INTRODUCTION

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Herpes simplex virus type 1 (HSV-1) is well known as the etiological agent of cold sores, venereal lesions, and neonatal encephalitis (38). Like all herpesviruses, the HSV-1 virion consists of an icosahedral capsid surrounded by a membrane envelope. The viral double-stranded DNA is contained inside the capsid, while a layer of protein called the tegument is found between the capsid and the membrane. The mature capsid is an icosahedral protein shell approximately 15 nm thick and 126 nm in diameter. Its principal structural features are 162 capsomers (150 hexons and 12 pentons) that lie on a T=16 icosahedral lattice. Each capsomer consists of a roughly cylindrical protruding domain that is extended laterally at its proximal end to create the capsid floor layer (3 to 4 nm thick). The floor is the only place where capsomers make direct contact with each other. Capsomers, however, are also connected indirectly by way of the triplexes, trigonal structures (320 in all) that lie above the floor layer with one triplex found at the local threefold position created by each group of three capsomers (7, 25, 27, 36, 41, 42).

VP5, the HSV-1 major capsid protein (molecular weight [MW], 149,075), is the predominant polypeptide component of the capsid; it is the structural subunit of both the hexons and pentons (16). Hexons are hexamers of VP5, while pentons are pentamers. The triplexes are composed of two minor capsid proteins, VP19C and VP23. Most, if not all, triplexes contain one molecule of VP19C and two of VP23 (16).

HSV-1 capsids are formed in the infected-cell nucleus, where they are also packaged with DNA prior to further virus maturation (7, 27). Assembly requires the three capsid structural proteins mentioned above plus a scaffolding protein. In cells infected with wild-type HSV-1, the primary scaffolding protein is pre-VP22a (also called ICP35; product of the UL26.5 gene), although VP21, a cleavage product of the polypeptide encoded by UL26, can also serve effectively as a scaffolding protein (34). During capsid assembly, the scaffolding protein binds to VP5 and forms a core internal to the shell proteins. Pre-VP22a is cleaved to VP22a and exits the capsid at or near the time DNA enters and is not found in the mature virion.

Intermediates in the capsid assembly process have been identified in studies involving use of a cell-free assembly system (14, 15, 35). The system is based on the use of a panel of recombinant baculoviruses (rBV) encoding HSV-1 capsid proteins. It is constituted by mixing extracts of rBV-infected insect cells containing HSV-1 proteins and incubating the mixture in vitro. Studies with the system have demonstrated that capsids are formed by way of partial and complete procapsid intermediates. Partial procapsids are arc- or dome-shaped structures in which a region of capsid shell partially surrounds a region of core. They grow into complete procapsids as the shell is enlarged and closed. The procapsid has T=16 icosahedral symmetry and the same diameter as the mature capsid, but the two differ in several important ways. First, procapsids are spherical, while the mature capsid is icosahedral. Second, the floor layer in the procapsid is not continuous, resulting in a structure that has large gaps between the capsomers. No comparable gaps are found in the mature capsid floor layer. Third, hexons are oval shaped in the procapsid rather than hexagonal as they are in the mature form. Fourth, the procapsid is a more fragile structure. For example, it is disassembled during incubation at 2°C, a treatment that does not affect the integrity of mature capsids.

Although the cell-free system described above has been employed productively to clarify features of the assembly pathway, the presence of insect cell proteins prevents it from being used to address the issue of whether cell-encoded proteins are required for capsid formation. Studies of pairwise interactions between HSV-1 capsid proteins are also complicated by the presence of cell proteins. In order to evaluate the potential involvement of cell-encoded proteins in capsid formation, we have undertaken an effort to purify HSV-1 capsid proteins biochemically from rBV-infected insect cells expressing them and to test the purified proteins for the ability to assemble into procapsids in vitro. The results described here demonstrate that procapsids can be formed readily from purified capsid proteins. They also suggest the nature of VP5-scaffolding protein complexes that may be involved in capsid assembly.

	MATERIALS AND METHODS

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Protein purification. All protein purification was initiated with Sf9 cells expressing the relevant herpesvirus protein(s) after infection with an appropriate rBV vector(s). VP5 and pUL80.5-H were purified from cells infected with BAC-UL19 and BAC-UL80.5-H, respectively, while triplexes were purified from cells coinfected with BAC-UL18 and BAC-UL38. Construction of the four rBV vectors and the methods employed for their propagation and growth on Sf9 cells have been described previously (19, 34). Protein purification was carried out beginning with cell lysates that were produced from pellets of rBV-infected cells. Cell pellets were diluted twofold with phosphate-buffered saline (PBS) containing 20 mM EDTA and protease inhibitors (1 tablet/5 ml; complete, mini; Boehringer Mannheim) and then subjected to three cycles of freezing and thawing to lyse the cells. Lysates were stored in 1-ml aliquots at 80°C until used. The lysate protein concentration was in the range of 15 to 20 mg/ml. Further purification steps were carried out at 4°C beginning with 2 ml of lysates that were thawed and clarified by centrifugation for 5 min at 16,000 × g.

(i) VP5. The clarified lysate was first subjected to centrifugation at 35,000 rpm (43,000 × g) for 30 min in a Beckman TL100 tabletop ultracentrifuge (TLA100.3 rotor). Sufficient saturated ammonium sulfate was then added to the resulting supernatant to yield an ammonium sulfate concentration of 29% saturation. The mixture was incubated at 4°C for 30 min and then centrifuged at 16,000 × g for 10 min to collect the precipitate which contained VP5. The precipitate was dissolved in 3 ml of PBS-10 mM EDTA and desalted on a Bio-Rad EconoPac 10G desalting column (10 ml volume) eluted with 20 mM Tris-HCl (pH 8). The sample was then filtered (0.22 µm pore size; Millipore) and applied to a 1-ml Pharmacia Resource Q anion-exchange column. The column was eluted with a 20-ml gradient of 0 to 1.0 M NaCl prepared in 20 mM Tris-HCl (pH 8), and fractions (1.0 ml each) containing VP5, which eluted at approximately 0.3 M NaCl, were pooled.

(ii) Triplexes. The clarified lysate of BAC-UL18/BAC-UL38-infected Sf9 cells was subjected to high-speed clarification in the TL100 ultracentrifuge, precipitated with ammonium sulfate, desalted, and filtered as described above for purification of VP5, except that the ammonium sulfate precipitate was dissolved in 20 mM Tris-HCl (pH 7.). Triplexes were further purified by chromatography on a 1-ml Pharmacia Resource S cation-exchange column. The sample was applied in 20 mM Tris-HCl (pH 7) and eluted with a gradient of 0 to 1.0 M NaCl in the same buffer. Triplex-containing fractions, as identified by SDS-polyacrylamide gel electrophoresis, were pooled, precipitated with ammonium sulfate, and dissolved in PBS-10 mM EDTA as described above for VP5.

(iii) pUL80.5-H. Sf9 cell lysates containing pUL80.5-H (MW, 39,855) were clarified by centrifugation at 16,000 × g for 15 min. Lysates were then adjusted to 29% saturated ammonium sulfate and incubated at 4°C, and the precipitate was collected as described above for VP5. The precipitate, which contained pUL80.5-H, was dissolved in 0.5 ml of PBS and fractionated further by sucrose density gradient ultracentrifugation. The sample was applied to the top of a 5.0-ml linear 10 to 30% sucrose gradient containing PBS prepared in a 5-ml Beckman Ultra-Clear SW 50.1 ultracentrifuge tube. Gradients were centrifuged at 35,000 rpm (115,000 × g) in an SW 50.1 rotor for 4 h at 4°C. After centrifugation, gradients were separated into 0.4-ml fractions which were examined by SDS-polyacrylamide gel electrophoresis for the presence of pUL80.5-H. The desired fractions were pooled, and pUL80.5-H was precipitated by adjusting the solution to 50% saturated ammonium sulfate. The precipitate was dissolved in sufficient PBS-10 mM EDTA to yield a protein concentration of 2 to 3 mg/ml and used in procapsid assembly and other studies as described below.

Procapsid assembly. Procapsids were assembled in reaction mixtures containing 10 µl of VP5 (1 to 2 mg/ml), 10 µl of triplexes (~1 mg/ml), and 10 µl of pUL80.5-H (2 to 3 mg/ml). Typical reaction mixtures therefore contained approximately 70 pmol of VP5, 84 pmol of triplexes and 500 pmol of pUL80.5-H. Reaction mixtures were adjusted to 25 mM EDTA-10 mM dithiothreitol, and protease inhibitors were added before incubation for 1 h at 37°C. After incubation, reaction mixtures were centrifuged for 2 min at low speed (16,000 × g), and product procapsids were then precipitated by addition of 1 µl of purified monoclonal antibody (MAb) 6F10 (4 mg/ml) (14, 29). Precipitates were allowed to form during 5 min of incubation at room temperature and then harvested by centrifugation at 16,000 × g for 30 s. The precipitate was resuspended in 50 µl of PBS, and the procapsids were dispersed by sonication before further operations were performed.

Sucrose gradient analysis of purified proteins. The oligomeric state of purified capsid proteins and protein complexes was analyzed by centrifugation on 0.7-ml sucrose density gradients. Linear 10 to 30% sucrose gradients containing PBS were prepared in Beckman Ultra-Clear ultracentrifuge tubes (5 by 41 mm (0.7-ml capacity). A 20-µl sample (30 µl in the case of pUL80.5-H) of the specimen to be analyzed was adjusted to 25 mM EDTA-10 mM dithiothreitol, appropriate protein standards were added (see below), and the mixture was layered on top of the gradient, which was centrifuged in a Beckman SW50.1 rotor operated at 35,000 rpm. Centrifugation was for 14 to 18 h in the case of VP5 and triplex specimens and 2.5 to 4 h in the case of pUL80.5-H and VP5-pUL80.5-H complexes. Most gradients were centrifuged at 4°C, although all specimens were also examined at 26 and 34°C. After centrifugation, gradients were separated into 14 equal fractions and an aliquot (20 µl) of each was analyzed by SDS-polyacrylamide gel electrophoresis followed by Coomassie blue staining. Stained gels were scanned while wet on a flatbed scanner (Hewlett-Packard ScanJet IIc; reflected light), and bands were determined quantitatively by use of ImageQuant (Molecular Dynamics) software. The integrated density of individual bands was plotted as a function of the gradient fraction number. The approximate MWs of proteins and protein complexes were determined with reference to protein standards by the use of the treatment of Martin and Ames (10): distance sedimented₁/distance sedimented₂ = (MW₁/MW₂)^2/3. The protein standards employed were bovine serum albumin (BSA; MW, 68,000); -amylase (MW, 200,000), and apoferritin (MW, 443,000).

Cryoelectron microscopy. Freshly assembled procapsids in PBS were concentrated by adding MAb 6F10 (29), and the resulting precipitate was prepared for cryoelectron microscopy by adsorption to a continuous thin carbon film supported on a thick holey carbon film. The drop was blotted onto a thin film, quenched in liquid ethane cooled by liquid nitrogen in a Reichert FC4 cryostation, transferred into a Gatan 626 cryoholder, and observed on a Philips CM200-FEG electron microscope as described by Zlotnick et al. (43). Micrographs were recorded at a nominal magnification of ×38,000 by using minimal electron dose techniques, producing radiation levels of ~8 electrons/Ų.

Image processing. A three-dimensional reconstruction of the procapsid was computed beginning with micrographs that were selected for analysis by visual appraisal (e.g., to assess density of particles and contrast) and by optical diffraction to assess the state of defocus and resolution. Four micrographs whose first contrast transfer function (CTF) zeroes were in the range of 1/22 to 1/24 Å were scanned at 26 µm/pixel on a Perkin-Elmer 1010MG microdensitometer yielding an effective pixel size of ~7 Å. A total of 94 capsid images were processed as previously described (35). The structure was solved by using the polar Fourier transform (PFT) method (1), with our earlier result for extract-assembled procapsids (35) as the starting model. As a control against imprinting the features of the starting model on the reconstruction, the calculation was repeated by using a density map of the mature B capsid (2) as a starting model. Identical results were obtained. After iterative cycles of refinement of orientation angles and origins, the 72 particles with the highest correlation coefficients were selected and a density map was calculated (5) to 25-Å resolution as assessed by the FRC3D criterion (2).

Other methods. Procapsid-containing precipitates were prepared for thin-section electron microscopy by fixation, embedding in Epon 812, and sectioning as previously described (11, 14). For negative-stain electron microscopy, specimens were stained with 1% (wt/vol) uranyl acetate as described by Thomas et al. (32). All thin-section and negative-stain electron micrographs were recorded on a JEOL 100CX transmission electron microscope operated at 80 keV. SDS-polyacrylamide gel electrophoresis was carried out on slabs of 12% polyacrylamide (6 by 9 cm) as described previously (13). Protein bands were identified by Coomassie blue staining and determined quantitatively by scanning as described above. Scans were taken in the dynamic range of the scanner, and the error of the measurement was estimated to be ±5%. Western immunoblotting was performed with similar gels in which proteins were transferred electrophoretically to Immobilon membranes and stained as described by Spencer et al. (29). Specific staining was carried out with (i) rabbit polyclonal antisera specific for VP5, VP19C, or VP23 (generously donated by G. Cohen and R. Eisenberg) or (ii) a rabbit polyclonal antiserum specific for the human cytomegalovirus (HCMV) scaffolding protein (UL80.5 gene product) (19). All specific antisera were used at a 1:5,000 dilution. HSV-1 B capsids, used as reference standards in SDS-polyacrylamide gel electrophoresis and immunoblotting studies, were prepared from BHK-21 cells infected with the 17MP strain of HSV-1 as previously described (16).

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Proteins employed for procapsid assembly. Procapsid assembly was carried out with VP5, triplexes (composed of VP19C and VP23), and a hybrid scaffolding protein called pUL80.5-H (MW, 39,855) (19). The latter consists of the N-terminal 364 amino acids of the HCMV scaffolding protein (UL80.5 gene product) linked to the C-terminal 25 amino acids of the corresponding HSV-1 protein (pre-VP22a; UL26.5 gene product). Our strategy of using the hybrid HCMV-HSV-1 scaffolding protein was based on our experience that although assembly of HSV-1 procapsids was accomplished with the endogenous HSV-1 scaffolding protein in unpurified form in cell extracts (14), our yields of this protein in subsequent attempts at expression and purification were insufficient for the current project. Since procapsids had also been assembled with the hybrid scaffolding protein in vivo and in vitro from cell extracts (19) and this protein proved more tractable in purification trials, it was used throughout the project. All proteins were purified from Sf9 cells expressing the relevant herpesvirus protein(s) as a result of infection with an appropriate rBV vector(s). In each case, protein purification began with a clarified cell lysate produced as described in Materials and Methods.

View larger version (68K): [in this window] [in a new window]   FIG. 1.   SDS-polyacrylamide gel electrophoresis of fractions obtained during purification of VP5 (lanes 2 to 5) and pUL80.5-H (lanes 8 to 10). For reference, HSV-1 B-capsid proteins are shown in lanes 1, 6, 7, and 11. The upper panel shows a gel after staining with Coomassie blue, while the lower two panels show Western immunoblots of the VP5 (left) and pUL80.5-H (right) regions of a comparable gel after staining with a rabbit polyclonal antibody specific for VP5 (lanes 1 to 6) or pUL80.5 (lanes 8 to 10), respectively. Amm., ammonium; Exch. Chrom., exchange chromatography; Suc., sucrose; Purif., purification.

View larger version (24K): [in this window] [in a new window]   FIG. 2.   Column eluate obtained during the ion-exchange chromatography step in the purification of VP5 (a) and triplexes (b). VP5 anion-exchange chromatography was performed on a Pharmacia Resource Q column eluted with a gradient of NaCl (0 to 1.0 M). Triplex chromatography was carried out on a Pharmacia Resource S cation-exchange column eluted with a gradient of NaCl (0 to 1.0 M) as described in Materials and Methods. Optical density at 280 nm was measured. The positions of VP5- and triplex-containing fractions, as judged by SDS-polyacrylamide gel analysis are indicated.

View larger version (37K): [in this window] [in a new window]   FIG. 3.   SDS-polyacrylamide gel electrophoresis of fractions obtained during triplex purification (lanes 1 to 3). For comparison, HSV-1 B-capsid proteins are shown in lane 4. The upper panel shows the gel after staining with Coomassie blue, while the bottom panel shows the results of Western immunoblotting of the VP19C-VP23 region in a comparable gel after staining with rabbit polyclonal antibodies specific for VP19C and VP23. Amm., ammonium; Exch. Chrom., exchange chromatography.

View larger version (133K): [in this window] [in a new window]   FIG. 4.   Electron microscopic analysis of purified pUL80.5-H. Material from the most rapidly migrating band of pUL80.5-H observed during the sucrose gradient purification step (see text) was examined by electron microscopy after negative staining as described in Materials and Methods. Note that pUL80.5-H particles are heterogeneous in size and shape. Bar, 100 nm.

Procapsid assembly. Procapsids were formed by mixing VP5, triplexes, and pUL80.5-H together and incubating them as described in Materials and Methods. The order of component addition did not appear to affect the quality or quantity of the procapsids formed. During the incubation period, the reaction mixture became visibly turbid as procapsids were assembled (data not shown). Turbidity, presumed to be due to procapsids, could not be removed by low-speed centrifugation (16,000 × g) at this stage. Procapsid formation required incubation at temperatures of approximately 26°C or higher. No procapsids formed if reaction components were maintained at 4°C.

View larger version (169K): [in this window] [in a new window]   FIG. 5.   Electron microscopy of procapsids assembled from purified proteins (a, b, and d), procapsids assembled from Sf9 cell extracts containing HSV-1 proteins as described previously (c and e) (14), and control HSV-1 B-capsids (f). Specimens were observed after cryopreservation (a), thin sectioning (b and c), or negative staining (d, e, and f) as described in Materials and Methods. Note that procapsids appear round in all of the preparations, indicating they are spherical. Note also that distinct shell and core layers can be seen in frozen hydrated (a) and thin-sectioned (b and c) preparations. Small procapsids with reduced diameters compared to the majority population (e.g., d; middle procapsids) were observed as a small and variable proportion of procapsids formed both from extracts and from purified components. Bars, 150 nm.

View larger version (120K): [in this window] [in a new window]   FIG. 6.   Three-dimensional reconstruction of procapsids assembled from purified components (b, d, and f). Also shown is the reconstruction computed previously of HSV-1 procapsids assembled in vitro from insect cell extracts containing VP5, VP19C, VP23, and pre-VP22a (a, c, and e; see reference 14). Both reconstructions are computed to approximately 2.5-nm resolution. In all cases, the reconstructions are shown as viewed along the icosahedral twofold axis of symmetry. The reconstructions are shown as outside views (a and b), inside views (c and d) and central thin sections (e and f). Outside views (a and b) are shown before (left half of each capsid) and after (right half) computational removal of MAb 6F10 from the distal tips of the capsomers (29). In the procapsid hemispheres (c and d), the scaffolding protein has been computationally removed from the cavity to expose the inner surface of the shell. Note the overall similarity between procapsids assembled in the two different ways. For example, in both cases, the procapsid is spherical rather than icosahedral, the hexons are oval rather than hexagonal (one is circled in panel a), and the floor layer is rudimentary (the inner shell surfaces in panels c and d). In both reconstructions, the porous nature of the procapsid shell is apparent from the holes (black patches) through it. Note that in both reconstructions, adjacent capsomers do not make direct contact with each other, but rather interact by way of the triplexes (a and b). Note also that capsomer channels appear closed in procapsids assembled from purified components while they are more open in procapsids formed from cell extracts. Bar, 10 nm.

View larger version (30K): [in this window] [in a new window]   FIG. 7.   SDS-polyacrylamide gel electrophoresis of proteins present HSV-1 B capsids (left) and in procapsids formed from purified proteins (right). Proteins were stained with Coomassie blue. Note that the amount of scaffolding protein is greater in procapsids (pUL80.5-H) than in B capsids (VP22a). Ab-H indicates the position of antibody heavy chain.

Oligomeric state of purified components. The oligomeric state of purified procapsid components was examined by sucrose density gradient ultracentrifugation. Samples of purified components were mixed with appropriate protein standards and centrifuged on sucrose gradients, and the gradients were separated into equal fractions as described in Materials and Methods. The positions of procapsid proteins in the gradient were determined by SDS-polyacrylamide gel electrophoresis of gradient fractions followed by densitometric scanning of the stained gel. The integrated density corresponding to each protein was then plotted as a function of the fraction number.

View larger version (27K): [in this window] [in a new window]   FIG. 8.   Sucrose density gradient analysis of purified VP5 (a) and triplexes (b). Gradients were prepared, and the fractions were analyzed by SDS-polyacrylamide gel electrophoresis as described in Materials and Methods. Centrifugation was done at 4°C for 14 (a) and 18 (b) h, respectively. Note that VP5 migrated between the BSA (MW, 68,000) and -amylase (MW, 200,000) markers.

View larger version (27K): [in this window] [in a new window]   FIG. 9.   Sucrose density gradient analysis of purified pUL80.5-H. Gradients contained 5 (top panel), 25 (middle), and 75 (bottom) µg of purified protein, respectively. The gradients were prepared, and the fractions were analyzed by SDS-polyacrylamide gel electrophoresis as described in Materials and Methods. Centrifugation was done for 2.5 h at 4°C. The MW positions shown in the top panel were calculated with reference to the sedimentation of protein standards using the equation of Martin and Ames (10). Note that the sedimentation behavior of pUL80.5-H was strongly influenced by protein concentration.

VP5-pUL80.5-H complexes. Sucrose gradient ultracentrifugation was also employed to examine complexes formed between VP5 and pUL80.5-H. Analysis was carried out at 4°C as described above for purified proteins, except that VP5 and pUL80.5-H were mixed prior to centrifugation. All experiments were carried out at a molar excess of pUL80.5-H over VP5, with pUL80.5-H-to-VP5 ratios in the range of 10:1 to 2:1. Figure 10 shows the results obtained with two specimens containing pUL80.5-H and VP5 at molar ratios of 4:1 and 8:1, respectively. In both cases, pUL80.5-H migrated as a broad band extending across most of the gradient, as it did in the absence of VP5. Migration of VP5, however, was found to be affected by the presence of pUL80.5-H, indicating an interaction between the two proteins. At a 4:1 ratio of pUL80.5-H to VP5, VP5 migrated as a band corresponding to a complex with an estimated MW of 200,000 to 270,000 (Fig. 10a), which was distinct from the band of VP5 sedimented in the absence of pUL80.5-H (arrow in Fig. 10a). The 200,000 to 270,000-MW band was also observed at an 8:1 pUL80.5-H-VP5 ratio, but in this case, the VP5 distribution extended further down the gradient, suggesting the existence of larger complexes, the largest corresponding to an estimated MW of ~500,000 to 600,000 (Fig. 10b). Only trace amounts of VP5 were found to migrate coincidentally with the major 10⁶-MW pUL80.5-H band (Fig. 10b, fractions 10 and 11). No VP5-containing complexes were found outside the region between approximately fractions 3 and 9 under any of the experimental conditions tested.

View larger version (19K): [in this window] [in a new window]   FIG. 10.   Sucrose density gradient analysis of VP5-pUL80.5-H mixtures. Mixtures contained 10 µg of pUL80.5-H plus 10 µg of VP5 (a; ~4:1 molar ratio of pUL80.5-H to VP5) and 30 µg of pUL80.5-H plus 18 µg of VP5 (b; ~8:1 molar ratio). Gradients were prepared, and the fractions were analyzed by SDS-polyacrylamide gel electrophoresis as described in Materials and Methods. Centrifugation was done for 2.5 h at 26°C. The MW positions indicated in the top panel were calculated with reference to the sedimentation of protein standards by use of the equation of Martin and Ames (10). Note that VP5 migrated in the form of complexes with estimated MWs in the range of 200,000 to 600,000, and the most rapidly migrating peak of pUL80.5-H bound VP5 poorly. Vertical arrows indicate the position of VP5 migration in companion gradients in which no pUL80.5-H was present.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Protein purification. It was attractive to initiate protein purification beginning with extracts of rBV-infected Sf9 cells because capsid proteins were known to be competent for procapsid assembly in such preparations. Procapsids are formed after mixing Sf9 cell extracts containing VP5, VP19C, VP23, and pre-VP22a (14, 15). The decision to purify VP19C and VP23 as a complex from cells coinfected with rBV encoding both proteins rather than as separate species was made for three reasons. First, it was observed that VP19C and VP23 participate in assembly only after they associate with each other to form triplexes (28). Neither protein can bind to nascent capsids without the other. Second, preformed triplexes were found to remain intact and to retain their assembly competence during purification. Third, we were unsuccessful in attempts to purify assembly-competent VP19C from Sf9 cells containing it. In extracts, VP19C proved sensitive to proteolytic digestion and the purified protein did not support procapsid formation (17). We suggest, therefore, that association with VP23 serves to stabilize VP19C against proteolysis or other processes that denature or inactivate it.

Protein oligomeric state. Sucrose density gradient analysis of purified VP5 demonstrated that it migrated between the BSA and -amylase markers, suggesting that it is a monomer in solution. No larger structures were observed at any concentration tested. Since MAbs cannot ordinarily precipitate protein monomers (9), the presence of VP5 as a monomer in solution accounts for the observation that it is not precipitated by specific MAbs such as 6F10 unless it is first complexed with scaffolding or scaffolding-plus-triplex proteins (15).

Procapsid assembly. Procapsids formed readily when purified VP5, triplexes, and pUL80.5-H were mixed and incubated. The ability of procapsids to assemble from purified proteins indicates that cell-encoded proteins are not required for assembly. The ability of procapsids to form without involvement of cell proteins is further suggested by the fact that only input virus proteins (and antibody) can be identified by SDS-polyacrylamide gel electrophoresis of the product procapsids (Fig. 7). As with cell proteins, cellular small molecules are also unlikely to play a major role in procapsid assembly from purified components. Purification of each of the three reaction components involves a desalting step that is expected to remove small molecules present in the original cell lysate.

VP5-scaffolding protein complexes. Sucrose gradient analyses of VP5-pUL80.5-H mixtures demonstrated that the two proteins interact readily to form complexes with estimated MWs in the range of 200,000 to 600,000 (Fig. 10). Taking into account the protein composition of the complexes as shown in Fig. 10, we interpreted them to be a population of oligomers in which individual species contain one or two VP5 and one to five or six pUL80.5-H molecules. Little or no free VP5 was observed in the gradients, suggesting that interaction between the two proteins is strong enough that complexes do not dissociate during sucrose gradient centrifugation. Interaction between VP5 and pUL80.5-H has been demonstrated in vitro (19), and interaction of VP5 with the HSV-1 scaffolding protein has been documented both in vivo and in vitro (8, 12, 20, 33). The present studies extend the earlier work by indicating the molecular size and composition of VP5-scaffolding protein complexes.

View larger version (12K): [in this window] [in a new window]   FIG. 11.   Schematic representation of the pathway proposed for HSV-1 procapsid formation. VP5 (gray) and the scaffolding protein (black) are considered to interact to form complexes involving one or two VP5 and one to six scaffolding molecules, as suggested by the results of the sucrose gradient analyses shown in Fig. 10. VP5-scaffolding protein complexes are then cross-linked to each other by triplexes to initiate procapsid formation or to extend regions of growing procapsid shell as found in partial procapsids. The thickening in each scaffolding protein molecule represents the domain by which scaffolding molecules interact with each other (20, 39).