capsid protein.

Based on these data, a possible model for the E1 protein was proposed in Fig. 8. This model explains why both hydrophobic domains of the HCV E1 proteins are required for the interaction with capsid protein in cells. The H2 domain is the retention signal for the E1 protein in the ER membrane, while the H1 domain is involved in binding with capsid protein.



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Fig. 8. Proposed model for the HCV E1 protein: the H2 domain is the retention signal for the E1 protein in the ER membrane, while the H1 domain is involved in binding with capsid protein.

 

   Acknowledgments
 
We thank Dr J.-H. Ou and Dr M.M.C. Lai for providing expression plasmids. This work has been supported by a grant from the National Science Council of Taiwan (NSC 90-2320-B-320-014).


   References
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
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Received 1 March 2002; accepted 23 July 2002.