Australian Centre for Hepatitis Virology, Macfarlane Burnet Institute for Medical Research and Public Health, Yarra Bend Road, Fairfield 3078, Victoria, Australia1
Author for correspondence: Elizabeth Grgacic. Fax +61 3 9282 2100. e-mail grgacic{at}burnet.edu.au
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Abstract |
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Introduction |
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The assembly of the envelope proteins and their involvement in entry of the virus are closely linked to a unique protein transport process adopted by the hepadnaviruses. This ability of the virus to translocate a large N-terminal portion of the fully translated L protein, and to limit this process to achieve mixed membrane orientations or topologies, are underlying features of hepadnavirus assembly and regulation. The L protein displays a more complex functional role than that expected of a viral structural protein and this is partly achieved by its mixed topology. The orientation of viral envelope proteins spanning the endoplasmic reticulum (ER) membrane is mirrored in the mature particle, so that those domains located in the ER lumen are found on the ectodomain of the particle and cytosolic domains remain internally disposed. Thus, the translocated form of L makes pre-S sequences available on the external surface of the mature virion for receptor binding (Klingmuller & Schaller, 1993 ; Le Seyec et al., 1999
), while maintenance of an internal pre-S domain enables the L protein to take on the suggested role of a matrix protein for interaction with the nucleocapsid (Bruss et al., 1994
) (Fig. 1A
). The L protein also controls the size of the pool of the replicative template, covalently closed circular DNA (cccDNA), at least partly by determining the assembly of core particles into virions and perhaps also by a more direct regulatory role (Summers et al., 1991
).
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Increasingly, TM domains are being identified as important structural elements in membrane protein assembly and for many enveloped viruses, budding is dependent on the formation of an envelope lattice through lateral interactions of the TM domains (Garoff et al., 1998 ). Lateral interactions may occur through hydrogen bonding of the side groups of polar residues, such as asparagine, arginine and glutamic acid, or by helix packing through the regular meshing of side chains, as exemplified by heptad repeats or leucine zipper motifs (Gurezka et al., 1999
; Ubarretxena-Belandia & Engelman, 2001
). The latter have been implicated in the folding and/or oligomerization of a variety of cellular and viral membrane proteins (Gurezka et al., 1999
), including gp41 of human immunodeficiency virus type 1 (Center et al., 1997
). Assembly of the HBV envelope involves the accumulation of S monomers along the ER, where initial contacts may involve such lateral TM interactions before they bud into the lumen and are stabilized by disulphide bonds into dimers, eventually forming higher-order oligomers. However, of the 14 cysteine residues in HBV, only the three conserved cysteines of the first hydrophilic loop are essential for particle secretion and these do not form intermolecular disulphide bonds (Mangold & Streeck, 1993
). That the DHBV envelope contains only these three cysteines further points to TM helices playing a role in the initial stages of hepadnavirus assembly.
The aim of this study was to investigate whether the transmembrane helices of the S protein play a role in translocation and morphogenesis. Through mutagenesis using constructs expressing the S and L envelope proteins independently, we have identified TM1 as an essential structural determinant in S, with charged, polar residues contributing to L protein translocation and particle assembly.
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Methods |
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Cells and transfections.
Chicken hepatocyte LMH cells (Leghorn male hepatoma) were maintained in Dulbeccos modified Eagles medium (DMEM-F12) supplemented with 10% foetal bovine serum. Transfections were carried out by the dextran sulphate method, as previously described (Grgacic et al., 1998 ) using 5 µg of DNA per well in six-well multiplates (Greiner).
Protease protection analysis.
Microsomes were prepared according to the method of Prange & Streeck (1995) with modifications. Transfected LMH cells (two 30 mm diameter wells) were washed in cold Tris-buffered saline (TBS; 50 mM TrisHCl, pH 7·5,150 mM NaCl). The monolayers in each well were incubated on ice with 400 µl 0·1x TBS for 10 min and then harvested by scraping, pooled and dispersed by drawing five times through a 26G needle. The homogenate was adjusted to 1x TBS with 5x TBS and centrifuged for 20 min at 2500 r.p.m. at 4 °C to remove unbroken cells and nuclei. The supernatant was removed and set aside while the pellet was again dispersed in 300 µl TBS and centrifuged as before. Supernatants were pooled and layered on to 2·7 ml 250 mM sucrose in TBS and centrifuged for 30 min at 38000 r.p.m. at 4 °C in an SW60 rotor (Beckman). The microsomal pellets were washed once with TBS and resuspended in 65 µl TBS. For trypsin protection analysis, the microsomal preparation was divided into three 20 µl aliquots. One sample was left untreated while the remaining two were treated with 25 µg/ml of trypsin (TPCK treated; Worthington Biochemical Corporation) with or without 0·5% NP-40 for 1 h on ice. Proteolysis was halted by the addition of 30 µg/ml aprotinin (Boehringer) and further incubated on ice for 20 min. Five µl of 5x Laemmli buffer was then added to each sample and boiled for 5 min prior to separation by 13% SDSPAGE followed by Western blotting to detect the L protein.
Western blot analysis.
Proteins were separated by 13% SDSPAGE and transferred to nitrocellulose membrane (Schleicher and Schüll) using a Trans-Blot SD semi-dry transfer cell (Bio-Rad). Membranes were blocked for 1 h with 3% skim milk in PBS plus 0·3% Tween 20 (PBST). Membranes were probed with monoclonal anti-S (7C.12) and anti-pre-S (1H.1) (Pugh et al., 1995 ) for 1 h in 1% skim milk in PBST, then washed with PBST and probed with goat anti-mouse Ighorseradish peroxidase (Amersham) in 1% skim milk in PBST. After a final wash in PBST (3x10 min), protein bands were visualized by enhanced chemiluminescence (ECL) (Amersham).
Pulsechase analysis and carbonate extraction.
Transfected LMH cells were starved in methionine/cysteine-free media (ICN) for 30 min before pulse-labelling with 150 µCi/ml TRAN35S-label (ICN) for 30 min. Cells were harvested immediately after labelling and after a 4 h and 24 h chase with unlabelled media. A crude microsome preparation was prepared as described above except that the ultracentrifugation through sucrose was omitted. Microsomes were extracted with 0·1 M Na2CO3 according to the method of Bruss & Ganem (1991b ) with modifications. Briefly, 200 µl microsome fractions were treated with 4 ml 0·1 M Na2CO3, pH 11·5, on ice for 30 min. The sample was then ultracentrifuged for 30 min at 100000 g using an SW60 rotor. The top 3·5 ml of supernatant was removed, the next 0·5 ml discarded and the remaining 200 µl was regarded as the pellet. The supernatant and the pellet were neutralized with acetic acid and adjusted to 1x RIPA buffer with 10x RIPA buffer (100 mM Tris, 2·5 M NaCl, 10 mM EDTA, 10% NP-40, 5% sodium deoxycholate, 1% SDS), and labelled envelope proteins were immunoprecipitated with monoclonal anti-S (7C.12) (Pugh et al., 1995
) at 4 °C for 16 h. Immune complexes were pelleted following incubation with 60 µl of a 10% slurry of Protein ASepharose for 1 h at 4 °C, washed twice with 1x RIPA buffer and boiled in Laemmli buffer for SDSPAGE and autoradiography.
Isolation of intracellular and extracellular particles.
For extracellular particles, media from transfected LMH cells was harvested 3 days post-transfection and clarified of non-adherent cells by centrifugation for 5 min at 2000 r.p.m.
For intracellular particles, cell monolayers were washed twice with PBS and harvested by scaping cells into 1 ml PBS. Harvested cells were freeze/thawed three times with vigorous vortexing upon thawing. The cytosol fraction (supernatant) was obtained by centrifugation for 1 min at 10000 r.p.m. in an Eppendorf centrifuge. This procedure has been used in this laboratory to release DHBV particles capable of infecting primary duck hepatocytes from transfected cells. Particles in the clarified media or cytosol fraction were diluted to 6 ml with PBS and pelleted for 3 h at 38000 r.p.m. in an SW40 rotor through 3 ml of 20% sucrose on to a 2 ml 70% sucrose cushion. The fraction at the 2070% interface was collected from the bottom and methanol-precipitated for 16 h at -20 °C, followed by separation by 13% SDSPAGE and analysis by Western blotting.
For the cell membrane fraction, following removal of the cytosol fraction the pellet was resuspended in 100 µl PBS with 1% NP-40 by vigorous vortexing and centrifuged for 1 min at 10000 r.p.m. in an Eppendorf centrifuge. The supernatant or membrane fraction was removed and 20 µl was mixed with 5 µl 5x Laemmli buffer for SDSPAGE.
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Results |
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L chains expressed in the presence of S in microsomal vesicles were protected from protease digestion and deemed to have been translocated across the ER membrane to the lumen (Fig. 1C, L/S), while L chains expressed without S were susceptible to digestion (Fig. 1C
, L
).
To assess the role of the transmembrane domains of S in L translocation, L+S- was co-transfected with a construct only able to synthesize S (L-S+) in which various deletions of the amphipathic TM domains were made (Fig. 2A). Microsomes were prepared from co-transfected LMH cells, treated with trypsin, and protection of L was assessed by Western blotting. TM1 appeared to be a critical structural element of the S protein, since deletion of this domain resulted in little or no detectable S expression, most likely due to aberrant protein folding and degradation. As a consequence, L protein was not protected from trypsin digestion in the absence of detergent (Fig. 2B
,
TM1). In contrast, S protein with a deletion in the region encompassing TM3 was tolerated and still able to protect L protein from protease attack (Fig. 2B
,
TM3).
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Discussion |
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Using constructs that express L and S independently, we have demonstrated that envelope protein functional diversity is not limited to the pre-S domain and the mixed topologies it achieves, but that TM1, a common domain of L and S, appears to have different functional roles in L and S. This study has indicated that TM1 in S is essential for proper S folding and stability. Similar studies on HBV S protein have shown that although TM1 was not essential for membrane insertion, it played a role in assembly either directly or indirectly (Bruss & Ganem, 1991a ; Prange et al., 1992
). While this deletion in DHBV S had a more drastic effect than previously observed in HBs, together these results and the apparent non-essential role of TM3 strongly pointed to TM1 as the region engaged in envelope interaction. In contrast, deletion of TM1 in L was well tolerated, allowing L to be membrane-inserted, translocated to the ER lumen (Fig. 3B
) and secreted with S in particles (data not shown). Thus, TM1 of L appears to be dispensable for the translocation/assembly process. Previously, we have shown that low pH treatment of particles released a hidden, potentially fusogenic, hydrophobic domain, identified as TM1 (Grgacic & Schaller, 2000
). These results and studies with synthetic peptides of TM1 implicate this domain of L in the viral fusion process (Rodriguez-Crespo et al., 1999
). We conclude that TM1 of L has a different functional role to that of S and, importantly, that the way L is incorporated into the particles must differ from S.
TM1 of S was shown to have two important structural determinants for particle assembly: (i) a leucine zipper-like motif on one face of the -helix, and (ii) two charged, polar residues, lysine 24 and glutamic acid 27, on the opposing face. Both these features were shown to play a role in the ability of S to facilitate L translocation as determined by the protease protection assay. Multiple mutations in the leucine zipper-like motif resulted in defects in L translocation and particle export, suggestive of aberrant particle formation. It is not clear whether L translocation is a prerequisite for particle secretion or whether a more global change had occurred in the envelope, which inhibited export. The partial glycosylation of these leucine zipper mutants implied such a change in S occurred, since the glycosylation site is located adjacent to TM2 well downstream of TM1. However, these changes were not sufficient to inhibit particle assembly.
The lack of particle assembly of the K24A/E27A S mutant suggests there is a relationship between assembly of S into particles and its ability to translocate L. Polar residues are unusual in transmembrane domains: the energetic cost of their insertion into a hydrophobic environment usually heralds some specific functional role (Bonifacino et al., 1991 ; Cocquerel et al., 2000
; Ubarretxena-Belandia & Engelman, 2001
). If exposed, they are predicted to be involved in inter-helical hydrogen bonding or, if charged, to form salt bridges with similar anti-parallel domains. Previously it has been predicted that the TM helices had the potential to form a proteinaceous channel for L translocation (Guo & Pugh, 1997
; Stirk et al., 1992
), and the data presented here would suggest that inter-helical bonding between polar residues of TM1 could facilitate this. Alternatively, or in addition, involvement of the charged residues may be through interaction of the helix with regions of the membrane-embedded loop (Grgacic et al., 2000
) (Fig. 1A
), which also contains several charged residues and may form the hydrophilic lining of the channel. One could further speculate that the same residues in L will necessarily be sequestered to stop unwanted helix interactions, again suggestive of a different conformational arrangement from S and possibly also accounting for the binding of Hsc70 chaperone to L (Hildt, 1997
; Prange et al., 1999
).
Although this study identified a link between particle assembly and L translocation, no direct interaction (dimerization) between the envelope proteins could be demonstrated in the ER membrane, either by chemical cross-linking with a membrane permeable cross-linker or by velocity sedimentation of envelope containing cell lysates (data not shown). This may be due to the technical difficulty in capturing non-covalent envelope interactions.
None the less, the results from this study provide further supportive evidence that L translocation in DHBV involves correct assembly of S into particles. These data, together with the demonstration of an intermediate L topology in DHBV, i.e. one where the pre-S domain is not fully translocated to the ER lumen but traverses the viral envelope in the mature particle, support the model that the envelope proteins assemble to form their own translocation channel. An envelope-formed channel would allow the long hydrophilic pre-S domain to not only traverse the viral envelope but also be retained in that topology in the mature particle for subsequent release by such conditions as low pH, as previously described (Grgacic & Schaller, 2000 ; Guo & Pugh, 1997
). Whether an envelope complex of this nature would still need, or be physically able, to engage with the host cell translocation machinery is not clear.
The mechanism of DHBV L translocation appears to be in contrast with recent studies in HBV (Lambert & Prange, 2001 ), which indicate that S is not required for L translocation. This difference in the translocation mechanism from DHBV may reflect use of a different folding pathway, indicated by the dominant retention of HBV L in the ER in the presence of excess S whereby L is largely excluded from subviral particles (SVPs) and cytosolic L is retained for virion formation (Gazina et al., 1998
). In contrast, DHBV L is not actively excluded from SVPs, which thus contain the same ratio of L:S as virions (Schlicht et al., 1987
). This difference in envelope folding between the human and avian hepadnaviruses is further supported by the inability of the avian envelope proteins to combine with mammalian HBs to form particles, while woodchuck and human HBs are readily interchangeable (Gerhardt & Bruss, 1995
). An important question still remains with both the S-dependent (DHBV) and S-independent (HBV) modes of L translocation and that is how the translocation is regulated to 50% of L chains to meet the essential, dual pre-S roles of capsid assembly and receptor interaction. For DHBV at least it may be a result of the physical constraints conferred by the number of S chains required for each translocating L chain.
While this essential translocation mechanism appears to differ between the avian and mammalian viruses, the strong conservation of the N-terminal third of the S domain and two polar residues at positions 24 and 27 of TM1 for all hepadnavirus envelope proteins points to this being a common determinant in particle morphogenesis (Fig. 4C). Moreover, the results of this study indicate that TM1 has different functional roles in L and S, which suggest that the assembly and incorporation of L into the viral envelope lattice may differ markedly from S, despite their common membrane-spanning domains.
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Acknowledgments |
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References |
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Received 29 January 2002;
accepted 26 February 2002.