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The previously unstudied vaccinia virus gene I2L is conserved in all orthopoxviruses. We show here that the 8-kDa I2 protein is expressed at late times of infection, is tightly associated with membranes, and is encapsidated in mature virions. We have generated a recombinant virus in which I2 expression is dependent upon the inclusion of tetracycline in the culture medium. In the absence of I2, the biochemical events of the viral life cycle progress normally, and virion morphogenesis culminates in the production of mature virions. However, these virions show an ~400-fold reduction in specific infectivity due to an inability to enter target cells. Several proteins that have been previously identified as components of an essential entry/fusion complex are present at reduced levels in I2-deficient virions, although other membrane proteins, core proteins, and DNA are encapsidated at normal levels. A preliminary structure/function analysis of I2 has been performed using a transient complementation assay: the C-terminal hydrophobic domain is essential for protein stability, and several regions within the N-terminal hydrophilic domain are essential for biological competency. I2 is thus yet another component of the poxvirus virion that is essential for the complex process of entry into target cells.
Variola virus and vaccinia virus are perhaps the most notorious members of the poxvirus family: the former is the etiological agent of smallpox, and the latter has long been used as the vaccine to protect against smallpox. Poxviruses are the only DNA viruses whose life cycle is restricted to the cytoplasm of infected cells; the large repertoire of proteins encoded by the ~200-kb DNA genome of these viruses affords them a high degree of genetic and physical autonomy from host cells. The virions themselves are ~250 by 350 nm in size and contain ~75 proteins which localize to a delimiting and protein-rich membrane, two proteinaceous lateral bodies of unknown function, and a central core (9). The core, in addition to its numerous structural components, contains the viral genome and a transcriptional apparatus that is responsible for mediating early gene expression immediately after virion binding and entry. The binding of virions to target cells involves interactions between several virion membrane proteins (A27, D8, and H3) and glycosaminoglycans (GAGs) on the target cell (4, 5, 7, 21-23, 29) and is also presumed to involve higher-affinity interactions between as-yet-unidentified virion proteins and cellular receptors. Subsequent release of the virion core into the cytoplasm can occur either by direct fusion of the virion and plasma membranes or by engulfment of the intact virion followed by pH-dependent fusion events which result in the release of the core from the endocytic compartment and its deposition in the cytoplasm (4, 31, 51, 54).
Approximately 90 genes are conserved within the genomes of all chordopoxviruses and are therefore viewed as encoding the repertoire of proteins essential for completion of the viral life cycle (60). Through combined genetic, biochemical, and bioinformatic analyses, the roles played by the majority of the proteins encoded by these genes are being elucidated. As might be expected, these proteins collectively mediate virion entry, gene expression, genome replication and maturation, virion morphogenesis, and virion egress. However, a few of the gene products encoded by these 90 conserved genes remain unstudied. Among these, we chose the I2L gene for study because it was predicted to be expressed at late times of infection and to encode a small membrane protein. We therefore reasoned that it might be involved in the biogenesis of the virion membrane, an incompletely understood process which has been of historical interest to our laboratory.
We show here that the I2 protein is expressed at late times of infection, associates with membranes, and is present within mature virions. Moreover, we show that, when I2L is repressed, the biochemical progression of the life cycle and the process of virion morphogenesis are unaffected. However, the specific infectivity of the virions that are produced in the absence of the I2 protein is reduced ~400-fold because of their inability to enter target cells. Thus, I2 is an essential protein that plays a heretofore unknown role in the earliest stage of the infectious cycle.
African green monkey kidney BSC40 cells, mouse L cells, and human thymidine kinase-negative (TK−) 143B cells were cultured in Dulbecco's modified Eagle's medium containing 5% fetal calf serum (Invitrogen, Carlsbad, CA) at 37°C in the presence of 5% CO2. Viral stocks were prepared by ultracentrifugation of cytoplasmic lysates through 36% sucrose; titration was performed on confluent monolayers of BSC40 cells, which were fixed and stained with 0.1% crystal violet in 3.7% formaldehyde at 48 postinfection (hpi). Restriction endonucleases, T4 DNA ligase, calf intestinal alkaline phosphatase (CIP), and Taq polymerase were purchased from Roche Applied Sciences (Indianapolis, IN). Geneticin (G418 sulfate), Lipofectamine 2000, protein molecular weight markers, DNA molecular weight standards, and Texas Red-phalloidin were purchased from Invitrogen (Carlsbad, CA). 14C-labeled protein molecular weight standards were obtained from Amersham/GE Life Sciences (Piscataway, NJ). 32Pi and [35S]methionine were purchased from Perkin-Elmer Life and Analytical Sciences, Inc. (Boston, MA). Ultrapure chemicals, protein A-Sepharose, and protein G-agarose were from Sigma Aldrich (St. Louis, MO). DNA oligonucleotides were synthesized by IDT (Coralville, IA). Monoclonal hemagglutinin (HA) antibody was acquired from Covance Research Products (Princeton, NJ). Antibodies to the A21, A28, and G3 proteins were kindly supplied by B. Moss (NIAID, NIH, Bethesda, MD). The antibody to the A5 protein was kindly supplied by M. Esteban (Centro Nacional de Biotecnología, Madrid, Spain).
The primers used to amplify wild-type, epitope-tagged, and mutant alleles of I2 used in the generation of plasmids and viruses described in this work are shown in Table Table11.
A cassette comprising the 3′ 346 bp of I1, the I2 promoter, the I2 open reading frame (ORF) in which 27 bp encoding the 9-amino-acid (aa) HA epitope was inserted immediately after the initiating ATG codon, and the 5′ 288 bp of the I3L ORF was prepared by overlap PCR using viral genomic DNA as a template. Two initial PCRs were performed using HAI2 1 plus HAI2 B and HAI2 C plus HAI2 4. The products derived from these two reactions overlapped by 17 bp and were combined and used as the templates for a second round of PCR performed with the outside primers HA-I2 1 and HA-I2 4. The PCR product generated was digested with BamHI and ligated with pUCneo (16) plasmid that had been previously digested with BamHI and treated with CIP.
Confluent 35-mm dishes of BSC40 cells were infected with wild-type (wt) virus (multiplicity of infection [MOI] of 0.03) and transfected with 5 μg of supercoiled pUCneo-I1-HA-I2-I3 plasmid DNA at 3 hpi with Lipofectamine 2000. At 16 hpi, G418 was added to 3 mg/ml to select for viruses in which the pUCneo-I1-HA-I2-I3 plasmid had integrated into the viral genome. At 48 hpi, cells were harvested and G418r viruses were isolated by two rounds of plaque purification in the presence of G418. PCR was performed with primers specific for the neomycin resistance (Neor) cassette (NEO) to confirm plasmid integration (16). G418r viruses were subjected to sequential rounds of plaque purification in the absence of drug, allowing for recombination between the tandem repeats of sequences derived from the I1L, I2L, and I3L genes that are present in this intermediate virus. To distinguish viruses containing only the wt allele from those containing the HA-tagged targeted locus, plaque isolates were screened by PCR using primers that flank the I1 3′ and I2 5′ junction (I2 probe 5′ and I2 probe 3′) and generate products of 248 or 221 bp that correspond to the targeted or wt locus, respectively.
To enable inducible expression of the protein I2, we generated vΔindI2, in which an inducible copy of the I2 gene has been placed within the TK locus and the endogenous I2 gene has been deleted.
The pJS4:tetR plasmid (59), based on the pJS4 plasmid (gift of B. Moss, NIH), contains two divergent promoters: one driving constitutive expression of the tetracycline (TET) repressor (tetR) and the other preceding a multicloning site. These two cassettes are flanked by the left and right halves of the TK gene, which enables their insertion into the genome by homologous recombination. The second promoter region was replaced with a fragment representing a TET-regulated copy of the I2 gene under the control of the endogenous I2 promoter. Overlap PCR was used to generate a 281-bp fragment containing the 222-bp I2L ORF, the 19-bp Tet operator sequence, and 40 bp of upstream sequence containing the I2 promoter. Using vaccinia virus genomic DNA as template, PCRs were performed using primers indI2-1 plus indI2-B and indI2-C plus indI2-4; the products of these reactions overlapped by 17 bp and were used together as the template for a second round of PCR performed with primers indI2-1 and indI2-4. The final PCR product was digested with ClaI and HindIII and ligated to pJS4:tetR DNA that had been similarly digested. Plasmids containing the fragments in the appropriate orientation were identified by restriction enzyme analysis; the final plasmid was designated pJS4:tetR↔ indI2.
To achieve insertion of the tetR-inducible I2 cassette into the TK locus, cells were infected with wt virus (MOI of 0.03) and transfected with linearized pJS4:tetR↔ indI2 DNA at 3 hpi. At 48 hpi, the cells were harvested and TK− virus was isolated by two rounds of plaque purification on human TK− cells in the presence of 25 μg/ml bromodeoxyuridine. Insertion of the cassette was confirmed by PCR using primers directed to the tetR sequence (Tetr5′ and Tetr3′) (56). Plaques of the correct genotype were expanded, and this virus was designated vindI2.
Once the inducible I2 gene had been inserted into the TK locus, the final step required for the generation of the inducible virus was the replacement of the endogenous I2 allele with NEO.
PCRs were performed with primers I1pucNeo A and I1pucNeo B, which enabled the amplification of a 300-bp I1L fragment containing Asp718 and BamHI sites at the 5′ and 3′ termini, respectively. Primers I3pucNeo B/H and I3pucNeo 5′ were used to amplify a 324-bp I3L fragment containing BamHI/HindIII and XbaI sites at the termini. The I1L fragment was digested with Asp718 and BamHI, and the I3L fragment was digested with BamHI and XbaI. These products were ligated simultaneously to pBSIIKS vector DNA (Stratagene; La Jolla, CA) that had been previously digested with Asp718 and XbaI and treated with CIP. The resultant plasmid, pBSIIKS-I1-I3, was then used for the next round of cloning. A 1.3-kb fragment containing the Neor gene under the regulation of a constitutive viral promoter was released from pUCneo (16) by digestion with BamHI and HindIII, and the 5′ overhangs were filled in with the Klenow fragment of Escherichia coli DNA polymerase. The plasmid pBSIIKS-I1-I3 was linearized at the internal BamHI site and treated with CIP, and the 5′ overhangs were similarly filled. This linearized plasmid was then ligated with the NEO gene; the resulting plasmid, which contained portions of the I1 and I3 genes flanking NEO, was designated pI2KO.
Cells were infected with vindI2 at an MOI of 0.03 and transfected with linearized pI2KO DNA. G418 was added at 15 hpi; cells were harvested at 48 hpi, and G418r viruses were isolated by two rounds of plaque purification in the presence of G418 (3 mg/ml) and TET (1 μg/ml). The final virus, in which the endogenous allele had been replaced with NEO and the inducible allele was inserted within the TK locus, was designated vΔindI2.
The I2 ORF, or variants thereof, was amplified from vaccinia virus genomic DNA by PCR using the primers indicated below. PCR products were digested with ClaI and BamHI and ligated with pUC1246 vector (56) that had been digested similarly and treated with CIP. (i) To generate pUC1246-I2Δ1-12, a single PCR was performed using primers Δ1-12-5′ and I2-3′. (ii) To generate pUC1246-I2-DK, primers DK-AA-5′ and I2-3′ were used. (iii) To generate pUC1246-I2-EDD, primer pairs EDD-AAA5′ with I2-3′, and I2-5′ with EDD-AAA3′ were used for the initial rounds of PCR. The products were used as templates for a second round of PCR with the outside I2-5′ and I2-3′ primers. (iv) To generate pUC1246-I2-DEK, primer pairs DEK-AAA 5′ with I2-3′ and I2-5′ with DEK-AAA 3′ were used for the first round of PCR. The products were used as templates for overlap PCR using the outside I2-5′ and I2-3′ primers. (v) To generate pUC1246-I2-FI, primer pairs FI-AA 5′ with I2-3′, and I2-5′ with FI-AA 3′ were used for the first round of PCR. The products were used as templates for overlap PCR using the outside I2-5′ and I2-3′ primers. (vi) Generation of pUC1246-I2-YFF was performed with primers YFF-AAA-5′ and I2-3′.
Confluent 35-mm dishes of BSC40 cells were infected with vΔindI2 (MOI of 3) in the presence (+) or absence (−) of TET. At 3 hpi, cells infected in the absence of TET were either left alone or transfected with 4 μg of the indicated plasmid using Lipofectamine 2000. At 24 hpi, cells were harvested and the yield of infectious virus was determined by titration on BSC40 cells. Accumulation of the transiently expressed protein within the infected/transfected cells was assessed by immunoblot analysis using the anti-I2 serum.
To prepare a recombinant immunogen, we constructed a plasmid encoding a trpE-I2 fusion protein which incorporated the hydrophilic portion of I2. Sequences encoding the N-terminal 47 aa of I2 were amplified using vaccinia virus genomic DNA as a template and primers I2-path 5′ and I2-path 3′. The 141-bp PCR product was digested with EcoRI and ClaI and then ligated with pATH11 vector (27) that had been similarly digested and treated with CIP. The resulting plasmid was designated pATH11-I2ΔC′.
E. coli strain HB101 transformants harboring pATH11-I2ΔC′ were induced to express the 41-kDa fusion protein. Cells were lysed, and the insoluble fraction was resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE); the trpE-I2ΔC′ fusion protein was visualized by copper staining, and the excised gel slice was used as an immunogen in rabbits. Antigen injection and serum procurement were performed by PRFAL (Canadensis, PA).
The 222-bp I2 ORF was amplified using viral genomic DNA as a template and the I2-pTM1/pUC1246 5′ and I2-pTM1/pUC1246 3′ primers. These primers inserted terminal NdeI site or BamHI sites, which were used to ligate the insert into pTM1-3XFLAG or pUC1246 vectors (32, 39, 56) that had been similarly digested and treated with CIP.
Confluent 60-mm dishes of BSC40 cells were infected with vTF7-3 or wt vaccinia virus. At 3 hpi, cells were transfected with 10 μg of pTM1-3XFLAG-I2 (vTF7-3 infection) or pUC1246-I2 (wt infection). Cell pellets prepared at 18 hpi were washed with 10 mM Tris (pH 7.4) and 100 mM NaCl and collected by centrifugation. The resulting cell pellets were resuspended in cold 100 mM Na2CO3 (pH 11.5) and disrupted with a Dounce homogenizer. Lysates were incubated on ice for 30 min and then sedimented at 150,000 × g for 30 min. The membrane pellet was resuspended in an equal volume of 100 mM Na2CO3. The supernatant and pellet fractions were acetone precipitated and then resuspended in equal volumes of protein sample buffer.
Proteins were resolved on SDS-polyacrylamide gels and transferred to nitrocellulose (Schleicher & Schuell, Keene, NH) in CAPS buffer [10 mM 3-(cyclohexylamino)-1-propanesulfonic acid, 10% methanol (pH 11.3)]. Filters were probed with primary antisera followed by the appropriate horseradish peroxidase-conjugated anti-rabbit or anti-mouse secondary antibodies. After development with the West Pico enhanced chemiluminescent reagent (Pierce, Rockford, IL), immunoreactive proteins were visualized using Kodak MR film or an Alpha Innotec FluorChem 8800 documentation system.
Confluent monolayers of BSC40 cells were infected with the appropriate virus and metabolically labeled with [35S]methionine for the indicated times; cytosine arabinoside (AraC) was included in the culture medium at 20 μM where indicated. Cells were lysed in phospholysis buffer (10 mM NaPO4 [pH 7.4], 100 mM NaCl, 1% Triton X-100, 0.1% SDS, 0.5% sodium deoxycholate), and clarified lysates were incubated with antibody for 4 h on ice followed by the addition of protein A or G for 1.5 h. Immune complexes were washed twice in lysis buffer and analyzed by SDS-PAGE. Gels were visualized by autoradiography using a Kodak low-emission screen.
BSC40 cells were infected with the virus of interest at an MOI of 2. At 24 hpi, cells were harvested and purified virions were purified from cytoplasmic lysates by ultracentrifugation through 36% sucrose followed by banding on 25 to 40% sucrose gradients. Virions were quantitated by determining the optical density (OD), using the conversion factor of 1 OD at 260 nm (OD260) unit = 1.2 × 1010 virion particles per ml. Infectious viral yield was determined by titration on BSC40 cells.
Banded virions were incubated at 37°C for 30 min in one of the following reaction mixtures: 100 mM Tris (pH 9.0), 100 mM Tris (pH 9.0) with 0.1% NP-40 (vol/vol), or 100 mM Tris (pH 9.0) with 0.1% NP-40 (vol/vol) and 50 mM dithiothreitol (DTT). Soluble and particulate fractions, representing the membrane and core components of the virion, were partitioned by sedimentation (16,000 × g for 30 min at room temperature) and analyzed by immunoblot analysis with an anti-HA monoclonal antibody and antisera recognizing representative proteins of the virion membrane and core.
Confluent 35-mm dishes of BSC40 cells were infected with vΔindI2 at an MOI of 10 in the presence or absence of inducer. At 2, 4, 6, and 8 hpi, cells were rinsed with methionine-free medium (DME-met) and pulsed for 30 min with DME-met containing 100 μCi/ml [35S]methionine.
Confluent 35-mm dishes of BSC40 cells were infected with vΔindI2 at an MOI of 10, in the presence or absence of inducer. As a control, a parallel sample was infected with wt virus in the presence of rifampin (RIF [100 μg/ml]) for the duration of the experiment. At 8 hpi, cells were rinsed with DME-met and pulsed for 30 min in DME-met containing 100 μCi/ml [35S]methionine. Cells were either harvested immediately as described above (pulse) or fed with DME plus 5% fetal bovine serum (FBS) (containing methionine) and harvested at 24 hpi (chase). For both the time course and the analysis of protein processing, samples were fractionated on 10 to 17% SDS-polyacrylamide gels and visualized by autoradiography.
Equal amounts of purified vΔindI2 ± TET or vTetR viral particles were diluted in 50 μl of water, boiled for 5 min, and applied (using a Bio-Dot microfiltration apparatus [Bio-Rad]) to a hydrated ZetaProbe membrane (Bio-Rad). Samples were denatured and renatured in situ, and the blot was then subjected to Southern dot blot hybridization using 106 cpm/ml of a radiolabeled probe representing the vaccinia virus HindIII E and HindIII F genomic fragments (20, 55). The levels of hybridized probe were visualized and quantitated by autoradiography and phosphorimager analysis.
Confluent monolayers of BSC40 cells were placed on ice for 15 min in the presence of 300 μg/ml cycloheximide, which was present in all further steps. Cells were inoculated in duplicate with 10,000 particles/cell of purified I2+ or I2− virions and maintained for 1 h on ice. Cells were then washed three times with phosphate-buffered saline (PBS) and pulsed with neutral fusion buffer (PBS with 10 mM MES [morpholineethanesulfonic acid]and 10 mM HEPES [pH 7.4]) or acidic fusion buffer (PBS with 10 mM MES and 10 mM HEPES [pH 5.5]) for 2 min at 37°C. Cells were then fed with Dulbecco's modified Eagle's medium plus 5% FBS (with cycloheximide) and incubated at 37°C for 3 h. Phase-contrast pictures were captured with a Nikon Coolpix camera.
BSC40 cells were infected at an MOI of 2 for ~18 h and then fixed for 15 min at room temperature in 4% paraformaldehyde-4% sucrose-PBS. Texas Red-phalloidin was used to stain actin, and DAPI (4′,6′-diamidino-2-phenylindole dihydrochloride) was used to detect DNA.
BSC40 cells were grown to confluence in four-well chamber slides and chilled on ice for 10 min prior to infection with 300 particles/cell of purified I2+ or I2− virions. After 1 h, the “no-shift” samples were washed with PBS and fixed with 4% paraformaldehyde in PBS (20 min on ice). “Shifted” samples were washed with PBS containing 300 μg/ml cycloheximide, fed with prewarmed media containing 5% FBS and 300 μg/ml cycloheximide, and incubated at 37°C for 1.5 h before being fixed. Fixed cells were then quenched, permeabilized, and blocked prior to incubation with anti-A17 or anti-39-kDa protein (A5) (generously provided by M. Esteban) sera followed by the appropriate fluorescent-conjugated secondary antibodies (Molecular Probes). DAPI was applied to visualize DNA. Images were taken on a Nikon Eclipse TE 2000-U microscope using a Photometrics Cool Snap EZ camera and Metamorph software. The images shown were selected from a z-series that had been subjected to deconvolution using a nearest-neighbor format.
Confluent 60-mm dishes of BSC40 cells were infected with vΔindI2 in the presence or absence of inducer. At 17 hpi, cells were fixed in situ with 1% glutaraldehyde in PBS for 1 h on ice and processed for conventional transmission electron microscopy. Samples were examined on a Hitachi H-600 electron microscope.
DNA and amino acid sequences of I2 poxvirus orthologs were retrieved from www.poxvirus.org. Sequence analyses were carried out with LaserGene software (DNASTAR, Madison, WI); amino acid alignments were performed using ClustalW. Autoradiographic films and electron microscopy negatives were scanned with an Epson 2400 scanner, adjusted in Adobe Photoshop (Adobe Systems, Inc., San Jose, CA) or Canvas 8.0 (Deneba Systems, Miami, FL), and figures were labeled with Canvas 8.0.
The goal of this work was to analyze the structure and function of the previously uncharacterized protein encoded by the vaccinia virus I2L ORF. I2L is predicted to encode a 72-aa protein with a calculated molecular weight of 8,400. Figure Figure1A1A presents a ClustalW multiple-sequence alignment of the amino acid sequence of the I2 proteins encoded by diverse chordopoxviruses. The first 33 aa of the protein are highly conserved; the aromatic/nonpolar and charged residues marked with the blue and red lines, respectively, were chosen for mutagenesis studies that will be described later in this report. The arrowhead represents the start of an N-terminal truncation mutant which was also analyzed. Each of the orthologs possesses a region of extreme hydrophobic character that begins at residue 48, as indicated by the green line; this region is predicted to be a C-terminal membrane-anchoring domain. This domain is easily seen in Fig. Fig.1B,1B, which represents a Kyte-Doolittle hydrophobicity plot. The blue bar in Fig. Fig.1B1B indicates the portion of the protein used for preparation of the anti-I2 antiserum.
The anti-I2 antibody that we generated showed good specificity and was useful in some analyses of the endogenous I2 protein, but there were assays in which detection of the endogenous protein with this antibody proved difficult. We therefore found it efficacious to construct a recombinant virus that expressed an N-terminally HA-epitope-tagged I2 protein from the endogenous locus (vHA-I2). The growth properties of this virus were indistinguishable from those of wt virus (not shown). To examine the temporal profile of I2 expression, cells were infected with wt or vHA-I2, in either the presence or absence of cytosine arabinoside (AraC), and were metabolically labeled with [35S]methionine from 3 to 8 hpi. Cell lysates were subjected to immunoprecipitation with preimmune serum (Fig. (Fig.2,2, lanes 1 and 2) or anti-I2 (lanes 3 and 4) or anti-HA (lanes 5 and 6) serum. A labeled protein of ~8.4 kDa (from the wt infections) or ~8.7 kDa (from vHA-I2 infections) was immunoprecipitated by the anti-I2 and anti-HA antibodies, respectively. I2 was not found in samples prepared from cells infected in the presence of AraC, indicating that I2 is an intermediate or late protein whose expression is induced following the onset of viral DNA replication.
Sodium carbonate treatment is an efficient means of partitioning integral or lipid-anchored proteins that are tightly associated with membranes away from extrinsic membrane proteins (18). To monitor the predicted association of I2 with membranes, sodium carbonate fractionation was therefore performed on cells infected with wt virus or vTF7-3 (expresses the T7 polymerase) (14, 17). The wt virus-infected cells were transfected with pUC1246-I2, and vTF7-3 infected cells were transfected with pTM13XFLAG-I2. The pellet and soluble fractions were separated by ultracentrifugation and subjected to immunoblot analysis using anti-FLAG, anti-I2, anti-I3, or anti-A17 antibodies (Fig. (Fig.2B).2B). I2 and 3XFLAG-I2 were found exclusively in the pellet, along with a known membrane component, A17 (43, 45, 64). As expected, the single-stranded DNA binding protein I3 (42) was found in the soluble fraction. To address whether I2 is encapsidated in mature virions, virions purified from wt and vHA-I2 infections were subjected to immunoblot analysis with monoclonal anti-HA antibody. As shown in Fig. Fig.2C,2C, the HA-I2 protein is indeed encapsidated. Furthermore, the subvirion localization of I2 was assessed by treating virions with NP-40 in the absence or presence of DTT. The membrane (soluble) and core (pellet) fractions were separated by sedimentation and analyzed by immunoblot analysis (Fig. (Fig.2D).2D). Treatment of HA-I2 virions with NP-40 was sufficient to release the I2 from the virion into the soluble fraction, whereas release of A17 required treatment with both NP-40 and DTT, as we have observed before (32). As expected, the F18 and L4 proteins remained in the core fraction (pellet) after both treatments. Cumulatively, these data show that I2 is expressed at late times of infection and is a component of the virion membrane that can be extracted with detergent alone.
The high degree of conservation of the I2 ORF in diverse poxviruses suggested that it might play an important role in the viral life cycle. To test this possibility, we constructed a recombinant vaccinia virus in which expression of I2 is regulated by the TET operator/repressor system (vΔindI2). The inducible copy of I2 and the TetR gene were inserted into the nonessential TK locus of the genome, and the endogenous I2 allele was replaced by NEO. A schematic representation of the relevant regions of the genome of this recombinant virus is shown in Fig. Fig.3A.3A. As shown in Fig. Fig.3B,3B, plaque formation by vΔindI2, but not by the parental vTetR virus (in which the TET repressor gene alone has been inserted into the TK locus), is dependent on the presence of the TET in the medium. Because the plaque assay measures both the production and spread of infectious virus, we also performed a quantitative assessment of the 24-h viral yield from vTetR and vΔindI2 infections (MOI of 2) performed in the presence or absence of TET. Although equivalent viral yields were obtained from cells infected with vTetR ± TET, ~100-fold less infectious virus was produced during vΔindI2 infections performed in the absence of TET compared to those performed in the presence of TET (Fig. (Fig.3C,3C, left panel). These data indicate that repression of I2 has a significantly deleterious effect on the production of infectious viral progeny during a single round of infection on BSC40 cells. The same phenotype was observed in mouse L cells and human TK− cells (not shown). The conclusion that I2L is an essential gene was further supported by our inability to generate a viral recombinant in which the endogenous I2L gene was deleted and replaced by a selectable marker (NEO) (data not shown).
Having determined that there was a severe decrease in the yield of infectious progeny when the expression of I2 was repressed, we wished to determine which stages of the viral life cycle were affected by the absence of I2. First, the temporal profile of viral gene expression was determined by metabolically labeling infected cells with [35S]methionine at sequential times after infection and visualizing the profile of nascent protein synthesis by SDS-PAGE and autoradiography. The protein expression profiles were indistinguishable whether vΔindI2 infections were performed in the presence or absence of inducer (Fig. (Fig.4A4A).
The fact that late protein synthesis occurred with normal timing and intensity was an indication that viral DNA replication had also occurred, because DNA replication is a prerequisite for intermediate and late gene expression. Although it seemed unlikely that a membrane protein would play a role in the maturation of replication intermediates, we nevertheless examined whether the repression of I2 had any impact on the processing of concatemeric intermediates to monomeric genomes and found that it did not (data not shown).
Since the biochemical milestones in the viral life cycle seemed to be unaffected by the repression of I2, we next turned to the process of virion morphogenesis. First, we investigated whether the precursors of the major virion proteins underwent normal proteolytic processing in the absence of I2; these processing events occur only during the later stages of virion morphogenesis. We therefore performed pulse-chase analyses in cells infected with wt virus (Fig. (Fig.4B,4B, lanes 1 to 6) or vΔindI2 virus (lanes 7 to 10) in the presence or absence of TET. Samples treated with RIF, an inhibitor of morphogenesis and hence of proteolytic processing (1, 34), were included as controls (lanes 5 and 6). The black arrows point to the precursors, whereas the gray arrows indicate the cleaved products. As expected, RIF inhibited processing during wt infections, whereas the inclusion or omission of TET had no effect on wt infections (compare lanes 2, 4, and 6). Most importantly, we observed that proteolytic processing was comparable when vΔindI2 virus infections were performed in the presence or absence of TET (compare lanes 8 and 10). These data indicate that the repression of I2 does not arrest virion assembly at an early stage.
To address the question of virion morphogenesis in more detail, we performed ultrastructural analysis of cells infected with vΔindI2 for 18 h in the presence or absence of inducer. In both cases, the full set of assembly intermediates, including electron-dense virosomes, membrane crescents, immature virions (IV), immature virions with nucleoid (IVN), and mature virions (MV) were observed. (Representative images for vΔindI2 without TET infections are shown in Fig. Fig.4C.)4C.) Morphogenesis was clearly able to proceed to completion in the absence of I2. Although a formal quantitation of intermediates versus MV was not performed, there appeared to be a modest increase in the number of IV and IVN forms and a modest decrease in the number of MV when I2 was repressed. These data suggested that morphogenesis might proceed at a somewhat slower rate when I2 is repressed. However, when cells were infected with and without TET and the viral yields were titrated at 12, 16, 20, and 24 hpi (MOI of 3 [Fig. [Fig.3C,3C, right graph]), we observed that viral yield appeared to plateau at 20 hpi in all cases. Thus, repression of I2 appears to block, rather than delay, the production of infectious virus.
During vaccinia virus infections, a subset of virions become enwrapped in double membranes derived from the Golgi apparatus, traffic to the cell surface on microtubules, and are released by exocytosis (50). Many of these extracellular virions (EV) remain tethered to the cell surface and initiate actin tail formation, which propels the EV toward adjacent cells and facilitates viral spread (49, 50). In our ultrastructural analyses, we observed numerous wrapped MV, also known as intracellular enveloped virions. To confirm that the process of EV formation was not impaired by the absence of I2, we assayed for the formation of actin tails in vΔindI2 infections performed in the absence of inducer. Infected cells were stained with Texas Red-conjugated phalloidin and examined by immunofluorescence microscopy; actin tails were easily observed in cells infected with vΔindI2 in the absence of inducer (Fig. (Fig.4D).4D). As a control, we confirmed that these actin tails were absent when infections were performed in the presence of brefeldin A (not shown), which blocks the formation of EV (58).
Since the assembly of mature virions did not appear to be compromised in the absence of I2, we purified I2-containing and I2-deficient virions and examined their protein and DNA content. As shown in Fig. Fig.5A,5A, I2-containing and I2-deficient virions form a light-scattering band at the same position. The particle yield from vΔindI2 infections performed in the absence of TET was decreased by ~2.3-fold compared to the yield from induced infections, but the yield of infectious virus was reduced by ~1,000-fold (Fig. (Fig.5B).5B). This 400-fold increase in the particle/PFU ratio of the I2-deficient virions indicates that they have an extremely low specific infectivity.
The DNA content of the purified virions was measured by dot blot hybridization (Fig. (Fig.5C),5C), the overall protein content was assessed by silver stain analysis (Fig. (Fig.5D),5D), and the presence of specific virion proteins was examined by immunoblot analysis (Fig (Fig5E).5E). No significant difference was observed between the I2-containing and I2-deficient virions in any of these assays.
Although the I2-deficient virions appeared to encapsidate the normal profile of proteins and DNA, they were not infectious. We therefore wanted to address at which stage this poor infectivity was first manifested and began by assessing their ability to direct early gene expression or induce cytopathic effect (CPE). Cells were infected with equivalent numbers of I2+ and I2− virions (corresponding to an MOI of 10 for the I2+ virions) and then maintained in the presence of [35S]methionine. The morphology of the cells was assessed at 4 hpi, and it was evident that the I2− virions did not induce the morphological changes, such as cell rounding, that were so obvious in cells infected with the I2+ virions (Fig. (Fig.6A).6A). Early gene expression was then assessed by harvesting the cells at 4 hpi and performing immunoprecipitation analyses with antibodies directed against a variety of early viral proteins (anti-E9, -D5, -I3, and -H5) (12, 15, 25, 30, 38, 42). Autoradiographs of the lysates and immunoprecipitates are shown in Fig. Fig.6B.6B. In cells infected with the I2-deficient virions, the synthesis of host proteins persisted and no expression of early viral proteins was detected, in stark contrast to the pattern observed after infection with I2+ virions. Thus, infection with I2-deficient virions was shown to arrest prior to the synthesis of early proteins.
The stages of infection that precede early protein synthesis include binding to target cells, entry into target cells, and transcription of early genes. Because I2 is a membrane protein, we deemed it far more likely that its absence would affect binding or entry rather than transcription. To test this prediction directly, we used the immunofluorescence-based assay that has been established to monitor virion binding and entry (48, 61). Cells were inoculated with 300 particles/cell of I2+ or I2− virions at 4°C for 1 h and then either processed for immunofluorescence microscopy or shifted to 37°C for 1.5 h in the presence of cycloheximide. Virions bound to the cell surface were identified with an antibody to the A17 membrane protein (32, 56), and virion cores deposited in the cytoplasm after virion entry were detected with an antibody to the A5 core protein (10). As shown in Fig. Fig.7,7, the anti-A17 antibody readily detected virions bound to cells after adsorption of either I2+ or I2− virions at 4°C; these virions appear as brightly stained green puncta on the cell surface. These data indicate that I2 is not required for virus attachment. As expected, no staining was observed with the anti-A5 serum under these conditions, as the A5 epitope is not available in intact virions. In the cells that had been infected with I2+ virions and then shifted to 37°C, numerous A5-positive green puncta were observed, indicative of the unmasking of the A5 antigen upon deposition of the viral cores within the cytoplasm. However, no such staining was observed in cells that had been infected with the I2− virions, strongly suggesting that virion entry depends upon the presence of the I2 protein.
Another assay that serves as a surrogate measure of virion entry is the “fusion-from-without” assay, which evaluates the ability of viral inocula to mediate cell fusion at acid pH (19, 48). The validity of this assay has been strengthened by recent demonstrations that proteins required for virion entry are also required for cell fusion, such as A28 and H2 (46, 48). Cultures were therefore inoculated with a high dose of vTetR (parental virus) or I2+ or I2− virions at 4°C for 1 h. The cells were then either left in neutral medium (pH 7.4) or briefly exposed to acidic medium (pH 5.5) and then incubated at 37°C for 3 h in the presence of cycloheximide. Cell fusion was monitored by phase-contrast microscopy; representative images are presented in Fig. Fig.88 (right panel). None of the infections maintained at neutral pH showed any evidence of cell fusion (left panels). However, multinucleate syncytia were evident in cells that had been infected with vTetR or I2+ virions and subjected to the pH 5 pulse (Fig. (Fig.8,8, right panels, arrows indicate large syncytium). In contrast, I2− virions were unable to trigger syncytium formation after acid treatment. These data provide further support for the conclusion that the I2 protein is essential for virion entry. In data not shown here, we also performed the “fusion-from-within” assay, in which CEV deposited on the cell surface during infection are assayed for their ability to trigger cell fusion after an acidic pulse (19, 48). Here again, I2+ but not I2− virions triggered cell fusion (data not shown).
During the course of these studies, numerous other proteins have been shown to play essential roles in vaccinia virus entry, and many of these interact to form a multiprotein entry/fusion complex (EFC) (3, 36, 37, 46-48, 52, 53, 57). We therefore addressed the question of whether repression of I2 might compromise the encapsidation of other proteins involved in virion entry. As shown in Fig. Fig.9,9, I2-deficient virions contained reduced levels of the A21, G3, and A28 proteins; the levels of these proteins were reduced fivefold from what was observed in I2+ virions. This result stands in contrast to the encapsidation of normal levels of other components of the virion membrane (A13, A17, and A14) or core (L4, F10, and A5).
As shown in Fig. Fig.1,1, I2 is a small protein with a hydrophilic N-terminal tail and a C-terminal hydrophobic domain that we assume serves as a membrane-anchoring domain. To gain an initial understanding of which domains and/or residues within the protein might be important for I2's biological function, we established a transient complementation assay. In this assay, cells were infected with vΔindI2 in the absence of TET and transfected with an empty vector or one containing various alleles of I2 under the regulation of a strong late promoter. We examined the wt I2 allele, an allele lacking the C-terminal membrane-anchoring domain (not shown), an allele lacking the N-terminal 12 aa, or alleles containing alanine substitution mutations at positions of clustered charged or aromatic residues. A graphic representation of these data is shown in Fig. Fig.10,10, along with an immunoblot analysis documenting the expression of the various transfected alleles. When TET was omitted from the medium, there was a 3-log difference in the viral yield that was not affected by transfection of an empty vector. The plasmid encoding wt I2 restored virus yield to the level seen when inducer was present; as shown in the immunoblot, the protein expressed from the transfected plasmid is present at levels that significantly exceed that of the endogenous protein, which is undetectable in this experiment. The allele encoding an I2 derivative lacking the C-terminal membrane-spanning domain lacked complementation activity, but we were also unable to detect the protein (not shown). We interpreted these data as evidence that, without the membrane-anchoring domain, the protein was highly unstable. Deletion of the highly conserved N-terminal 12 aa (Δ1-12) also resulted in a complete loss of complementation activity, although this protein was expressed as well as the wt protein. (Note that this protein has an anomalous electrophoretic migration, since it lacks 12 aa but comigrates with the wt protein.) Mutation of any of three clusters of charged residues, E17DD→AAA, D2K→AA, and D33EK→AAA, had no effect on complementation activity. All of these mutants except E17DD were readily detected by the anti-I2 serum. Mutation of the E17DD motif may disrupt the epitope recognized by the serum, as there is complete restoration of virus production in the presence of the mutation but no reactivity in the immunoblot. We next mutated two conserved hydrophobic regions. One allele, in which Y5, F9, and F12 were mutated to alanine, retained wt complementation activity. More importantly, the second allele, which contained F23I→AA substitutions, was unable to complement the I2 deficiency despite being expressed at high levels.
The heretofore unstudied protein I2 is encoded by one of the 90 genes that is conserved in all chordopoxviruses. I2L is predicted to encode a protein of 73 aa, and we have verified that the protein is indeed expressed at late times of infection. Using a recombinant virus encoding HA-tagged I2 from the endogenous locus, we also showed that the I2 protein is packaged in mature virions; fractionation studies indicate that I2 is found in the membrane of purified virions. Given our results, it is somewhat surprising that I2 was not identified as a virion component in recent studies profiling the composition of virions by mass spectroscopy (6, 41, 65). We attributed this discrepancy to the fact that I2 is a very small protein and may also be present at low copy number. The I2 protein can be extracted from the virion membrane with NP-40 alone, as can the A13 protein, which is also a very small protein that contains a single membrane-spanning domain (32, 59). In contrast, the A14 and A17 proteins, which form inter- and intramolecular disulfide bonds and each span the membrane twice, can only be extracted with NP-40 and DTT (32). The integral association of I2 with membranes in vivo and the incorporation of I2 into the virion membrane are likely to be mediated by the extremely hydrophobic domain present at the C terminus of the protein. Because this domain is required for stable expression of the protein, we could not directly test the hypothesis that it serves as a membrane targeting domain for I2.
To address the function of I2 in the viral life cycle, we generated an inducible recombinant in which the expression of I2 from its own promoter was dependent upon inclusion of TET in the culture medium. When I2 was repressed, gene expression, DNA replication, and genome maturation proceeded normally. Moreover, core proteins appeared to undergo normal proteolytic processing, and electron microscopic analysis confirmed that the full spectrum of morphogenetic intermediates was present. Mature virions were readily detected, as were wrapped and enveloped viruses. However, the yield of infectious virus at 24 hpi was reduced by 100- to 1,000-fold in the absence of I2. Virion production was not compromised, but the particle/PFU ratio of the I2-deficient virions was ~400 fold greater than that of those containing I2. Nevertheless, these I2-deficient virions had a wt appearance and contained a normal complement of protein and genomic DNA.
The infectivity defect seen with the I2-deficient virions manifested itself at an early stage. These virions did not induce any CPE, nor were they able to direct the synthesis of early proteins. Furthermore, they were also unable to mediate “fusion from without”: cells inoculated with I2− virions did not induce syncytia after being pulsed with pH 5.5. Acid-induced syncytium formation is now known to mimic viral entry: the brief treatment with low pH mimics the environment presented by the endosomal pathway of entry/fusion that can be utilized efficiently by poxvirus particles (51, 54). This suggestion that I2-deficient virions were defective in virion entry was reinforced by immunofluorescence analysis, which verified that I2-deficient virions retain the ability to bind to target cells but lack the ability to enter cells, as defined by deposition of subviral cores within the cytoplasm.
Binding to target cells is mediated, at least in part, by the association of several virion proteins (A27, H3, and D8) with GAGs on the cell surface as well as the interaction of the A26 protein with laminin (5, 7, 22, 23, 29). There is a presumption that a higher-affinity and/or more specific interaction must take place between as-yet-unknown virion proteins and cellular receptor(s). Neutralizing antibodies have been generated to the H3, A27, and L1 proteins. H3 and A27 are not essential for the production of infectious virus, making them unlikely candidates for receptor-binding proteins. The possibility that L1 may play this role has been difficult to test, because repression of L1 has been shown to block the assembly of mature virions (40). Thus, we cannot be sure how virions that are positive for GAG and laminin interactions, but negative for receptor interactions, would score in the virion binding assay. We are therefore following convention in saying that I2-deficient virions can bind to cells, but we cannot rule out that the possibility that I2 may indeed contribute, either directly or indirectly, to the establishment of a “binding state” that may be a prerequisite for entry.
Production of viral particles that bind to, but cannot enter, cells has recently been associated with mutations in numerous vaccinia virus genes. The A16, A21, A28, G3, G9, H2, J5, L5, and F9 proteins have been shown to be dispensable for virion morphogenesis and cell attachment but essential for core deposition into the host cell (3, 24, 36, 37, 46-48, 52, 53, 57). A conserved feature shared by all but one of these proteins (G3) is the presence of disulfide bonds formed by the virally encoded redox machinery. Eight of these proteins (all but F9) have been shown to associate in a multiprotein complex now known as the EFC (47). The role of individual proteins within the complex is just beginning to be deciphered, but it is known that the EFC does not form in the absence of the virion membrane, nor does it form if either A21 or A28 are absent. H2 and A28 are known to interact directly (35). The F9 protein associates with the EFC at substoichiometric levels but is not required for EFC formation (3). F9 shows significant sequence similarity to the L1 protein, which was previously found to play a role in virion morphogenesis (40). However, very recent data show instead that L1 is dispensable for virion morphogenesis but, like F9, associates with the EFC and is essential for virion entry (2).
The EFC has also been shown to associate with a heterodimeric complex containing the A56 (HA) and K2 proteins, which are known as fusion regulatory proteins. This interaction is mediated by A16 and G9 (62, 63). In the absence of either A56 or K2, infected cells form syncytia spontaneously at neutral pH. These data led to the hypothesis that the A56/K2 proteins prevent syncytium formation by interacting with A16/G9 and blocking EFC-mediated fusion. It seems reasonable to propose that, during the entry of wt viruses, the EFC-A56/K2 interaction is disrupted in the acidic environment of the endosome, activating fusion of the virion and vesicular membranes and resulting in core release.
The data presented here indicate that the I2 protein is the 10th protein that is required for virion entry. Although I2 was not identified as part of the EFC, its presence within the EFC is still possible. It may have been overlooked because of its small size (molecular weight of 8,000), or it may, like the F9 and L1 proteins, be present in substoichiometric levels (3). A fivefold reduction in the levels of some of the EFC proteins was observed within I2-deficient virions, suggesting that I2 might play a role in the stabilization of the EFC or in recruitment of EFC proteins to nascent virion membranes. This 5-fold reduction in EFC proteins seems unlikely to account for the 400-fold decrease in the specific infectivity of the I2-deficient virions. However, it may be that the fivefold reduction does indeed bring the level of the EFC below a threshold needed for efficient entry; it is also possible that the reduction in EFC proteins is greater than our immunoblot analyses indicate. Alternatively, I2 might contribute to virion entry in an EFC-independent manner.
The small size of the I2 protein and the absence of disulfide bonds make I2 quite distinct from the entry proteins that comprise the EFC. One of the striking features that emerges from comparative analysis of the I2 orthologs present in diverse poxviruses is the C-terminal hydrophobic domain. We propose that this region is a membrane targeting domain (MTD) that, like the MTDs of the poxviral H3 protein (11) and other tail-anchored proteins, mediates posttranslational insertion of I2 into nascent virions, although cotranslational insertion is also possible. The N-terminal portion of the protein contains several highly conserved regions. Results from a limited structure/function analysis revealed that three clusters of charged residues within this portion are not essential for I2's biological function. However, the highly conserved N-terminal 12 aa, which are also hydrophobic in character, are required. Similarly, perturbation of a small hydrophobic/aromatic cluster (FI23I24) within this N-terminal domain renders I2 biologically inactive. The hydrophobic and aromatic residues within the N-terminal domain of I2 may mediate protein-protein interactions within the virion membrane or between the virion and the target cell. Alternatively, the N terminus of I2 may participate directly in membrane fusion.
The process through which poxviruses enter cells has recently become an area of intense study. First, it seems striking that at least 11 viral proteins are required for entry. Second, new insights into the mechanism of entry itself are emerging rapidly. Historically, fusion of the virion membrane with the plasma membrane was thought to occur (4, 13, 33). More recently, it has emerged that the majority of virions are internalized into an endosomal compartment, with subsequent membrane fusion and cytoplasmic core deposition occurring only upon compartment acidification (51, 54). The initial internalization of virions has recently been shown to occur via macropinocytosis, a process which is upregulated when the binding of virions to cells induces PAK1-mediated signaling events that result in massive cell blebbing (31). Macropinocytic uptake requires the presence of phosphatidylserine in the virion membrane, suggesting that virion uptake mimics the engulfment of apoptotic fragments. The presence of cholesterol in the plasma membrane of the target cell has also been shown to be important for virion entry (8).
Deposition of the subviral core within the cytoplasm requires fusion of the virion membrane with the membrane of an endocytic compartment (or, less frequently, with the plasma membrane). Because none of the EFC proteins has been shown to have fusogenic activity, we have no insight into how this process occurs. The viral A17 protein has been shown to mediate cell-cell fusion when expressed at high levels on its own in insect cells and to do so when coexpressed with the A27 protein in mammalian cells (26). Because A17 is essential for the early stages of virion morphogenesis (28, 43, 44, 64), it has not been possible to test its role in virion entry. Thus, a role for A17 as a mediator of fusion in the context of infection remains to be formally tested.
The intricacy of the poxvirus life cycle has proven to be illuminating for virologists, immunologists, and cell biologists alike. Recent studies have shown that process of poxvirus entry is surprisingly complex, and further study of how I2 collaborates with many other viral proteins to facilitate the initial interactions of poxvirus virions with their target cells should prove to be of great interest.
This work was sponsored by the NIH/NIAID Regional Center of Excellence for Bio-Defense and Emerging Infectious Diseases Research (RCE) Program. We acknowledge membership within and support from the Region V “Great Lakes” RCE (NIH award 1-U54-AI-057153). P.T. also acknowledges the support of NIH RO1 063630.
We thank Angelika for technical support and Clive Wells for excellent assistance with electron microscopy and acknowledge the invaluable input of all of the members of the Traktman laboratory.
Published ahead of print on 13 August 2008.