INTRODUCTION

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Kaposi's sarcoma (KS), a multicentric angiogenic tumor of mixed cellularity, is the leading neoplasm of patients with AIDS. Molecular and seroepidemiologic data demonstrate that a rhadinovirus, KS-associated herpesvirus (KSHV), also known as human herpesvirus 8, is the infectious cause of KS (17, 21, 26, 28, 35, 39). KSHV is also associated with primary effusion lymphoma (PEL), a clonal B-cell tumor, and multicentric Castleman's disease, a rare lymphoproliferative disorder (7, 36). Although the identity of the cell population initially infected with KSHV remains unclear, infected but asymptomatic individuals often demonstrate latent virus in their circulating B cells (47) and macrophages (1). In KS lesions, the virus is present in the hallmark spindle cells and in some of the endothelial cells lining vascular spaces (4, 37).

Since most otherwise healthy KSHV-infected individuals remain disease free, a healthy cellular immune response probably keeps active viral replication in check. In contrast, immunosuppression can lead to viral reactivation, replication, and widespread dissemination. It is in this setting that pathogenic progression can ensue. Tumor formation probably requires not only an initial infection of critical numbers of target cells but also the continual recruitment of new cells to replace those lysed from low levels of spontaneous lytic viral replication (11, 37). Similarly, human-to-human transmission, even in the absence of overt disease, presumably relies on horizontal spread of KSHV. Such processes clearly depend on viral replication, including successful formation of infectious particles. As with all herpesviruses, the first structures to appear following the initiation of KSHV replication are the capsidsthe icosahedral particles that fill the nucleus and, when fully mature, harbor the linear viral genome.

Herpesvirus structure and assembly. Studies of alpha- and betaherpesviruses indicate that mature herpesviruses comprise three distinct structural layers plus an inner DNA core. Most information on herpesvirus structure stems from work on these two branches of the herpesvirus family, with the largest portion reflecting data from herpes simplex virus type 1 (HSV-1). In alpha- and betaherpesviruses, the innermost layer is the capsid, consisting of a highly ordered icosahedral structure with a triangulation number (T) of 16 (reviewed in references 14 and 38). Only a portion of the synthesized capsids undergoes viral DNA packaging. When this occurs, the encapsidated DNA is present as a single linear copy of the viral genome and is free of nucleosomes or other DNA binding proteins (2). The KSHV genome also follows this paradigm, assuming a linear form in virions and a circular form during latency (9, 32). A fraction of these DNA-containing capsids, as well as some DNA-free capsids (empty or A capsids [see below]), then acquire first a spherical halo of proteins known as the tegument and second a surrounding envelope with a cadre of integrated proteins. Capsids probably acquire the two outer layers while budding through the nuclear membrane into the cytoplasm (5, 14). Groups of these enveloped particles then transcytose within vesicles toward the plasma membrane. Fusion of the vesicles with the plasma membrane releases the viral particles into the extracellular space.

Capsid architecture. One of the first and essential steps in the cascade of events that leads to viral production is the synthesis of the highly ordered capsid structures. During lytic replication of HSV-1, for example, multiple capsid forms arise (12). These include (i) A capsids (empty icosahedral shells lacking DNA or any other discernible internal structure), (ii) B capsids (shells containing an inner array of scaffolding protein), and (iii) C capsids (shells with packaged DNA and no scaffolding protein). Although their interpretation remains controversial, early pulse-chase experiments suggest that B capsids may mature to C capsids that, in turn, serve as the infectious virus precursors (30). The A capsids probably represent dead-end products derived from either the inappropriate loss of DNA from a C capsid or the premature release of scaffolding protein from a B capsid (without concurrent DNA packaging) (18, 20).

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion References

Cell culture. BCBL-1 is a B-cell line derived from a primary effusion lymphoma that is latently infected with KSHV; no Epstein-Barr virus (EBV) DNA is present in this line (33). The cells were maintained as described previously (33), except that the medium was additionally buffered with 20 mM HEPES (pH 7.3).

Isolation of KSHV capsids. KSHV virions and released capsids were isolated essentially as we have described previously for whole virions (33). In brief, 1 to 2 liters of BCBL-1 grown to 2 × 10⁵ to 3 × 10⁵ cells/ml were treated with both 20 ng of 12-O-tetradecanoyl phorbol 13-acetate (TPA) per ml and 0.3 mM sodium butyrate for 12 to 18 h. The cells were then changed to their standard medium and incubated for another 6 to 7 days. The medium was centrifuged (600 × g for 5 min and then 2,000 × g for 30 min) to sediment the cells, nuclei, and large debris, and then the viral and subviral particles were pelleted from this cleared medium by ultracentrifugation (50,000 × g for 2 h). The pellet was resuspended in DNase buffer (10 mM MnCl₂, 50 mM Tris HCl [pH 7.5]) with 0.03 U of DNase I (Roche Molecular Biochemicals) per ml and incubated for 30 min at 37°C. The reaction was stopped on ice with 20 mM EDTA (pH 8.0). Tris HCl (pH 8.0) and NaCl were then added to give final concentrations of 20 and 250 mM, respectively, and a protease inhibitor cocktail (PILL; Roche Molecular Biochemicals), diluted as specified by the manufacturer was added. Triton X-100 was then added to a final concentration of 2%, and the mixture was incubated overnight at 4°C. This mixture was sonicated in a bath for 15 s and then sedimented (75,000 × g for 30 min) through a 35% (wt/vol) sucrose cushion made up in 20 mM Tris HCl (pH 8)-250 mM NaCl-1 mM EDTA (MTNE). The resulting pellet was resuspended, sonicated as above, loaded (60 µl/gradient) onto a 600-µl 20 to 50% sucrose-MTNE gradient, and centrifuged at 75,000 × g for 40 min. We then collected 40-µl fractions from the top of the gradient for electron microscopy (EM) and protein analyses.

Detection of encapsidated KSHV DNA. KSHV capsid DNA isolation was performed essentially as described previously (33). In brief, the capsid samples were incubated with DNase I as described above, followed by inactivation of the enzyme by the addition of 15 mM EGTA and then digestion with 20 µg of proteinase K (Gibco BRL) per ml in the presence of sodium dodecyl sulfate (SDS) for 1 h at 55°C. After phenol-chloroform-isoamyl alcohol extraction and ethanol precipitation of the encapsidated DNA, the samples were applied to a GeneScreen hybridization membrane (NEN) using a dot blot apparatus and probed for KSHV-specific DNA using a fluorescein-labeled single-stranded probe complementary to the 3' end of KSHV open reading frame 73 (orf73) (random primer fluorescein labeling kit; NEN). The membrane was exposed to Hyperfilm (Amersham), the exposed film was digitally scanned, and the relative intensities of the signals were quantified using GelExpert Software (Nucleotech).

KSHV capsid antibodies. Antibodies to the KSHV MCP were raised in rabbits injected with either internal peptide DQNYDNPQNR (antiserum was a gift from S.-J. Gao) or KAGVQTGSPGN (Animal Pharm Services, Inc., Healdsburg, Calif.). Rabbit polyclonal antisera specific for ORF65 were kind gifts from both S.-J. Gao (unpublished data) and G. Miller (15). Horseradish peroxidase (HRP)-conjugated anti-rabbit antibodies were from Jackson ImmunoResearch Laboratories, Inc. (West Grove, Pa.).

Protein electrophoresis and immunoblotting. Samples containing capsid proteins were separated by SDS-PAGE. Proteins were detected either by staining with Coomassie blue or silver (Silver Stain Plus; Bio-Rad, Hercules, Calif.) or by electroblotting onto polyvinylidene difluoride membranes (Immobilon-P; Millipore) for Western analyses. Immunoblots were preincubated in Block (phosphate-buffered saline [PBS] containing 0.05% Tween 20 [PBST] and 5% nonfat dry milk) for 1 h at room temperature. Primary antibodies were added at the indicated dilutions, and the mixtures were incubated at room temperature for 1 to 2 h and washed three times for 5 min in PBST. The membranes were then incubated with the HRP-conjugated secondary antibody (Jackson Laboratories, Inc.) diluted (1:10,000) in PBST and washed as above. The membranes were then developed using the Renaissance detection kit (NEN) as recommended by the manufacturer, and reactive proteins were detected by autoradiography.

Protein band densitometry. Coomassie blue-stained SDS-polyacrylamide gels underwent digital scanning using a Molecular Dynamics personal densitometer SI. The relative content of individual protein bands was determined using GelExpert Software (Nucleotech).

Mass spectrometry. The details of the mass spectrometric (MS) determination of tryptic peptides are described elsewhere (16). In brief, the gel piece containing the protein band of interest was digested with trypsin and the peptides formed were extracted from the polyacrylamide in 50% acetonitrile-5% formic acid. These extracts then underwent liquid chromatography-MS analysis, and the peptides were eluted from the column by an acetonitrile-0.1 M acetic acid gradient. The digest was analyzed by acquiring full-scan mass spectra (LCQ ion trap mass spectrometer; Finnigan, San Jose, Calif.) to determine the peptide molecular weights and product ion spectra to determine the amino acid sequence in sequential scans. The data were analyzed by database searching using the Sequest search algorithm against the NR database (National Center for Biotechnology Information [NCBI]). Peptides that were not matched were searched against the EST database (NCBI) using Sequest and interpreted manually.

EM. Pellets containing capsids were prepared for EM by fixation, embedding in Epon 812, and sectioning as described previously (19), except that fixation was carried out in 4.5% (wt/vol) glutaraldehyde-4% (wt/vol) paraformaldehyde in 0.1 M sodium phosphate (pH 7.2) overnight at room temperature. Negative staining was performed with 1% (wt/vol) uranyl acetate (40). All thin section and negative-stain electron micrographs were recorded on a Philips 400T transmission electron microscope operated at 80 keV.

Immuno-EM. Samples containing capsids resuspended in MTNE were placed onto carbon-coated copper grids for 45 to 60 s, changed to TBS (100 mM Tris 7.5, 50 mM NaCl), and blocked for 1 h in BB (0.1% fish skin gelatin, 5% goat serum, and 5% bovine serum albumin in TBS). The grids were then incubated for 1 h in antiserum diluted 1:50 in BB and then washed six times for 15 s and then once for 15 min with BB. Colloidal (10-nm-diameter) gold-conjugated goat anti-rabbit immunoglobulin G (IgG) (Electron Microscopy Services) was then added and incubated for 0.5 to 1 h. The grid was washed three times for 15 s with BB and then four times for 15 s with TBS. The grid was then rinsed twice with PBS and fixed with 1% glutaraldhyde in PBS for 2 min. This solution was then removed, and the grid was rinsed three times with TBS. Samples were stained with uranyl acetate and viewed by EM as described above.

Capsid mass determinations. Scanning transmission EM (STEM) determinations of the molecular mass of individual capsid particles were carried out at Brookhaven National Laboratories (Brookhaven, N.Y.). The STEM facility and its analysis capabilities have been reviewed (46). In brief, capsid samples were placed on a thin (2- to 3-nm) carbon film supported by a thick holey film over a titanium grid. Tobacco mosaic virus was also applied as an internal standard. The grid was washed five times with 300 mM ammonium acetate and then ten times with 20 mM ammonium acetate, blotted to a thin layer of liquid, plunged into liquid nitrogen slush, and freeze-dried under vacuum overnight before being transferred to the microscope. The microscope was operated in a dark-field mode in which annular detectors collect nearly all the scattered electrons. A digital image was obtained which consists of 512 by 512 pixels, showing the number of scattered electrons from each pixel. The number of scattered electrons in each pixel is directly proportional to the mass thickness in that pixel.

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

A source of stable KSHV capsids. In a herpesvirus-infected cell, lytic replication often results in nuclei filled with viral capsids. Electron micrographs have documented similar phenomena within the nuclei of PEL cells supporting KSHV replication (29, 33). In studying HSV-1 and cytomegalovirus (CMV) capsid structures, investigators have used such nuclei as the source of capsids. However, as mentioned above, this approach has been unsuccessful with gammaherpesviruses such as EBV. The reason for this is unclear. One possibility is that for gammaherpesviruses many of the intranuclear capsids may be inherently unstable, acquiring additional structural integrity only as they mature and exit the nucleus. In fact, the only successful characterizations of the protein composition of EBV capsids used released virions as the starting material (10).

View larger version (130K): [in this window] [in a new window]   FIG. 1.   Transmission electron micrographs of KSHV capsid species at sequential stages of purification. (A and B) Capsid mixtures pelleted from BCBL-1 medium before (A) and after (B) Triton X-100 extraction. Capsids surrounded with material suggestive of a tegument layer are indicated by wide black arrowheads, empty capsids are indicated by arrows, ring-filled capsids are indicated by white arrowheads, and core-containing capsids are indicated by narrow black arrowheads. (C to E) Gradient fractions enriched for each of these last three capsid morphologies (fractions 7, 9, and 12 from the gradient depicted in Fig. 4). Bar, 0.1 µm.

Isolation and purification of distinct KSHV capsid species. To characterize more carefully the different KSHV capsid species we observed by EM, we next isolated each subpopulation. If the mixture of particles present in the pelleted medium represented the KSHV homologs of the A, B, and C capsids of alpha- and betaherpesviruses, it follows that they would probably have masses sufficiently different to allow their separation by velocity sedimentation. In fact, after sedimentation of the KSHV capsid mixture through a linear sucrose gradient, two closely spaced, light-scattering bands were present toward the center of the gradient and a third, quite faint band was present toward the bottom. Since the A, B, and C capsids of HSV-1 display a similar sedimentation profile under identical conditions (data not shown), we reasoned that the light-scattering bands in our KSHV capsid preparations probably represented the KSHV homologs of these three capsid species.

Protein characterization of KSHV capsids. To determine the protein composition of the capsids, we analyzed the gradient fractions from a series of KSHV capsid preparations by PAGE, concentrating our initial efforts on the two distinct and more abundant capsid species that sedimented toward the middle of the sucrose gradients. Coomassie blue staining of the gels revealed a unique set of protein bands that arose in the same fractions that also contained EM evidence of either empty or ring-filled capsids (Fig. 2A). This set comprised four protein bands (Fig. 2B, bands 1, 2, 4, and 5) that had apparent molecular masses of approximately 140 to 160, 36, 28, and 16 kDa. In addition, a fifth species (labeled band 3 in Fig. 2B), migrating with an apparent molecular mass of approximately 34 kDa, was disproportionately represented in the fraction that contained predominantly capsids with the inner ring structure (Fig. 1D and 2, fraction 11). Importantly, neither Coomassie blue nor silver staining detected this 34-kDa protein in fractions containing only empty capsids (Fig. 1C and 2, fraction 9) despite the presence of the other four capsid-associated protein bands discussed above. The consistently low yield of the capsids with a dense core (the most rapidly sedimenting species) prevented their detection on Coomassie blue-stained gels. However, silver staining of gradient fractions subjected to SDS-PAGE revealed that this underrepresented population of capsids likewise contained the same four proteins shared by the other two capsid species but, as with the empty capsids, lacked the 34-kDa protein (data not shown). The presence of this third population of capsids and its sucrose gradient sedimentation profile relative to those of the other two species was best discerned in Western and silver-stained blots probed for the largest capsid-associated band (see below). We have yet to determine the origin of the other proteins present throughout the sucrose fractions, but their appearance even in regions free of capsids argues that they most probably represent proteins from or adherent to cellular debris of different sizes.

View larger version (61K): [in this window] [in a new window]   FIG. 2.   Velocity sedimentation of KSHV capsids through a 20 to 50% sucrose gradient. (A) Coomassie blue-stained SDS-10% polyacrylamide gel of eight sequential fractions (numbered at the bottom) containing KSHV capsids sedimented through the gradient (note that fraction 13 was underloaded). (B) Fractions 9 and 11 (from panel A) juxtaposed for comparison and with a slightly lighter exposure. The five protein bands that rose above background and sedimented as components of a particle through the gradient are indicated to the right. Note that the capsids in fraction 9 lack band 3. Molecular mass markers (in Kilodaltons) are indicated to the left.

KSHV major capsid protein. Since the single largest contributor to the overall mass of the capsid shell of herpesvirus capsids is MCP, it seemed reasonable to predict that the most prominent capsid-associated protein in our preparations (Fig. 2B, band 1) might represent the KSHV MCP. Sequence homology suggests that orf25 in KSHV encodes the MCP homolog and that the protein, ORF25, would have a molecular mass of 153 kDa (34). We subjected the fraction with the largest amount of band 1 from another capsid preparation to electrophoresis through an SDS-4 to 20% gradient polyacrylamide gel. Better separation of the larger proteins in this type of gel demonstrated more clearly that the band 1 protein migrated with an approximate molecular mass of 150 kDa (Fig. 3A), close to that predicted for ORF25. We confirmed that this band was the KSHV ORF25/MCP by using rabbit polyclonal antiserum that recognizes a unique peptide (KAGVQTGSPGN) encoded by orf25. Figure 3B shows that the MCP peptide-directed serum specifically detected the protein even in the relatively crude pregradient capsid mixture. The antiserum did not react significantly with the other bands in the gel that were detected by Coomassie blue. A second antiserum raised against another MCP peptide (DQNYDNPQNR) likewise specifically reacted with capsid band 1 (data not shown).

View larger version (35K): [in this window] [in a new window]   FIG. 3.   Capsid band 1 is ORF25/MCP. (A) Magnified view of band 1 (arrowhead) stained with Coomassie blue after separation on a 4 to 20% polyacrylamide gel. The band migrates at approximately 150 kDa. (B) Coomassie blue-stained 12% polyacrylamide gel (lane 1) and immunoblot (lane 2) of the capsid mixture prior to gradient purification. The immunoblot was probed with rabbit anti-MCP peptide polyclonal antisera. Molecular mass markers (in kilodaltons) are indicated to the right of each panel.

View larger version (24K): [in this window] [in a new window]   FIG. 4.   MCP profile across fractions from a sucrose gradient reveals distinct sedimentation velocities of the different KSHV capsid species. The intensity of silver staining of MCP (band 1) is plotted against fraction number (fractions were collected from the top of the gradient as indicated). The first peak (fractions 7 to 9) contains empty and ring-filled capsids, and the second, smaller peak (fractions 12 and 13) contains mainly the less abundant core-filled capsids (see the text). Maximal Coomassie blue staining of MCP (fraction 8) was designated 100%. The silver-stained gels containing fractions 1 to 7 and 8 to 16, respectively, are aligned below the graph, and the position of MCP is indicated by horizontal arrows (a vertical black line marks the break between the two gels).

Identification of the 34-kDa capsid protein as the KSHV scaffolding homolog. B capsids of HSV-1 and CMV capsids have a protein scaffolding within their shells. This structure is absent in the other (A and C) capsid species that arise during lytic replication. The presence of the scaffolding endows B capsids with a larger mass than A capsids, so that the former sediment at slightly higher rates through a sucrose gradient. Our EM and sedimentation velocity data support the notion that KSHV lytic replication similarly leads to the accumulation of A and B capsid species. We observed that the two closely spaced bands present in our gradients corresponded to two distinct capsid morphologies present in the fractions from these bands (Fig. 1C and D). The only discernible morphologic difference between the two capsid populations was the inner ring structure. As a result, these capsids would probably have similar protein profiles, differing only by the one or two proteins that would explain the discrepancies in their morphology and sedimentation behavior. In fact, as shown above, both had the common set of four capsid-associated proteins, but the slightly faster-sedimenting capsids demonstrated a unique protein migrating on SDS-PAGE with a mass of 34 kDa. This protein was absent from the lane containing the uppermost gradient band of capsids (compare fractions 9 and 11 in Fig. 2B).

View larger version (51K): [in this window] [in a new window]   FIG. 5.   Rabbit antisera raised against a recombinant protein encoded by the 3' half of orf17 recognizes a KSHV lytic protein in TPA-induced BCBL-1 cells. Overproduced recombinant protein (arrow) from bacterial lysates (lane 1) comigrates with its endogenous counterpart from TPA-induced BCBL-1 cells (lane 2) in Western analyses probed with the rabbit antiserum (anti-ORF17.5). The protein is essentially absent in extracts from uninduced BCBL-1 cells (lane 3). The secondary antibody was HRP-conjugated goat anti-rabbit IgG. Molecular mass markers are shown to the left. Vertical lines indicate where separate lanes from the single gel were juxtaposed for optimal comparison.

View larger version (74K): [in this window] [in a new window]   FIG. 6.   The 34-kDa capsid band 3 reacts with anti-ORF17.5 antiserum. KSHV capsids from gradient fractions 9 and 11 (Fig. 2B) were subjected to SDS-PAGE and then either stained with Coomassie blue (lanes 1 and 2) or immunoblotted and probed with anti-ORF17.5 antiserum (lanes 3 and 4), as in Fig. 5. The 34-kDa ORF17.5 capsid protein (arrow) was present in the gradient fraction containing ring-filled capsids (lanes 2 and 4) but absent in the fraction containing empty capsids (lanes 1 and 3). Molecular mass markers (in kilodaltons) are indicated to the left. Vertical lines between lanes indicate where separate lanes from the gel and Western blot were juxtaposed for direct comparison.

MS identification of the remaining three protein components common to all the KSHV capsid populations. Although immunoblots helped identify the KSHV capsid proteins MCP (ORF25) and scaffolding protein (ORF17.5) (Fig. 2, bands 1 and 3, respectively), we used MS to help assign the appropriate KSHV orfs to the three remaining candidate capsid proteins (Fig. 2B, bands 2, 4, and 5). MS determines both the overall mass and amino acid sequence of tryptic peptide fragments from protein bands removed from Coomassie blue-stained gels (see Materials and Methods). The technique usually provides a sufficient portion of a protein's sequence to allow it to be identified unambiguously (see below). Since the KSHV genome is essentially completed sequenced, identification of the proteins that give rise to MS-derived peptide sequences should, in theory, be possible by searching the protein and DNA databases (22, 34). To rule out the formal possibility that the proteins we identified as capsid associated were instead cellular contaminants, we searched the MS-derived peptide sequences against entire protein databases rather than focusing exclusively on those predicted from the sequence of the KSHV genome.

View larger version (49K): [in this window] [in a new window]   FIG. 7.   Results of MS of KSHV capsid bands 2, 4, and 5. Coomassie blue-stained capsid bands 2, 4, and 5 (Fig. 2B) were subjected to MS (see Materials and Methods). The resultant peptides from each band and their respective sequences (oval shaded regions) are shown superimposed on the deduced amino acid sequences of ORF62, ORF26, and ORF65, respectively.

ORF62 and ORF26 are the KSHV capsid triplex homologs. As discussed above, the capsids of alpha- and betaherpesviruses studied to date contain, in between each adjacent set of MCP capsomers, a heterotrimeric complex referred to as the triplex. Triplexes in these other herpesviruses are composed of two distinct proteins. In HSV-1, for example, each triplex consists of one molecule of VP19C and two molecules of VP23. Although the predicted sequence of ORF62 is only loosely homologous to VP19C, it is 31% identical and 51% similar to the predicted EBV triplex counterpart, the BORF1 gene product. (Amino acid comparison of these EBV and HSV-1 triplex homologs, in turn, reveals 30% identity and 49% similarity.) Likewise, the KSHV ORF26 is 21% identical and 41% similar to the HSV-1 triplex component VP23, whereas it is 49% identical and 69% similar to the predicted amino acid sequence of the protein encoded by the EBV homolog, BDLF1. To test further the notion that ORF62 and ORF26 represent the two components of the putative KSHV triplex, we calculated whether they were present in purified capsids with the expected 1:2 stoichiometry. We estimated the relative amounts of these proteins in both silver-stained and Coomassie blue-stained gels of our capsid preparations (compare, as an example, Fig. 2B, bands 2 and 4, in fractions 9 and 11), quantifying the staining intensity of each (see Materials and Methods) and taking into account the predicted molecular masses of the two proteins. Although these measurements are only rough estimates, given the potential variability in staining between these two proteins, in five capsid preparations we found that the molar ratio of ORF62 to ORF26, in both A- and B-type capsids, was approximately 1:2 (range 1:1.7 to 1:2.2) regardless of which of the two stains was used. These data, coupled with the sequence homologies we describe above, argue for the identification of ORF62 and ORF26 as the components of the KSHV triplex.

The smallest capsid-associated structural protein, ORF65, decorates the outside of KSHV capsids. In the hexomeric capsomers of alpha- and betaherpesviruses, each MCP component is associated with a single small capsid protein in a 1:1 ratio (3). Across the herpesvirus family, these MCP-interacting proteins often show little sequence homology to one another; however, each is the smallest structural component of their respective capsids and each has a highly basic isoelectric point. The predicted EBV counterpart, the 20-kDa small viral capsid antigen (sVCA, encoded by BFRF3), has a pI of 10.8. In the case of HSV-1, the likely structural homolog, VP26, has a mass of 12.1 kDa and a pI of 11.1. Since ORF65, with a predicted molecular mass of 18.6 kDa and a predicted pI of 9.6, is the smallest structural capsid protein present in stoichiometric amounts in our purified KSHV capsid preparations (Fig. 2B, band 5), it seemed reasonable to propose that it is the likely KSHV homolog of sVCA and VP26 (even though the latter has no significant amino acid homology to ORF65) (3).

View larger version (96K): [in this window] [in a new window]   FIG. 8.   Immuno-EM reveals the presence of ORF65 on the surface of purified KSHV B capsids. (A and B) Purified KSHV B capsids were incubated with either anti-ORF65 rabbit antiserum (A) or with unimmunized rabbit serum (B) followed by colloidal gold-conjugated goat anti-rabbit IgG antibodies and then subjected to EM (see Materials and Methods). (C) Purified HSV-1 B capsids were also incubated with the anti-ORF65 rabbit antiserum and then secondary antibody as above. Arrows indicate the single capsids magnified approximately fivefold to the right of each panel in the electron micrographs. Colloidal gold appears as dark, uniformly sized circles. Bar, 0.1 µm.

Only the rapidly sedimenting capsids contain KSHV DNA. Although nonlinear with absolute protein concentration, the relative signal intensities from HRP-conjugated secondary antibodies on immunoblots probed for ORF25/MCP provided a sensitive and rapid means of identifying the fractions containing each of the capsid populations and, particularly, the more elusive core-filled species that sedimented toward the bottom of the gradient (Fig. 1E). Armed with the ability to detect the different capsid populations, we tested the three capsid species for the presence of encapsidated KSHV DNA. Our prediction was that the third gradient band, composed of core-filled capsids, represented KSHV C capsids, each containing a single copy of the viral genome. We isolated potentially encapsidated (DNase-resistant) DNA from each gradient fraction and analyzed it by dot blot Southern analysis using a chemiluminescent probe complementary to KSHV orf73 (see Materials and Methods). The digitized profile of the resultant autoradiograph, superimposed on the profile of MCP shown above (Fig. 4), revealed that the majority of KSHV DNA cosediments with fractions containing the low-abundance, rapidly sedimenting capsid population (Fig. 9). In contrast, the middle gradient fractions containing the more abundant A and B capsid species showed, as expected, an absence of KSHV DNA. These results indicate, therefore, that the capsids shown in Fig. 1E and sedimenting in the second, smaller peak within sucrose gradients are KSHV C capsids.

View larger version (16K): [in this window] [in a new window]   FIG. 9.   Profile of encapsidated DNA from KSHV capsids separated by velocity sedimentation through a sucrose gradient. Relative amounts of KSHV-specific DNA (solid squares) in each fraction from the same gradient shown in Fig. 4 were measured by Southern dot blot analysis (see Materials and Methods). For direct comparison, the MCP profile (open diamonds) from Fig. 4 is also shown. The KSHV DNA peaks in fractions 12 and 13, correlating with the second MCP peak, which contains the core-filled capsids (Fig. 1E).

Capsid mass determinations reflect those predicted by the stoichiometry and molecular masses of the individual capsid components. The relative sedimentation velocities of the three KSHV capsid species suggested that the mass of a B capsid is slightly greater than that of an A capsid but that both are lower than that of the genome containing C capsid. Knowing the orf for each of the five major capsid protein components and therefore the predicted molecular mass of each, we were able to estimate the overall molecular mass of each capsid species. Such calculations were based on the assumption that KSHV capsid geometry parallels that of the alpha- and betaherpesviruses, namely, that each shell contains 150 hexameric and 12 pentameric capsomers, giving a total of 960 molecules of MCP. Likewise, each would also have 320 triplexes (since each triplex joins three MCP molecules, each on a different capsomer). Comparisons of electron micrographs of KSHV and HSV-1 capsids (data not shown) helped support these compositional inferences. Furthermore, our three-dimensional reconstructions of KSHV capsid cryoelectron micrographs demonstrated that KSHV capsids share, with other herpesviruses, the same basic capsomer and triplex architecture (see the accompanying paper [44a]).

View larger version (35K): [in this window] [in a new window]   FIG. 10.   STEM analysis of a mixed population of KSHV capsids. KSHV capsids pelleted from the medium of TPA-induced BCBL-1 cells were subjected to Triton X-100 extraction (but no gradient fractionation) and then analyzed by STEM to determine their masses. Each bar in the graph indicates the frequency with which individually identified particles had a specific mass. Downward arrows indicate the modes and corresponding A, B, and C capsid morphologies.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Beginning with infected-cell supernatant, we have isolated intact KSHV capsids in quantities sufficient for structural and biochemical analyses. To date, this has not been possible for gammaherpesvirus capsids derived from the nuclei of lytically infected cells. Although the reasons for the apparent inability of these intranuclear capsids to withstand the stresses inherent in standard herpesvirus purification procedures remain unclear, recent data suggest that very early forms of HSV-1 capsids (procapsids) possess a more spherical and less angular shell and that this species has less structural integrity compared with the angularized and presumably more mature form that arises subsequently (24, 43). If parallel processes occur in KSHV formation but such maturation is delayed relative to that in the alpha- or betaherpesviruses, then selecting a pool of KSHV capsids at a later stage of development might permit their successful isolation. KSHV capsids that accumulate in the medium of induced PEL cells represent such a population. Supporting this notion, Wu et al. have recently isolated KSHV capsids from the medium of infected cells (48). Our work has shown, in thin-section electron micrographs, that the capsid mixture pelleted from the medium prior to detergent treatment contains a variety of capsid species that are released from TPA-induced BCBL-1 cells (Fig. 1A). This release of previrion forms probably arises from the lysis of cells supporting productive virus replication. Since capsid formation is probably a continuous process during productive infection, it follows that cell death and the accompanying cellular disruption lead to the release of partially formed particles still in the early stages of formation. In addition to mature virions, this would include, therefore, partially and fully formed capsids with and without overlying tegument and envelope layers.

The ability to purify A, B, and C KSHV capsids has allowed us to determine the proteins common to each as well as to delineate the compositional differences among them. Western analyses and MS on preparations of the capsids demonstrated that the basic shell of the KSHV capsid consists of four major protein components, ORF25, ORF62, ORF26, and ORF65. Integrating the present data and those from capsid reconstructions (44a) with the known capsid composition of other herpesviruses, we propose tentative names for the capsid proteins that reflect more accurately their structural roles in the virion (Table 2). When not in conflict with this goal, these names also preserve consistency with the nomenclature of homologous proteins from the other herpesvirus subfamilies. Therefore, we suggest continuing to use MCP (the major capsid protein) for ORF25 but TRI-1 and TRI-2 (the two putative triplex components) for ORF62 and ORF26, respectively, and SCIP (the small capsomer-interacting protein) for ORF65. In particular, we suggest dropping the somewhat misleading designation "minor capsid protein" for ORF26 (and replacing it with TRI-2) since the protein is the second most abundant and third largest component of the capsid shell. In turn, we propose SCIP over sVCA, the name adopted from the EBV nomenclature (15), for two reasons: (i) the EBV nomenclature implies that only this capsid protein is antigenic, an untested hypothesis in KSHV, and (ii) SCIP refers to the likely structural position of the protein in KSHV (as well as that of its homologs in other herpesviruses).

Our analyses have also identified a fifth capsid structural protein, the scaffolding protein, present in stoichiometric amounts only in B capsids and demonstrating an apparent molecular mass of 34 kDa. (We have chosen to use the name "scaffolding protein" rather than "assembly protein" since the former designation is not only descriptive but also the most widely used throughout the alpha- and betaherpesvirus structural and assembly literature.) Interestingly, as with other herpesviruses, the KSHV protease mRNA (encoded by orf17) encodes a long precursor protein with downstream sequence that is colinear and in frame with that of the scaffolding proteins (34, 45). The scaffolding protein, in alpha- and betaherpesviruses, however, derives not from proteolytic processing of the protease precursor but, rather, from the translation of a distinct coterminal mRNA that initiates downstream from the protease termination codon (14). Northern analyses have demonstrated that replicating KSHV gives rise to an approximately 950-base (in addition to the longer pre-protease) mRNA that hybridizes with sequences complementary to the downstream half of KSHV orf17 (45). Recently, Chang and Ganem have mapped the 5' end of this KSHV transcript and have named the corresponding viral gene orf17.5 (8). These data, along with our own (M. Cruise, unpublished results), suggest that KSHV scaffolding expression probably follows the same pattern as seen with the other herpesvirus subfamilies.

Assuming that the scaffolding protein initiates at the first methionine downstream of the protease R site (as in other herpesviruses), the resultant mature scaffolding protein would have a predicted molecular mass of approximately 28.5 kDa rather than the 34 kDa we observed. This discrepancy, however, is not unexpected since the scaffolding proteins from other herpesviruses also demonstrate aberrantly low electrophoretic mobilities. In HSV-1, for example, the scaffolding protein homolog, VP22a, in its mature state has a molecular mass of 31 kDa but an electrophoretic mobility close to 40 kDa (23). The apparent conservation of expression in these regions among the different herpesviruses argues that the systematic designation of ORF17.5 is appropriate for this KSHV protein. However, in light of the present data confirming its structural role in KSHV capsids and in keeping with the nomenclature goals stated above, we suggest the use of the more descriptive name of scaffolding.

The least abundant capsid species that we detected were the C capsids. Although these capsids demonstrated the same four shell proteins as A and B capsids, they were the only capsids that contained KSHV DNA (Fig. 9 and Table 1). Their low abundance in medium from induced BCBL-1 cells may explain, at least in part, the low infectivity found in experiments using this cell line as a source of infectious particles (31). We are presently evaluating other PEL lines to determine if infectivity rates correlate with C capsid and/or virion production.

By comparing the total signal from the encapsidated KSHV DNA on such blots to that from cloned orf73 DNA standards (data not shown), we were able to estimate that the final yield of purified C capsids (i.e., genome equivalents) was approximately 2 × 10⁵ per ml of medium. (Note that there were 2 × 10⁵ BCBL-1 cells/ml at the time of TPA induction.) Since our EM analyses of pelleted medium indicated that approximately 10 to 15% of the total capsids accumulated during the 7 days after TPA induction are C capsids (see above and Fig. 1B), it follows that the final yield of all capsid particles was approximately 1.3 × 10⁶ to 2 × 10⁶/ml. Finally, if 10 to 20% of the BCBL-1 cells (passaged for 2 to 4 months) supported lytic KSHV replication after TPA induction, each such cell gave rise to 30 to 100 total capsid particles.

It is also important to note that such calculations of the absolute yield of capsids from the BCBL-1 cells are probably minimal estimates since a portion of the capsids probably degrade during the prolonged period between TPA induction and capsid harvest. Further, even the relative contributions of A, B, and C capsids to this total may be somewhat distorted. Our work with HSV-1 capsid preparations, for example, indicates that HSV-1 B and C capsids slowly lose their contents over time even at 4°C (W. W. Newcomb, unpublished observations). This phenomenon, if it occurred during our KSHV capsid preparations, would lead to an underestimation of B and C capsid production and an overestimation of A (empty) capsid production.

KSHV capsids as a source of more detailed structural analyses. The identification, stochiometry, and localization of the major structural proteins of alpha- and betaherpesvirus capsids are well known from a combination of biochemical and cryoEM studies (14, 38). The present work provides the first such insights into a gammaherpesvirus, combining KSHV capsid purification with compositional and stoichiometric determinations. Consistent with the structure of capsids from the other two herpesvirus subfamilies, KSHV capsids demonstrate the expected arrangements of capsomers and triplexes as well as the other capsid proteins (namely, SCIP/ORF65 and the scaffolding protein). Further, the results have allowed us to interpret accurately three-dimensional reconstructions from cryoEM analysis in a separate study (44a), which would otherwise rely solely on extrapolation from work on other herpesviruses. Wu et al. recently presented a three-dimensional reconstruction of the KSHV capsid, suggesting that ORF65 was absent in their preparations (48). This conclusion was based on the absence of density near the tip of the KSHV MCP, where, in HSV-1 capsids, the supposed structural homolog (VP26) lies. In contrast, our data demonstrate that ORF65 is in fact present in KSHV capsids. This is clear from four sets of experiments employing (i) PAGE, (ii) Western analyses, (iii) MS, and (iv) immuno-EM. Since the isolation procedure used by Wu et al. is not radically different from ours, loss of ORF65 in their samples as they suggest, although formally possible, seems improbable. A more likely explanation is that ORF65 is present in their capsid samples as well but that it lies in a region distinct from that occupied by VP26 in HSV-1 capsids. Only a through biochemical analysis of their capsids would bear this out. We address this discrepancy more thoroughly in the accompanying structrual study (44a).