Centro de Investigación en Sanidad AnimalINIA, Valdeolmos, E-28130 Madrid, Spain1
Author for correspondence: Rafael Blasco. Fax +34 91 620 22 47. e-mail blasco{at}inia.es
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Abstract |
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Introduction |
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The best studied member of the poxvirus family, vaccinia virus (VV), belongs to the genus Orthopoxvirus. The classification of SPV in a separate genus, based originally on immunological studies, was later reinforced by DNA hybridization, restriction mapping and DNA sequencing (Massung & Moyer, 1991a , b
). The limited sequence information available for SPV, including sequences close to the genome ends as well as internal regions (Feller et al., 1991
; Massung et al., 1993
; Schnitzlein & Tripathy, 1991
) has revealed enough similarity to other poxviruses to allow alignment of gene maps, despite extensive sequence divergence.
The major protein component of the envelope of extracellular vaccinia virions (EEV), P37, is encoded by the F13L open reading frame (ORF) (Hirt et al., 1986 ). It is a 372 amino acid polypeptide expressed at late times during infection and is targeted to Golgi-derived membranes where it is incorporated into the virion by a wrapping process (Hiller & Weber, 1985
; Schmelz et al., 1994
). Thus P37 is present in the enveloped forms of the virus, but is absent from intracellular mature virus (IMV) (Payne, 1978
). P37 is a peripheral membrane protein, and lines the inner surface of the EEV envelope (Roos et al., 1996
; Schmutz et al., 1995
). Palmitylation of the protein, a process that is facilitated by other viral proteins, is required for its membrane-association, proper intracellular targeting and correct functioning of the protein (Borrego et al., 1999
; Grosenbach & Hruby, 1998
; Grosenbach et al., 1997
; Schmutz et al., 1995
).
Deletion of the P37 gene results in a block of the virus envelopment process, and abolishes extracellular virus formation and cell-to-cell virus transmission (Blasco & Moss, 1991 , 1992
). In addition, other virus-induced phenomena that require virus envelopment, such as engrossed actin tails and cellcell fusion at acid pH, are also blocked (Blasco & Doms, 1993
; Blasco & Moss, 1992
; Cudmore et al., 1995
; Sanderson et al., 1998
).
Considering that P37 is the major protein in the EEV envelope, its topology relative to the EEV envelope and the phenotype of P37 knock-out mutants, it is likely that P37 acts by mediating the interaction between the surface of intracellular virions and wrapping membranes. Recent reports suggest that in addition to this structural role, P37 may have enzymatic activities related to lipid metabolism. Indeed, the P37 sequence contains motifs distinctive of the phospholipase D superfamily (Koonin, 1996 ; Ponting & Kerr, 1996
), and shows phospholipase activities which may be important for function (Baek et al., 1997
; Sung et al., 1997
).
Because of the importance of P37 in VV envelope morphogenesis and dissemination, we mapped and sequenced the SPV homologous gene, and present here a characterization of the corresponding protein.
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Methods |
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Cloning and sequencing.
SPV DNA fragment HindIII B (Massung & Moyer, 1991a ) was excised from an agarose gel and digested with EcoRI. A 5·5 kb HindIIIEcoRI fragment derived from the left-hand end of the HindIII B fragment was cloned into plasmid pUC19 generating plasmid pSPV-HE. A 1·9 kb NcoIBamHI fragment derived from pSPV-HE was cloned in plasmid pUCPTS-3 (Barcena & Blasco, 1998
) to generate pSPV-NB. Both pSPV-HE and pSPV-NB were used as templates for sequencing the P42 gene using the dideoxy chain termination method. Both DNA strands were sequenced.
Sequence analysis.
Pairwise alignment of amino acid sequences was performed using the Align program (FASTA program package, version 2.0) (Pearson, 1990 ). Pairwise percentage identities (or similarities) were calculated by dividing the number of identical residues (and conservative changes) by the total number of positions in the aligned sequences, including gaps. Multiple alignment of amino acid sequences was generated using the program Clustal W version 1.7 (Thompson et al., 1994
). Hydrophobicity analysis of the amino acid sequences was based on the algorithm of Kyte & Doolittle (1982)
.
Plasmid construction.
Plasmid pRB24 was constructed to insert foreign genes in place of F13L, under the control of the natural F13L promoter. The plasmid contains F13L recombination flanks, XhoI and BglII restriction sites immediately downstream of the P37 promoter, and a -glucuronidase (
-gus) cassette to provide visual identification of recombinant viruses.
The process of plasmid construction is outlined in Fig. 3(A) (see Results). First, the
-gus gene was amplified by PCR from plasmid pBI101 (Clontech) using primers 5' CAGGTCAGAATTCTATGTTACGTCC 3' (EcoRI site underlined) and 5' GGAGAGTTGCTAGCTCATTGTTTGCC 3' (NheI site underlined), and inserted into the EcoRINheI sites of plasmid pRB21 (Blasco & Moss, 1995
), to construct pRB21-
gus. Then, a SphIXhoI DNA fragment of pRB21-
gus, containing the P37 gene, was removed and replaced by a PCR product obtained using pRB21 as template and oligonucleotides 5 GGGGCATGCGATAAAGTTTCGAAACAGCAAAA 3' (SphI site underlined) and 5' CGATGCCTCGAGATCTATTTAGTTACATAAAAAC 3' (XhoI and BglII sites underlined) as primers, generating plasmid pRB24. Subsequently, the genes corresponding to SPV protein P42 or VV protein P37 were inserted into pRB24, generating plasmids pRB24-P42 and pRB24-P37 respectively. The gene encoding P42 protein was amplified by PCR from plasmid pSPV-HE using primers 5 AAATAAAGGATCCGTATGTGGTGG 3' (BamHI site underlined) and 5' ACGTCCTGGATCCAAATATATTTTC 3' (BamHI site underlined). The PCR product containing the P42 gene was partially digested with BamHI, ligated to pRB24 plasmid digested with BglII, and checked by DNA sequencing. Finally, a BamHI fragment from pSG-P37 plasmid (B. Borrego and others, unpublished results), containing the P37 gene, was isolated and ligated to BglII-digested pRB24 to generate pRB24-P37.
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Antisera.
A rat monoclonal antibody, 15B6 (Schmelz et al., 1994 ), was used to detect the P37 protein. Antiserum against an 18-mer synthetic peptide corresponding to the C terminus of P42 (NH2-CDIFERDWKNNSNTPITN-COOH) was obtained. The peptide, conjugated with diphtheria toxoid protein, was injected subcutaneously into rabbits in three doses. The first dose contained 500 µg of protein in complete Freunds adjuvant and the following two contained 500 µg of protein in incomplete Freunds adjuvant.
Immunofluorescence.
PK-15 cells grown on coverslips were infected with SPV or VV (strain WR). After 23 h (for SPV) or 7 h (for VV) at 37 °C, cells were washed twice with PBS at room temperature, fixed by addition of ice-cold 4% paraformaldehyde, and incubated for 12 min at room temperature. All subsequent incubations were carried out at room temperature. After washing once with PBS, cells were permeabilized by incubation with 0·1% Triton X-100 in PBS. After washing with PBS, cells were incubated for 5 min with PBS containing 0·1 M glycine, and then with primary antibodies diluted in PBS20% foetal calf serum for 30 min. Anti-peptide antiserum was diluted 1:75, and anti-P37 hybridoma supernatant was diluted 1:50. After washing for 5 min in PBS, the cells were incubated for 30 min with secondary antibodies: rabbit anti-mouse IgG or swine anti-rabbit IgG conjugated with TRITC rhodamine (Dako) diluted 1:200 in PBS20% foetal calf serum. After a final wash, coverslips were mounted using FluorSave mounting medium (Calbiochem).
Western blotting.
Proteins were electrophoresed in 12% SDSpolyacrylamide gels and transferred to nitrocellulose membranes by electroblotting. After transfer, the membranes were incubated overnight at 4 °C in blocking buffer (PBS containing 0·1% Tween 20 and 5% non-fat dry milk). The membranes were then incubated with monoclonal antibody anti-P37 (diluted 1:2000) or antisera anti-P42 (diluted 1:100) in PBS0·1% Tween 20 containing 1% BSA for 1 h at 37 °C. After four 10 min washes with PBS0·1% Tween 20, the membranes were incubated for 1 h at 37 °C with rat or rabbit anti-IgG antibody (diluted 1:2000) conjugated with horseradish peroxidase in PBS0·1% Tween 20 containing 1% BSA. After four 10 min washes with PBS0·1% Tween-20, bound antibodies were detected using the enhanced chemiluminescence (ECL) kit from Amersham.
Subcellular fractionation of infected cells.
CV-1 cell monolayers were infected with recombinant VV at an m.o.i. of 10 p.f.u. per cell. At 12 h post-infection (p.i.) cells were scraped into the medium, collected by low-speed centrifugation, washed once with PBS and resuspended in 20 mM HEPES pH 7·6, 5 mM KCl, 1 mM MgCl2, 150 mM NaCl. Subcellular fractionation was then performed as described by Grosenbach et al. (1997) . Briefly, cells were lysed by Dounce homogenization to yield a total cell extract (TCE). The TCE was subjected to centrifugation at 700 g for 10 min at 4 °C to produce a nuclear pellet fraction (NP). The supernatant was centrifuged at 15000 g for 30 min at 4 °C to obtain the virus-containing fraction (P15). Finally, the supernatant of this centrifugation was further separated into soluble (S100) and insoluble (P100) protein fractions by centrifugation at 100000 g for 1 h at 4 °C. Portions of the resulting fractions were analysed by Western blot. Subcellular fractionation of SPV-infected cells was carried out as described above using PK-15 cell monolayers infected at an m.o.i. of 10 for 48 h.
Preparation and analysis of [3H]palmitylated VV and SPV proteins.
Infected cells were radiolabelled with [3H]palmitic acid essentially as described by Child & Hruby (1992) . Briefly, confluent monolayers of CV-1 cells were infected with the different recombinant VVs at an m.o.i. of 10. At 16 h p.i., cell cultures were labelled for 4 h with 200 µCi/ml [9,10-3H]-palmitic acid (Amersham, 51 Ci/mmol). Radiolabelled SPV-infected cells were prepared as described above using PK-15 cells infected for 48 h. For SDSPAGE analysis, infected-cell extracts were prepared by washing the cell monolayers with PBS followed by solubilization in 2% SDS. After electrophoresis in 11% SDSpolyacrylamide gels, the gels were fixed, fluorographed, dried, and exposed to an X-ray film at -70 °C.
Virion purification and CsCl gradient analysis.
For equilibrium centrifugation, confluent monolayers of RK-13 cells in six-well plates were infected with virus at an m.o.i. of 10. At 7 h p.i., medium was replaced with 1 ml of solution comprising two-thirds of methionine-free MEM and one-third of complete MEM supplemented with 25 µCi [35S]methionine. At 24 h p.i., 1 ml of complete MEM with 5% foetal calf serum was added, and the incubation continued for another 24 h. Extracellular virus was then recovered from the medium after clarification by low-speed centrifugation and was pelleted through a cushion of 4 ml 36% sucrose. Finally, the virus was loaded on a gradient made by overlaying 2, 3 and 4 ml of CsCl solutions with densities of 1·30, 1·25 and 1·20 g/ml, respectively, and centrifuged in an SW41 rotor at 32000 r.p.m. for 60 min at 15 °C. Fractions (six drops) were collected from the bottom of the tube, and aliquots of each fractions were assayed for radioactivity.
Similarly, EEV and IMV forms of SPV were purified by banding in CsCl density gradients. Confluent monolayers of PK-15 cells in six-well plates were infected with SPV virus at an m.o.i. of 10. At 48 h p.i., culture medium was replaced with medium containing 25 µCi [35S]methionine as described above, and the incubation was continued for another 24 h. In parallel, PK-15 cells in a 175 cm2 flask were infected with SPV at a multiplicity of 10 and incubated for 72 h. From both cultures, extracellular virus was recovered from the medium after clarification by low-speed centrifugation, and cell-associated virus was liberated by rupturing the cells in 10 mM TrisHCl (pH 9) by Dounce homogenization. At this step, both the radiolabelled and non-radiolabelled preparations of the extracellular and cell-associated virus were mixed, pelleted through a sucrose cushion and subjected to buoyant density centrifugation as described above. Fractions were assayed for radioactivity and density. Finally, radioactive peaks corresponding to fractions with densities of 1·23 g/ml and 1·27 g/ml were pooled and used as purified EEV and IMV, respectively.
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Results |
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Our sequencing revealed the typical poxvirus late promoter motif TAAAT, and the similar TAAAAT motif, upstream of the putative ATG initiation codon (see Fig. 1). The early transcription termination sequence TTTTTNT occurs in three different places within the ORF, suggesting an exclusively late expression pattern.
SPV P42 amino acid sequence
The calculated molecular mass of the product of the SPV ORF is 41779 Da, which is similar to the calculated size of VV P37, and somewhat larger than the size of 37 kDa estimated previously by its mobility in PAGE. On the basis of its predicted size, the putative SPV polypeptide was termed P42.
A comparison of the amino acid sequences of P37 homologues from different poxviruses is shown in Fig. 2. SPV P42 was most similar to the MV protein (73·1% identity) and VV P37 (54·3% identity), suggesting that SPV is more closely related to leporipoxviruses and orthopoxviruses than to the other poxvirus genera included in this study. Indeed, this result is in agreement with poxvirus phylogenies based on thymidine kinase sequences (Blasco, 1995
).
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Construction of recombinant viruses
In order to study SPV P42, we constructed several recombinant VVs in which either VV P37 or SPV P42 genes were placed downstream of the natural VV P37 promoter. To facilitate the isolation of recombinant viruses, plasmid pRB24 was constructed, as outlined in Fig. 3(A). This plasmid contains flanks for insertion into the P37 locus and a
-gus cassette to allow visual identification of recombinant virus plaques. The coding sequences of VV P37 or SPV P42 ORFs were inserted downstream of the P37 promoter, to generate recombinant viruses expressing either protein. As shown in Fig. 3(B)
, the DNA sequence between the promoter and the initiation ATG was slightly changed from wild-type virus due to the cloning. Recombinant viruses I-
, I-
P37 and I-
P42 were derived from VV vRB10, a VV (IHD-J strain) P37 deletion mutant, by recombination with plasmids pRB24, pRB24-P37 or pRB24-P42, respectively. Recombinant viruses were isolated by several consecutive rounds of plaque isolation in the presence of X-Gluc.
Expression of SPV P42
An antiserum was raised against a synthetic peptide corresponding to the C terminus of the protein. The antiserum specifically recognized SPV P42, when this was expressed from VV recombinant I-P42 (Fig. 4A
). In SPV-infected PK-15 cells, the antiserum recognized a polypeptide with an apparent molecular mass of 40 kDa, in good agreement with the predicted molecular mass for P42 (Fig. 4A
).
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Palmitylation of SPV P42
Because of the overall sequence conservation between VV P37 and SPV P42, and the conservation of the two cysteine residues that are modified by palmitylation in VV P37, we wished to determine whether SPV P42 was modified by palmitylation. Labelling with [3H]palmitic acid during infection by SPV produced two major labelled proteins, one of which had an apparent molecular mass of 42 kDa (Fig. 5, lane SPV). This protein comigrated with immunoprecipitated, or palmitate-labelled, VV P37.
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Localization of SPV P42
In order to study the subcellular distribution of SPV P42, and compare it to that of VV P37, cells infected with I-P37, I-
P42 or SPV were subjected to subcellular fractionation as detailed in Methods. After fractionation, samples were analysed by Western blot (Fig. 6A
) with anti-P37 or anti-P42 antibody. P37 was detected in significant amounts in the nuclear fraction (NP) and the virus-containing fraction (P15), which potentially contains some of the cytoplasmic membrane-bound organelles. No P37 was detected in the particulate cytoplasmic fraction (P100) or in the soluble cytoplasmic fraction (S100). These results are consistent with those reported previously (Grosenbach & Hruby, 1998
; Grosenbach et al., 1997
). After fractionation of SPV-infected cells, P42 showed roughly the same pattern, suggesting that P37 and P42 share a similar intracellular distribution. Fractionation of I-
P42-infected cells also produced a similar pattern (Fig. 6A
), indicating that the SPV P42 protein expressed in the context of a VV infection attains its correct subcellular distribution.
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The intracellular localization of P42 was also explored by immunofluorescence staining of infected cells. Staining with anti-P42 antibody in SPV-infected cells revealed a strong juxtanuclear signal, and some dispersed punctate staining (Fig. 7B), similar to that of P37 in VV-infected cells (Fig. 7D
). This localization pattern is characteristic of VV EEV envelope proteins, which at late times are enriched in the trans-Golgi network (juxtanuclear staining), and are also present in intracellular or cell-associated enveloped virus particles (punctated staining). The strong juxtanuclear labelling suggests efficient trans-Golgi network targeting of P42 in the context of SPV infection. Also, the punctate pattern obtained (which showed fewer and in general bigger structures than in the case of VV) is consistent with the incorporation of P42 in the enveloped forms of the virus, by a mechanism similar to that described for VV.
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VV recombinants expressing VV P37 (I-P37) or SPV P42 (I-
P42) as well as P37 deleted virus (I-
) were assayed for plaque formation, EEV release and EEV infectivity. Monolayers of BSC-1 cells were infected with wild-type VV (IHD-J) or the recombinant viruses and incubated under an agarose overlay. At 48 h p.i. the cell monolayers were stained with crystal violet to visualize virus plaques (Fig. 8A
). I-
P37 virus formed clearly visible plaques, which were slightly smaller than normal IHD-J virus plaques, probably due to differences in the sequence around the P37 promoter (shown in Fig. 3B
). As expected for a P37 deletion mutant (Blasco & Moss, 1991
), virus I-
produced no visible plaques in 48 h and formed small plaques after 67 days (not shown). I-
P42 plaques were indistinguishable from I-
plaques, indicating that P42 expression did not rescue the defect of the VV P37 deletion mutant. When the monolayers were maintained under a liquid medium overlay, the characteristic comet-like plaques typical of IHD-J virus formed after 2 days for IHD-J and I-
P37, and after 7 days for I-
and I-
P42 (data not shown), indicating that EEV release was not enhanced by expression of P42. These observations were not dependent on the cell type, similar results being obtained with swine PK-15 cells (data not shown).
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Both the development of virus plaques (Fig. 8A) or the release of high virus titres into the culture medium (Fig. 8B
) are dependent on the production and release of infectious virus. The phenotype of I-
P42 virus could reflect either a block in EEV formation or a defect in EEV infectivity. To distinguish between these possibilities, infected RK13 cells were labelled with [35S]methionine and the virus in the medium was analysed by CsCl gradient centrifugation (Fig. 8C
). I-
P37 produced a high peak of radioactivity in fractions corresponding to the density of the EEV band (1·23 g/ml). I-
and I-
P42 had drastically reduced levels of EEV particles, although detection of enveloped virions above background was possible. Thus, expression of P42 was not able to rescue EEV formation in a VV P37 deficient background.
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Discussion |
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In addition to sequence homology, our characterization of SPV P42 revealed significant resemblance to its VV counterpart. Similar to VV P37, the SPV protein showed an exclusively late expression pattern, had a similar intracellular distribution, was incorporated into EEV and was absent from IMV. These data led us to propose that both proteins share the same function in the morphogenesis of their respective enveloped virions. That SPV P42 is functional is supported by the observation that despite its slower replication kinetics with respect to VV (Massung et al., 1991b ), SPV forms larger plaques, and releases more EEV than VV P37 deletion mutants (J. Bárcena, J. M. Sánchez-Puig & R. Blasco, unpublished). Deletion of the SPV gene, in a similar way to the deletion of its VV counterpart (Blasco & Moss, 1991
) would be of interest to unequivocally determine the function of P42. However, SPV P42 deletion will presumably require the isolation of poorly plaquing or non-plaquing viruses, which is hampered by the low effectiveness of selection in SPV when using marker genes commonly used for VV selection (J. Bárcena & R. Blasco, unpublished).
Several studies point to a close connection between P37 function, palmitylation and intracellular targeting. By mutation of the palmityl acceptor cysteine residues, it has been shown that proper subcellular localization of P37, as well as functionality, are dependent on palmitylation of the protein (Grosenbach & Hruby, 1998 ; Grosenbach et al., 1997
). Also, the infection context, and probably the interaction with other envelope proteins, is important for efficient palmitylation and targeting of P37 (Borrego et al., 1999
). Since SPV P42 appears to have the same localization as VV P37, and the palmitylation site is conserved, we considered it likely that P42 was also modified by palmitylation. Indeed, our data suggest that P42 is the major palmitylated protein in a normal SPV infection (Fig. 5
).
Broad similarities between SPV P42 and VV P37 led us to attempt to rescue the VV P37 deficiency by expressing SPV P42 in the VV genetic background. Notably, when expressed in VV, SPV P42 seemed to be normally palmitylated and targeted. However, no complementation of the P37 deficiency was apparent, either for virus transmission or EEV formation. There are several possible explanations for this lack of cross-species functionality. Despite all of the above observations, we cannot formally rule out the possibility that the two proteins have different functions. Also, it is possible that each protein is adapted to function exclusively in cells of the host of their respective viruses. However, we favour an explanation based on the likely requirements for proteinprotein interactions between P37 (or P42) and other viral proteins involved in EEV formation. If those interactions are required for function, it is not surprising that a structural protein like SPV P42 may not function properly in the context of a different virus, like VV, provided that its protein partners have also undergone enough divergent evolution. From that point of view, the likely scenario is that concerted evolution of several virus proteins would result, eventually, in a lack of function of one single protein when isolated from its genetic context.
The absence of cross-species complementation is in sharp contrast with observations obtained with thymidine kinase genes, which are functional after being transferred between different poxviruses (Boyle & Coupar, 1986 ; Gruidl et al., 1992
; Scheiflinger et al., 1997
) or even between herpesviruses and poxviruses (Mackett et al., 1982
; Panicali & Paoletti, 1982
). Probably, complementation is easily achieved in the case of enzymes, where providing the enzymatic activity could be sufficient to provide successful complementation. Conversely, when protein interactions are required for function, the divergence between different viruses is expected to result in lack of complementation. This system, or similar situations when a particular function is affected by inter-genus divergence, has potential practical applications. For instance, one could study the protein(s) that interact with SPV P42 by incorporating additional SPV genes in the VV background. Also, recombinant viruses containing SPV P42VV P37 protein chimaeras could allow study of the protein regions involved in the proteinprotein interactions required for function.
SPV is a potential vector for the construction of recombinant vaccines for pigs. The possible widespread use of such a vector raises several concerns related to its safety. First, the possibility of horizontal transmission to an unintended host, and in particular to humans, should be addressed. In this respect, SPV shows an extremely narrow host range in vivo, and lack of transmission of SPV to humans and other species has been reported (Schwarte & Biester, 1941 ). A second concern regarding the use of SPV, or other poxviruses, is that the virus vector may change its biological properties, alter its host range or pathogenicity, through mutation or recombination with a naturally occurring poxvirus. Our results highlight the divergence of SPV with respect to VV, and reinforce the notion that suipoxviruses and orthopoxviruses are evolutionarily distant, and therefore unlikely to undergo genetic rearrangements.
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Acknowledgments |
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Footnotes |
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References |
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Barcena, J. & Blasco, R. (1998). Recombinant swinepox virus expressing beta-galactosidase: investigation of viral host range and gene expression levels in cell culture. Virology 243, 396-405.[Medline]
Blake, N. W., Porter, C. D. & Archard, L. C. (1991). Characterization of a molluscum contagiosum virus homolog of the vaccinia virus p37K major envelope antigen. Journal of Virology 65, 3583-3589.[Medline]
Blasco, R. (1995). Evolution of poxviruses and African swine fever virus. In Molecular Basis of Virus Evolution, pp. 255-269. Edited by A. J. Gibbs, C. H. Calisher & F. García-Arenal. Cambridge: Cambridge University Press.
Blasco, R. & Doms, R. W. (1993). Membrane fusion activity of vaccinia virus. In Viral Fusion Mechanisms, pp. 413-424. Edited by J. Bentz. Boca Raton, FL: CRC Press.
Blasco, R. & Moss, B. (1991). Extracellular vaccinia virus formation and cell-to-cell virus transmission are prevented by deletion of the gene encoding the 37,000-dalton outer envelope protein. Journal of Virology 65, 5910-5920.[Medline]
Blasco, R. & Moss, B. (1992). Role of cell-associated enveloped vaccinia virus in cell-to-cell virus spread. Journal of Virology 66, 4170-4179.[Abstract]
Blasco, R. & Moss, B. (1995). Selection of recombinant vaccinia viruses on the basis of plaque formation. Gene 158, 157-162.[Medline]
Borrego, B., Lorenzo, M. M. & Blasco, R. (1999). Complementation of P37 (F13L gene) knock-out in vaccinia virus by a cell line expressing the gene constitutively. Journal of General Virology 80, 425-432.[Abstract]
Boyle, D. B. & Coupar, B. E. (1986). Identification and cloning of the fowlpox virus thymidine kinase gene using vaccinia virus. Journal of General Virology 67, 1591-1600.[Abstract]
Calvert, J. G., Ogawa, R., Yanagida, N. & Nazerian, K. (1992). Identification and functional analysis of the fowlpox virus homolog of the vaccinia virus p37K major envelope antigen gene. Virology 191, 783-792.[Medline]
Cao, J. X., Koop, B. F. & Upton, C. (1997). A human homolog of the vaccinia virus HindIII K4L gene is a member of the phospholipase D superfamily. Virus Research 48, 11-18.[Medline]
Carroll, M. W. & Moss, B. (1995). E. coli beta-glucuronidase (GUS) as a marker for recombinant vaccinia viruses.Biotechniques 19, 352-354.[Medline]
Child, S. J. & Hruby, D. E. (1992). Evidence for multiple species of vaccinia virus-encoded palmitylated proteins. Virology 191, 262-271.[Medline]
Cudmore, S., Cossart, P., Griffiths, G. & Way, M. (1995). Actin-based motility of vaccinia virus. Nature 378, 636-638.[Medline]
Feller, J. A., Massung, R. F., Turner, P. C., Gibbs, E. P., Bockamp, E. O., Beloso, A., Talavera, A., Vinuela, E. & Moyer, R. W. (1991). Isolation and molecular characterization of the swinepox virus thymidine kinase gene. Virology 183, 578-585.[Medline]
Foley, P. L., Paul, P. S., Levings, R. L., Hanson, S. K. & Middle, L. A. (1991). Swinepox virus as a vector for the delivery of immunogens. Annals of the New York Academy of Sciences 646, 220-222.[Medline]
Grosenbach, D. W. & Hruby, D. E. (1998). Analysis of a vaccinia virus mutant expressing a nonpalmitylated form of p37, a mediator of virion envelopment. Journal of Virology 72, 5108-5120.
Grosenbach, D. W., Ulaeto, D. O. & Hruby, D. E. (1997). Palmitylation of the vaccinia virus 37-kDa major envelope antigen. Identification of a conserved acceptor motif and biological relevance. Journal of Biological Chemistry 272, 1956-1964.
Gruidl, M. E., Hall, R. L. & Moyer, R. W. (1992). Mapping and molecular characterization of a functional thymidine kinase from Amsacta moorei entomopoxvirus. Virology 186, 507-516.[Medline]
Hiller, G. & Weber, K. (1985). Golgi-derived membranes that contain an acylated viral polypeptide are used for vaccinia virus envelopment. Journal of Virology 55, 651-659.[Medline]
Hiller, G., Eibl, H. & Weber, K. (1981). Characterization of intracellular and extracellular vaccinia virus variants: N1-isonicotinoyl-N2-3-methyl-4-chlorobenzoylhydrazine interferes with cytoplasmic virus dissemination and release. Journal of Virology 39, 903-913.[Medline]
Hirt, P., Hiller, G. & Wittek, R. (1986). Localization and fine structure of a vaccinia virus gene encoding an envelope antigen. Journal of Virology 58, 757-764.[Medline]
Jackson, R. J. & Hall, D. F. (1998). The myxoma virus EcoRI-O fragment encodes the DNA binding core protein and the major envelope protein of extracellular poxvirus. Virus Genes 17, 55-62.[Medline]
Koonin, E. V. (1996). A duplicated catalytic motif in a new superfamily of phosphohydrolases and phospholipid synthases that includes poxvirus envelope proteins.Trends in Biochemical Sciences 21, 242-243.[Medline]
Kyte, J. & Doolittle, R. F. (1982). A simple method for displaying the hydropathic character of a protein. Journal of Molecular Biology 157, 105-132.[Medline]
Mackett, M., Smith, G. L. & Moss, B. (1982). Vaccinia virus: a selectable eukaryotic cloning and expression vector. Proceedings of the National Academy of Sciences, USA 79, 7415-7419.[Abstract]
Massung, R. F. & Moyer, R. W. (1991a). The molecular biology of swinepox virus. I. A characterization of the viral DNA. Virology 180, 347-354.[Medline]
Massung, R. F. & Moyer, R. W. (1991b). The molecular biology of swinepox virus. II. The infectious cycle. Virology 180, 355-364.[Medline]
Massung, R. F., Jayarama, V. & Moyer, R. W. (1993). DNA sequence analysis of conserved and unique regions of swinepox virus: identification of genetic elements supporting phenotypic observations including a novel G protein-coupled receptor homologue. Virology 197, 511-528.[Medline]
Panicali, D. & Paoletti, E. (1982). Construction of poxviruses as cloning vectors: insertion of the thymidine kinase gene from herpes simplex virus into the DNA of infectious vaccinia virus. Proceedings of the National Academy of Sciences, USA 79, 4927-4931.[Abstract]
Payne, L. (1978). Polypeptide composition of extracellular enveloped vaccinia virus. Journal of Virology 27, 28-37.[Medline]
Pearson, W. R. (1990). Rapid and sensitive sequence comparison with FASTP and FASTA. Methods in Enzymology 183, 63-98.[Medline]
Ponting, C. P. & Kerr, I. D. (1996). A novel family of phospholipase D homologues that includes phospholipid synthases and putative endonucleases: identification of duplicated repeats and potential active site residues. Protein Science 5, 914-922.
Roos, N., Cyrklaff, M., Cudmore, S., Blasco, R., Krijnse-Locker, J. & Griffiths, G. (1996). A novel immunogold cryoelectron microscopic approach to investigate the structure of the intracellular and extracellular forms of vaccinia virus. EMBO Journal 15, 2343-2355.[Abstract]
Sanderson, C. M., Frischknecht, F., Way, M., Hollinshead, M. & Smith, G. L. (1998). Roles of vaccinia virus EEV-specific proteins in intracellular actin tail formation and low pH-induced cell-cell fusion. Journal of General Virology 79, 1415-1425.[Abstract]
Scheiflinger, F., Falkner, F. G. & Dorner, F. (1997). Role of the fowlpox virus thymidine kinase gene for the growth of FPV recombinants in cell culture.Archives of Virology 142, 2421-2431.[Medline]
Schmelz, M., Sodeik, B., Ericsson, M., Wolffe, E. J., Shida, H., Hiller, G. & Griffiths, G. (1994). Assembly of vaccinia virus: the second wrapping cisterna is derived from the trans Golgi network.Journal of Virology 68, 130-147.[Abstract]
Schmutz, C., Rindisbacher, L., Galmiche, M. C. & Wittek, R. (1995). Biochemical analysis of the major vaccinia virus envelope antigen. Virology 213, 19-27.[Medline]
Schnitzlein, W. M. & Tripathy, D. N. (1991). Identification and nucleotide sequence of the thymidine kinase gene of swinepox virus.Virology 181, 727-732.[Medline]
Schwarte, L. H. & Biester, H. E. (1941). Pox in swine.American Journal of Veterinary Research 2, 136-140.
Sullivan, J. T., Mercer, A. A., Fleming, S. B. & Robinson, A. J. (1994). Identification and characterization of an orf virus homologue of the vaccinia virus gene encoding the major envelope antigen p37K. Virology 202, 968-973.[Medline]
Sung, T. C., Roper, R. L., Zhang, Y., Rudge, S. A., Temel, R., Hammond, S. M., Morris, A. J., Moss, B., Engebrecht, J. & Frohman, M. A. (1997). Mutagenesis of phospholipase D defines a superfamily including a trans-Golgi viral protein required for poxvirus pathogenicity. EMBO Journal 16, 4519-4530.
Thompson, J. D., Higgins, D. G. & Gibson, T. J. (1994). CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research 22, 4673-4680.[Abstract]
Tuboly, T., Nagy, E. & Derbyshire, J. B. (1993). Potential viral vectors for the stimulation of mucosal antibody responses against enteric viral antigens in pigs. Research in Veterinary Science 54, 345-350.[Medline]
van der Leek, M. L., Feller, J. A., Sorensen, G., Isaacson, W., Adams, C. L., Borde, D. J., Pfeiffer, N., Tran, T., Moyer, R. W. & Gibbs, E. P. (1994). Evaluation of swinepox virus as a vaccine vector in pigs using an Aujeszkys disease (pseudorabies) virus gene insert coding for glycoproteins gp50 and gp63. Veterinary Record 134, 13-18.[Medline]
Received 9 October 1999;
accepted 14 December 1999.