Department of Virology, Room 333, The WrightFleming Institute, Faculty of Medicine, Imperial College of Science, Technology & Medicine, St Marys Campus, Norfolk Place, London W2 1PG, UK1
Author for correspondence: Geoffrey L. Smith. Fax +44 207 594 3973. e-mail glsmith{at}ic.ac.uk
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Abstract |
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Introduction |
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An overview of morphogenesis |
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Recognition of intracellular and extracellular virus |
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An important conclusion from these studies, and one pertinent to the current development of second generation smallpox vaccines, was that tests to measure neutralizing antibody that are relevant to immunological protection should utilize EEV rather than IMV (Boulter & Appleyard, 1973 ; Appleyard & Andrews, 1974
). The EEV neutralization test is difficult because of the presence of contaminating IMV in EEV preparations and the fragility of the EEV outer envelope (Boulter & Appleyard, 1973
). However, Appleyard et al. (1971)
described two methods for measuring antibody to EEV: (i) the anti-comet test; and (ii) the modified neutralization test using EEV pretreated with antibody against inactivated IMV. Some strains of VV [such as rabbitpox and International Health Department (IHD)-J] release high levels of EEV and if these viruses are allowed to grow on cell monolayers they give rise to characteristic comet-shaped plaques in which the head of the comet represents the primary plaque and the comet tails represents secondary plaques caused by unidirectional spread of EEV by convection currents (Law et al., 2002
). The formation of comets is inhibited by antibody to EEV but not IMV.
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IMV formation |
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The nature and origin of the crescents are disputed. Early investigators proposed that these were formed from a single lipid bilayer that was synthesized de novo and lacked continuity with cellular membranes (Dales, 1963 ; Dales & Mosbach, 1968
). Subsequently, it was proposed that the crescent was composed of a pair of tightly apposed membranes that were derived from and were continuous with cell membranes of the intermediate compartment (IC) between the endoplasmic reticulum (ER) and the Golgi stack (Sodeik et al., 1993
). Another study reported no continuity between virus and cellular membranes and only a single lipid bilayer (Hollinshead et al., 1999
). Recently, additional reports claimed IMV has two (Risco et al., 2002
) or more membranes (Griffiths et al., 2001
). The de novo model of membrane biosynthesis contradicts dogma stating that membranes grow from existing membranes. However, a single membrane around the outside of IMV simplifies the virus re-entry mechanism (see below). In contrast, the double membrane model fits with our knowledge of cell biology, but creates a topological difficulty during virus re-entry: namely, how the multiple membranes surrounding the virus are shed to release the core into the cytosol. The issue is fundamental to aspects of virus morphogenesis and re-entry and additional study is needed. This review considers events after IMV formation and builds on an earlier review (Smith & Vanderplasschen, 1998
).
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Egress of IMV from factories |
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The movement of IMV particles from the factory to the wrapping membranes requires microtubules and the A27L protein (Sanderson et al., 2000 ), which is present on the IMV surface and is a target for antibodies that neutralize IMV infectivity (Rodriguez et al., 1985
). Repression of A27L gene expression caused a deficiency in IEV formation, a small plaque size and 20-fold reduced EEV production (Rodriguez & Smith, 1990
). The A27L protein is required for both transport and wrapping since loss of A27L prevented IMV transport and an Ala-25 to Asp substitution permitted transport but wrapping was inhibited (Sanderson et al., 2000
). This multi-functional protein also forms a complex with two other IMV proteins (A17L and A14L) (Rodriguez et al., 1993
) and promotes cell-to-cell fusion (Rodriguez et al., 1987
).
In another study virus particles were found to accumulate near the microtubule organizing centre (MTOC) and this accumulation was prevented by disruption of microtubules by nocodazole or by expressing dominant negative mutants of p50/dynamitin, which disrupts the function of dyneindynactin (Ploubidou et al., 2000 ). These observations supported the requirement for microtubules for IMV transport. Later during infection the MTOC was disrupted (Ploubidou et al., 2000
).
Late during infection, a greater proportion of IMV particles remains unwrapped and may either stay in the cytosol until cell lysis, bud through the plasma membrane or become occluded in ATIs. ATIs are proteinaceous bodies that appear late in infection (Ichihashi et al., 1971 ) and are composed predominantly of a single polypeptide (160 kDa in cowpox virus) (Patel et al., 1986
). The majority of orthopoxviruses, including VV, do not make ATIs because the gene encoding the 160 kDa protein is disrupted. However, several strains of cowpox virus and raccoonpox virus make ATIs (Ichihashi et al., 1971
; Patel et al., 1986
). The ATI enhances IMV stability after cell death and aids virus transmission between hosts.
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Proteins of IEV, CEV and EEV |
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In addition, genes A36R (Parkinson & Smith, 1994 ; van Eijl et al., 2000
) and F12L (Zhang et al., 2000
; van Eijl et al., 2002
) encode proteins that are present on IEV. Although some A36R and F12L proteins co-purify with EEV preparations, immunoelectron microscopy showed that they are absent from CEV and EEV envelopes (van Eijl et al., 2000
, 2002
). These proteins facilitate egress of IEV on microtubules (F12L) or CEV by actin polymerization (A36R) and therefore are termed transport proteins.
An interesting feature of the proteins encoded by these genes is that A33R, A36R, A56R, B5R and F13L proteins are palmitoylated (Grosenbach et al., 2000 ). Expression of most of these proteins individually by Semliki Forest virus vectors enabled the location of each protein to be studied in the absence of other VV proteins. The B5R, F13L and A34R proteins were present in intracellular vesicles, whereas the A33R and A56R proteins accumulated at the cell surface (Lorenzo et al., 2000
).
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Wrapping of IMV to make IEV |
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Early during infection the majority of IMV particles are wrapped to form IEV, whereas later during infection IMV predominate (Ulaeto et al., 1996 ), possibly due to depletion of wrapping membranes. The interaction of IMV with the wrapping membranes involves the cytosolic face of the wrapping membrane and the surface of IMV. The A27L protein on the IMV surface is implicated in this interaction (Sanderson et al., 2000
); however, no direct physical evidence for this has been published. Glycoproteins A33R, A34R, A56R and B5R have the majority of their polypeptide chains positioned within the lumen of the wrapping membranes and have only a short tail (1530 amino acids) in the cytosol (Fig. 3
), whereas the non-glycosylated proteins A36R, F12L and F13L have the majority of their polypeptide chain in the cytosol and so are better placed to interact with IMV. Interestingly, proteins F12L and A36R are associated predominantly with the outer IEV membrane such that after fusion of IEV with the plasma membrane they are absent from CEV (van Eijl et al., 2000
, 2002
). How they are excluded from the inner IEV wrapping membrane is unclear. One possibility is that during the progressive wrapping of IMV, A36R and F12L become displaced due to the bulk of their polypeptide chain being between the IMV surface and the wrapping membrane.
Analysis of virus mutants lacking individual genes has shown that the F13L (Blasco & Moss, 1991 ) and B5R (Engelstad & Smith, 1993
; Wolffe et al., 1993
) proteins are each required for efficient wrapping, whereas F12L, A33R, A34R, A36R and A56R are not (Table 1
). Without A34R there is an increased production of EEV yet fewer IEV are seen (Duncan & Smith, 1992
; Wolffe et al., 1997
; Law et al., 2002
).
Wrapping of IMV is inhibited by a drug, N1-isonicotinoyl-N2-3-methyl-4-chlorobenzoylhydrazine (IMCBH) (Kato et al., 1969 ; Payne & Kristensson, 1979
; Hiller et al., 1981a
), that prevents targeting of the F13L protein to the wrapping membranes (Hiller et al., 1981a
). Passage of VV in the presence of IMCBH resulted in generation of drug-resistant virus containing an Asp to Tyr mutation within the F13L protein (Schmutz et al., 1991
). The F13L protein is modified by acylation (palmitic and oleic acid) (Hiller & Weber, 1985
; Child & Hruby, 1992
; Payne, 1992
). Mutation of Cys-185 and Cys-186 to serine prevented palmitoylation leaving the F13L protein soluble in the cytoplasm and preventing wrapping (Grosenbach et al., 1997
; Grosenbach & Hruby, 1998
; Grosenbach et al., 2000
).
An interesting feature of the F13L protein is its limited amino acid similarity to phospholipase D (PLD) (Koonin, 1996 ; Ponting & Kerr, 1996
). Although no PLD activity was detected in cells expressing F13L, mutagenesis of a motif conserved in PLDs disrupted F13L function and only tiny plaques were formed (Sung et al., 1997
). Others reported that F13L is a broad specificity lipase with phospholipase C, phospholipase A and triacylglycerol lipase activity (Baek et al., 1997
). Mammalian PLD1 is expressed in the Golgi membranes and regulates vesicular budding (Bednarek et al., 1996
; Colley et al., 1997
). A similar role has been suggested for F13L based on the observation that F13L expressed without other VV proteins localizes in the Golgi (Lorenzo et al., 2000
) or post-Golgi vesicles (Husain & Moss, 2001
) and that F13L causes redistribution of B5R from the TGN to endosomal membranes unless the conserved PLD motif is mutated (Husain & Moss, 2001
). Consistent with a requirement for PLD activity in VV morphogenesis, the PLD inhibitor butanol-1 inhibited VV morphogenesis but expression of cellular PLD could not substitute for loss of the F13L protein (Husain & Moss, 2002
).
The other protein required for wrapping of IMV is B5R. This has four short consensus repeats (SCR) that are characteristic of regulators of complement activation (Takahashi-Nishimaki et al., 1991 ; Engelstad et al., 1992
) (Fig. 2
). The N-terminal signal peptide is proteolytically removed (Isaacs et al., 1992
) and some of the protein is also cleaved near the transmembrane domain to produce a secreted 35 kDa protein of unknown function (Martinez-Pomares et al., 1993
). The B5R protein is acylated (Payne, 1992
) by addition of palmitic acid at Cys-301, and possibly a second unidentified site (Grosenbach et al., 2000
), and forms higher molecular mass complexes in the absence of reducing agent (Engelstad et al., 1992
; Payne, 1992
). The B5R protein affects virus host-range in some cell types (Takahashi-Nishimaki et al., 1991
; Martinez-Pomares et al., 1993
).
Virus mutants lacking B5R are very inefficient at wrapping IMV to IEV, have 5- to 10-fold lower levels of EEV, form a small plaque and are attenuated in vivo (Takahashi-Nishimaki et al., 1991 ; Engelstad & Smith, 1993
; Martinez-Pomares et al., 1993
; Wolffe et al., 1993
) (Table 1
). The signals necessary for correct targeting of B5R to the Golgi membranes reside in the transmembrane/cytoplasmic tail since fusion of these domains to other proteins such as human immunodeficiency virus gp120 (Katz et al., 1997
), green fluorescent protein (GFP) (Ward & Moss, 2000
; Hollinshead et al., 2001
; Rodger & Smith, 2002
) or vesicular stomatitis virus G protein (Ward & Moss, 2000
) directed these chimaeras to IEV and EEV. Deletion of one or more SCR domains impaired wrapping of IMV and caused a small plaque phenotype, but EEV production was enhanced 10- to 50-fold (Herrera et al., 1998
; Mathew et al., 1998
; Rodger & Smith, 2002
). Loss of the cytoplasmic tail did not affect wrapping or reduce plaque size (Lorenzo et al., 1998
; Mathew et al., 2001
), but the protein was less rapidly transported through the exocytic pathway (Mathew et al., 2001
). Another study reported reduced accumulation of the C-terminally truncated protein in the Golgi membranes and concluded that the cytoplasmic tail had a role in retrieving B5R from the plasma membrane (Ward & Moss, 2000
). Addition of an ER retrieval sequence to the C terminus of B5R caused the relocation of B5R to the ER and a reduced plaque size, but did not prevent IEV and EEV formation (Mathew et al., 1999
).
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Transport of IEV to the cell surface |
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The movement of a large virion (250x350 nm) through the cytoplasm by diffusion is a slow and inefficient process (Sodeik, 2000 ) and so to expedite egress, VV uses cellular transport pathways. Two mechanisms have been proposed. One was that IEV particles induce polymerization of actin to drive these virions to the cell surface (Cudmore et al., 1995
, 1996
; Frischknecht et al., 1999b
). This proposal was consistent with the prior observations that VV-infected cells produce numerous actin bundles (diameter 0·3 µm) resembling filopodia or specialized microvilli with enveloped virions at their tip (Stokes, 1976
; Hiller et al., 1979
, 1981b
; Krempien et al., 1981
; Blasco et al., 1991
). These structures were seen only late during infection and required VV particle formation (Hiller et al., 1979
). However, the proposal by Cudmore et al. (1995)
was problematic. First, cytochalasin D prevented actin tail formation but CEV particles were still found on the cell surface (Payne & Kristensson, 1982
). Second, there are several virus mutants that are unable to produce actin tails (Table 1
) but which still form CEV and EEV and in some cases with enhanced EEV levels. Third, the drug PP1, which inhibits tyrosine phosphorylation [which is necessary for actin tail formation (Frischknecht et al., 1999a
, b
)], did not prevent CEV formation (Hollinshead et al., 2001
). Evidently, transport to the surface is not reliant on actin polymerization. Lastly, the actin tails are found on one side only of the virus particle. This is reminiscent of intracellular bacteria such as Listeria and Shigella that also induce actin polymerization, but in those cases a bacterial protein located at one end only of the bacterium directs actin polymerization to that site only (Goldberg & Theriot, 1995
; Smith et al., 1995
). Yet with VV, the A36R protein, which is required for actin tail polymerization (see below), is distributed evenly over the IEV surface (van Eijl et al., 2000
); so how is the polymerization polar?
The second model proposes that IEV move to the cell surface on microtubules, and actin tails form at the cell surface beneath CEV particles. The proposal that actin tails form only at the cell surface was based on the observation that the A36R protein was on IEV and beneath CEV on the cytosolic face of the plasma membrane, but was absent from CEV and EEV (van Eijl et al., 2000 ). This location is ideal to induce actin tail formation to drive the particle away from the cell. Subsequently, several groups utilizing GFP-labelled virions reported that IEV movement to the cell surface requires microtubules (Geada et al., 2001
; Hollinshead et al., 2001
; Rietdorf et al., 2001
; Ward & Moss, 2001
). IEV were found to move along defined pathways rather than randomly in the cytosol, their movement was inhibited reversibly by nocodazole, and they moved in a stopstart manner with an average speed of 60 µm/min characteristic of microtubular transport but 20-fold greater than VV movement on actin tails (2·8 µm/min) (Cudmore et al., 1995
).
These observations demonstrate that microtubules are used at two stages during VV egress: first, for transport of IMV from the virus factories toward the MTOC, and second, for transport of IEV from the MTOC to the cell surface (Fig. 1). Evidence that disruption of dynein-dynactin (Ploubidou et al., 2000
) inhibited IMV movement, while disruption of kinesin inhibited IEV movement (Rietdorf et al., 2001
), and the presence of different proteins on the surface of IMV and IEV is consistent with this. For IMV, the A27L protein is implicated directly or indirectly in microtubular movement, but which IEV proteins are involved? IEV proteins F12L, F13L and A36R are candidates because they are predominantly cytosolic (Fig. 3
). Of these, F13L is required for IEV formation (Blasco & Moss, 1991
) whereas F12L and A36R are not. In the absence of A36R, IEV are transported to the cell surface and CEV are visible by confocal and electron microscopy (Sanderson et al., 1998a
; Wolffe et al., 1998
; van Eijl et al., 2000
; Hollinshead et al., 2001
), although another study reported that A36R is needed for IEV movement on microtubules (Rietdorf et al., 2001
). The third protein, F12L, seems a better candidate for microtubular movement.
The F12L protein is a 6570 kDa protein that is conserved in chordopoxviruses (Zhang et al., 2000 ). Immunoelectron microscopy using an epitope-tagged F12L revealed the protein is located on the IEV surface and is absent from CEV and EEV. In this respect it resembles A36R, but one difference is the absence of F12L beneath CEV at the cell surface (van Eijl et al., 2002
). A mutant lacking the F12L protein made IEV, but IEV were not transported to the cell surface, EEV levels were reduced, the plaque size was small and the virus was highly attenuated (Zhang et al., 2000
; van Eijl et al., 2002
). Disruption of the corresponding gene in fowlpox virus also caused a small plaque phenotype and decreased EEV production (Ogawa et al., 1993
). The VV F12L deletion mutant is the only mutant reported to make IEV particles that are not transported and thus is a prime candidate for interactions with microtubules. The mode of interaction of F12L with the IEV is not understood.
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Actin tail formation |
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The A36R protein was originally described as a component of the EEV surface based upon its co-purification with EEV and its sensitivity to digestion by exogenous trypsin (Parkinson & Smith, 1994 ). However, subsequent analysis showed that the protein was absent from CEV and EEV (van Eijl et al., 2000
). The A36R protein has a type 1b membrane topology with the majority of the amino acids in the cytosol (Fig. 2
) (Röttger et al., 1999
; Grosenbach et al., 2000
; van Eijl et al., 2000
), explaining why the six potential sites for attachment of N-linked carbohydrate are unused (Parkinson & Smith, 1994
). Although the A36R protein is expressed by all orthopoxviruses examined, the protein varies in length and sequence near the C terminus (Pulford et al., 2002
) and in ectromelia virus strain MP1 the protein is significantly shorter (160 amino acids versus 220 in VV). Mutagenesis demonstrated that truncated versions of A36R can still induce actin tail formation, but phosphorylation of Tyr-112 [an amino acid conserved in all sequenced A36R proteins (Pulford et al., 2002
)] is essential and can be inhibited by PP1 (Frischknecht et al., 1999b
). After phosphorylation A36R interacts with Nck leading to recruitment of N-WASP to the site of actin assembly (Frischknecht et al., 1999b
). The recruitment of A36R to IEV requires the A33R protein, which functions as a chaperone and with which A36R forms a non-covalent complex (Wolffe et al., 2001
). Tyrosine phosphorylation of A36R is reduced in the absence of A34R or F13L and inhibited in the absence of A33R (Wolffe et al., 2001
). The A36R protein is also phosphorylated on serine and threonine residues (Wolffe et al., 2001
) and is acylated via Cys-25 (Grosenbach et al., 2000
). Deletion of A36R causes a dramatic attenuation (Parkinson & Smith, 1994
) comparable to that resulting from loss of F12L (Zhang et al., 2000
).
A direct comparison of the plaque-size phenotype of all mutants listed in Table 1 (Law et al., 2002
) highlighted the role for actin tails in cell-to-cell spread. These mutants include those lacking F12L (Zhang et al., 2000
), F13L (Blasco & Moss, 1991
; Cudmore et al., 1995
), A33R (Roper et al., 1998
), A34R (Duncan & Smith, 1992
; McIntosh & Smith, 1996
; Wolffe et al., 1997
; Sanderson et al., 1998a
), A36R (Parkinson & Smith, 1994
; Sanderson et al., 1998a
; Wolffe et al., 1998
; Frischknecht et al., 1999b
; Röttger et al., 1999
) and B5R (Engelstad & Smith, 1993
; Wolffe et al., 1993
; Mathew et al., 1998
; Sanderson et al., 1998a
; Röttger et al., 1999
). One report that a mutant lacking the SCR domains of B5R produced a normal size plaque but failed to produce actin tails (Herrera et al., 1998
) did not fit with this model, but upon re-examination this mutant was found to form a small plaque (Rodger & Smith, 2002
). The A56R protein is the only IEV/CEV/EEV protein not needed for efficient actin tail formation (Sanderson et al., 1998a
).
Two other observations are noteworthy regarding VV-induced actin tail formation. First, VV gene A42R encodes a profilin-like protein (an actin-binding protein) but this is not required for formation of actin tails or for virus maturation and egress (Blasco et al., 1991 ). Second, VV infection induces cell migration and subsequent cellular projections up to 160 µm long that often are branched and require drastic rearrangement of actin cytoskeleton of the host cell (Sanderson et al., 1998b
). Cell migration required early virus gene expression only, whereas formation of projections required both early and late virus gene expression (Sanderson et al., 1998b
).
Actin tails can continue to grow from the cell surface for considerable distances (Hiller et al., 1979 ) and facilitate virus penetration of surrounding cells. These tails can also re-enter the same cell (Hollinshead et al., 2001
). Eventually, as the tail grows longer, it may be detached from the cell still containing the CEV at its tip. Alternatively, the CEV may be released to form EEV.
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Release of EEV |
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Why does VV retain CEV? |
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Haemagglutination and haemadsorption |
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Comparisons of the HA sequences from different orthopoxviruses showed that the N-terminal Ig domain is more highly conserved than regions between the Ig domain and the transmembrane domain (Aguado et al., 1992 ; Cavallaro & Esposito, 1992
). Replacement of the Ig domain with a single-chain antibody specific for the tumour-specific antigen ErbB2 enabled the fusion protein to be incorporated into the EEV envelope and for the EEV to bind to ErbB2 (Galmiche et al., 1997
). Thus it may be possible to alter the tropism of EEV as a step towards specific anti-tumour therapy. Transcriptional and immunoblot analyses revealed that the HA is expressed from both early and late promoters but the majority of HA accumulates late (Brown et al., 1991b
).
An unusual feature of the HA is that it functions to inhibit cellcell fusion. This was demonstrated by comparison of the HA+ IHD-J and HA- IHD-W strains that are fusion (F)- or F+, respectively (Ichihashi & Dales, 1971 ). Co-infection with both viruses prevented fusion (Ichihashi & Dales, 1971
). In contrast, treatment of IHD-J-infected cells with HA-specific mAb induced fusion (Seki et al., 1990
). Analysis of 21 haemadsorption-negative mutants (Shida & Matsumoto, 1983
) showed that 19 of these failed to express cell surface HA and were F+, and five HA-positive revertants were F- (Seki et al., 1990
). The other two mutants expressed HA at the cell surface, but had single amino acid substitutions that caused either loss of haemadsorption activity but retention of fusion inhibitory activity (Glu-121 to Lys), or loss of both activities (Cys-103 to Tyr) (Seki et al., 1990
). Two other VV proteins also affect cellcell fusion: the K2L serine protease inhibitor is a fusion-inhibition protein like HA (Law & Smith, 1992
; Turner & Moyer, 1992
; Zhou et al., 1992
); and the A27L IMV surface protein promotes fusion (Rodriguez et al., 1987
).
Deletion of the A56R gene does not affect virus morphogenesis, plaque size or EEV release, but the plaques are syncytial and the deletion mutant shows attenuation if injected intracranially into mice (Flexner et al., 1987 ) but not if administered intranasally (G. L. Smith, unpublished data). The function of the HA in the virus life-cycle is not understood; in particular it is curious to have a fusion-inhibition protein on the surface of EEV.
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Incorporation of cellular proteins into EEV |
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Several host membrane proteins that are present in the TGN, early endosomes or plasma membrane fractions have been found in EEV preparations e.g. CD46, CD55, CD59, MHC class I and others (Vanderplasschen et al., 1998b ; Krauss et al., 2002
). Where investigated by electron microscopy these have also been found in IEV, CEV or EEV at low levels. Presumably these proteins are incorporated into the IEV outer membranes during wrapping. Biologically, the presence of CD55 protected EEV against destruction by homologous complement (Vanderplasschen et al., 1998b
).
Host proteins from the ER, IC and early Golgi membranes were not found in EEV preparations suggesting these membranes are not utilized for EEV formation. Similarly, these antigens were not detected in IMV preparations (Krauss et al., 2002 ). This demonstrated that if membranes of the IC are utilized to form IMV particles there must be a mechanism to exclude host antigens from these membranes during morphogenesis.
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Proteinprotein interactions |
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Mechanisms of virus spread |
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EEV interactions with antibody and complement |
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There have been conflicting reports about the neutralization of EEV by antibody. Early reports indicated that neutralization of EEV by antibody was possible if antibody was derived from convalescent serum after a live infection (reviewed in Boulter & Appleyard, 1973 ), Later reports that EEV were not neutralized (Ichihashi, 1996
; Vanderplasschen et al., 1997
) have been refuted (Galmiche et al., 1999
; Law & Smith, 2001
) although higher concentrations of serum or purified antibody are needed to achieve the same degree of neutralization as obtained with IMV. A serum against purified EEV antigens or only against the extracellular domain of B5R each inhibited EEV infectivity (Galmiche et al., 1999
). Further analysis indicated that SCR domain 1 of B5R was a target for this neutralizing antibody and that infectivity was reduced by inhibition of binding to cells and by virus aggregation (Law & Smith, 2001
).
The B5R protein remains the only EEV antigen identified as a target for neutralizing antibody, although immunization of animals with A33R protein or recombinant DNA, or passive transfer of antibody to A33R protein, also induced protection against challenge (Galmiche et al., 1999 ; Hooper et al., 2000
). This observation is relevant to the use of VV strain LC16m8 as the smallpox vaccine in Japan. LC16m8 was introduced during the latter years of the smallpox eradication campaign because of its increased safety compared to the parental Lister strain (Hashizume et al., 1985
). However, this virus does not make the B5R protein and the reduced plaque size of LC16m8 was attributable to this defect (Takahashi-Nishimaki et al., 1991
). Given that B5R is the only established target for EEV neutralizing antibody, this virus might have diminished potency as a smallpox vaccine.
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EEV binding and entry |
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The study of EEV binding and entry has lagged behind that of IMV because of the low amounts of virus, the difficulty in obtaining EEV preparations that are free from IMV contamination and the fragility of the EEV outer envelope. For IMV, the A27L (Chung et al., 1998 ), D8L (Maa et al., 1990
; Hsiao et al., 1999
), and H3L (Lin et al., 2000
) proteins have all been demonstrated to bind to cell surface glycosaminoglycans, and an IgM mAb to a cell surface antigen blocked the binding of IMV to the cell surface (Chang et al., 1995
). For EEV, no specific virus protein has been demonstrated to bind to a cell molecule, although the A34R and B5R proteins may have a role due to the increased release of EEV when these proteins are mutated (Blasco et al., 1993
; McIntosh & Smith, 1996
; Herrera et al., 1998
; Mathew et al., 1998
) and the reduced specific infectivity of A34R-deficient EEV (McIntosh & Smith, 1996
). Even the factor to which HA binds on rooster erythrocytes (the haemagglutination reaction) is unknown.
To study EEV binding and entry it is necessary to either use pure preparations of EEV or to distinguish IMV and EEV particles in mixed populations and measure each simultaneously. Although physical methods exist to separate IMV and EEV due to their different buoyant densities (Boulter & Appleyard, 1973 ), these processes result in damage to the EEV outer envelope so that an increased proportion of infectivity is neutralized by IMV-specific mAb (Ichihashi, 1996
; Vanderplasschen & Smith, 1997
). Such damaged virions might bind to cells via either EEV or IMV antigens. To overcome these difficulties, EEV binding was studied using fresh EEV preparations and confocal microscopy. The IMV and EEV particles were distinguished using mAbs specific for the IMV or EEV surface (Vanderplasschen & Smith, 1997
, 1999
). Using this methodology it was shown that: (i) IMV and EEV bind to different cell types with differing relative efficiency; (ii) treatment of cells with Pronase, trypsin or neuraminidase affected IMV and EEV binding differently; (iii) a mAb that blocked the binding of IMV to the cell surface (Chang et al., 1995
) did not affect EEV binding; and (iv) IMV and EEV bound to distinct sites on the cell surface (Vanderplasschen & Smith, 1997
). Evidently, IMV and EEV bind to different receptors.
The mechanism of EEV entry is not understood. A fundamental issue is the number of lipid bilayers that must be shed from the virion to enable the core to access the cytosol. If IMV has a single membrane then EEV has two, and if IMV has two or more membranes EEV has three or more. A single fusion event cannot enable the EEV core to enter the cytosol and the mechanism of EEV entry must result in loss of one more membrane than IMV. Early studies on VV entry (using IMV) reported that entry was via pinocytosis (Dales, 1965 ). Other workers reported fusion at the plasma membrane (Armstrong et al., 1973
; Chang & Metz, 1976
; Janeczko et al., 1987
) and showed electron micrographs of the IMV surface membrane in continuity with the plasma membrane (Armstrong et al., 1973
; Chang & Metz, 1976
). In addition to thin section electron microscopy Chang & Metz (1976)
detected virus antigen on the cell surface after virions had penetrated the cells, consistent with cell surface fusion.
Several methods to study EEV entry have been used including electron (Krijnse Locker et al., 2000 ) and confocal microscopy (Vanderplasschen et al., 1998a
; Krijnse Locker et al., 2000
) to follow the appearance of cores within the cytosol, loss of radioactively labelled virions from the cell surface (Payne & Norrby, 1978
) and a lipid mixing assay based upon dilution of a fluorescent probe (Doms et al., 1990
). While these studies all conclude that EEV enters more rapidly than IMV, despite having to shed an additional lipid membrane, there is discrepancy about whether the penetration is affected by pH, where fusion takes place and whether drugs that affect actin influence entry. When comparing the data from these different studies the method used to obtain the EEV preparation should be considered. EEV purified by centrifugation and with an additional labelling procedure may have an increased proportion of virions with damaged outer envelopes and thus might bind to cells via either IMV or EEV proteins. This type of preparation should be avoided. Using fresh EEV from the supernatant of infected cells and an IMV mAb to neutralize IMV, Ichihashi (1996)
proposed a model for EEV entry that required a low-pH step. In this model EEV are taken up by pinocytosis into intracellular vesicles that become acidified. At reduced pH the outer EEV membrane is disrupted and the IMV particle released into the vesicle fuses with the vesicle membrane releasing the core into the cytosol. In support of this model, Vanderplasschen et al. (1998a
) noted that drugs that raise the intracellular pH reduced the uptake of EEV but not IMV, and a low-pH shock caused rupture of the EEV outer membrane so that virion infectivity was neutralized by an anti-IMV mAb. On the other hand Doms et al. (1990)
reported that the rate of fusion of IMV and EEV was not affected by pH. Another study reported that IMV, but not EEV, induce signalling and the formation of actin-containing cell surface protrusions (Krijnse Locker et al., 2000
). Another model for IMV entry proposed that IMV enter cells without a membrane fusion event: IMV were suggested to unfold' outside the cell and cores were then somehow able to pass across the plasma membrane (Krijnse Locker et al., 2000
; Griffiths et al., 2001
; Sodeik & Krijnse Locker, 2002
). This proposal is inconsistent with the images of IMV membrane in continuity with the plasma membrane (Armstrong et al., 1973
; Chang & Metz, 1976
). Additional studies are needed to determine the exact mechanisms of VV entry.
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Summary |
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Acknowledgments |
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References |
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