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Journal of Virology, September 2007, p. 9396-9407, Vol. 81, No. 17
0022-538X/07/$08.00+0     doi:10.1128/JVI.00986-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Structural Features of the Scaffold Interaction Domain at the N Terminus of the Major Capsid Protein (VP5) of Herpes Simplex Virus Type 1

Eugene Huang,^1, Edward M. Perkins,² and Prashant Desai^1*

Viral Oncology Program, The Sidney Kimmel Comprehensive Cancer Center,¹ Department of Biology and Integrated Imaging Center, Johns Hopkins University, Baltimore, Maryland²

Received 7 May 2007/ Accepted 13 June 2007

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Protein-protein interactions drive the assembly of the herpes simplex virus type 1 capsid. A key interaction occurs between the C terminus of the scaffold protein and the N terminus of the major capsid protein (VP5). Results from alanine-scanning mutagenesis of hydrophobic residues in the N terminus of VP5 revealed seven residues (I27, L35, F39, L58, L65, L67, and L71) that reside in two predicted alpha helices (helix 1^22-42 and helix 2^58-72) that are important for this bimolecular interaction. The goal of the present study was to further characterize the VP5 scaffold interaction domain (SID). Amino acids at the seven positions were replaced with L, M, V or P (I27); I, M, V, or P (L35, L58, L65, L67, and L71); and H, W, Y, or L (F39). Replacement with a hydrophobic side chain did not affect the interaction with scaffold protein in yeast cells or the ability of a virus specifying the mutation from replicating in cells. The mutation to the proline side chain abolished the interaction in all cases and was lethal for virus replication. Mutant viruses with proline substitutions in helix 1^22-42 at positions 27 and 35 assembled large open capsid shells that did not attain closure. Proline substitutions in helix 2^58-72 at either position 59, 65, or 67 abolished the accumulation of VP5 protein, and, at 58 and 71, although VP5 did accumulate, capsid shells were not assembled. Thus, the second SID, SID2, is highly structured, and this alpha helix (helix 2^58-72) is likely involved in capsomere-capsomere interactions during shell accretion. Conserved glycine G59 in helix 2^58-72 was also mutated. G59 may act as a flexible "hinge" in helix 2^58-72 because decreasing the movement of this side chain by replacement with valine impaired capsid assembly. Thus, the N terminus of VP5 and the alpha helices embedded in this domain, as in the capsid shell proteins of some double-stranded DNA phages, are a key regulator of shell accretion and stabilization.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Herpesviruses encode six proteins that come together in a highly coordinated and regulated fashion to form a protective coat surrounding the virus genome. The assembly pathway of herpes simplex virus type 1 (HSV-1) produces three capsid structures detected in sucrose gradients of lysates from infected cells. They are called "A,""B," and "C" capsids, because of their sedimentation profiles in the gradient; C capsids sediment furthest down the gradient (13). "C" capsids contain the virus genome, "B" capsids contain the internal scaffold proteins, and "A" capsids are empty (reviewed in references 26, 28, and 30). A fourth capsid particle, which is sensitive to temperature and sedimentation and which has been observed by ultrastructural analysis of infected cells, is the procapsid and is thought to be the earliest spherical structure formed in HSV-1-infected cells (20, 35).

Seven proteins make up "B" capsids. They are VP5, VP19C, precursor 21 (p21), p22a, VP23, VP24, and VP26. VP5, VP19C, VP23, and VP26 form the outer capsid shell, whereas the scaffold proteins (p22a and p21) occupy the internal space of the capsid (reviewed in references 26 and 30). An important interaction for assembly of an icosahedral capsid structure is that between the scaffold proteins and VP5 (11, 32, 34). This interaction has been confirmed using in vitro and genetic methods (9, 17, 21, 27, 33). The C-terminal 25 amino acids of the scaffold protein are necessary and sufficient for the interaction with the major capsid protein (3, 9, 17-19, 22, 33, 38).

An elegant study by Hong et al. (17) showed that the C terminus of p22a, which interacts with VP5, is a predicted amphipathic helix. The hydrophobic residues of this amphipathic helix are important for binding with VP5 (17). Thus, the hydrophobic residues of VP5 were candidates for this bimolecular interaction. An important clue as to the region of VP5 required for binding to the scaffold protein came from a replication-defective mutant virus that contained a blocked maturation cleavage site in the scaffold protein (23). Second-site revertants of this mutation were easily isolated, and almost all mapped to the gene encoding VP5 (UL19). Many of these second-site mutations mapped to the N terminus, within the first 100 amino acids of this large polypeptide (10, 37). Protein analytical programs were used to identify a hydrophobic region in the N terminus of VP5 (31). Based on these findings an alanine scanning mutagenesis approach was used to change 24 hydrophobic residues in this region (36). The mutants were analyzed for interaction with p22a in the yeast two-hybrid system. Only alanine substitutions at I27, L35, F39, L58, L65, L67, and L71 abolished the interaction with the scaffold protein (36). These amino acids are embedded in two predicted alpha helices in this binding domain, helix 1 (amino acids 22 to 42 [helix 1^22-42]) and helix 2 (amino acids 58 to 72 [helix 2^58-72]) (1, 29). We introduced all 24 mutations into the HSV-1 genome using a relatively quick marker rescue, marker transfer assay that was originally devised by Stanley Person for the transfer of glycoprotein B mutants into the virus (8). All seven mutants that did not interact with the scaffold protein were not capable of growth on Vero cells. Ultrastuctural analysis of the mutant viruses showed visually the effect of disrupting the VP5-p22a interaction. In the nuclei of infected cells large capsid shells that could not attain closure were detected. These structures were formed by the unregulated accretion of capsomeres with each other, indicative of the importance of the interaction between VP5 and p22a for icosahedral capsid shell formation (36).

An important question is whether there is a requirement for a specific hydrophobic group at these seven sites or whether the change to alanine alters the structural positioning of the binding domain. The scaffold interaction domains (SID) reside in the two alpha helices, helix 1^22-42 and helix 2^58-72, predicted by cryo-electron microscopy (cryo-EM) (39) and protein structure analyses (1, 29). The goal of the present study was to target for mutagenesis these seven hydrophobic residues to probe the structural features of the VP5 hydrophobic interactive surface and the requirement of a specific hydrophobic group or contact position and analyze the contribution of a conserved glycine at position 59, which resides in helix 2^58-72 and which may be important for the configuration of the scaffold binding domains.

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Cells and viruses. Vero cells and transformed Vero cell lines were grown in minimum essential medium-alpha medium supplemented with 10% fetal calf serum (Gibco-Invitrogen) and passaged as described by Desai et al. (7). C32 cells complement the growth of VP5 (UL19) and VP23 (UL18) mutants (23), whereas 31 cells complement the growth of only VP5 mutants (36). Virus stocks of KOS (HSV-1) and the mutant viruses were prepared as previously described (7). K18-19G is a mutant virus that contains a deletion in both UL18 (VP23) and UL19 (VP5) genes (36). This virus required the C32 cell line for propagation and served as the recipient genome for all marker rescue (of VP23) and marker transfer (of VP5 mutants) experiments. K5R was the marker-rescued virus of K18-19G, and its replication properties as well as those of K18-19G have been previously described (36).

Plasmids. The template for mutagenesis of the VP5 N terminus was a plasmid designated pKVP5N (36). This plasmid encodes the N-terminal 226 amino acids of VP5.

Mutagenesis. Site-directed mutations were made by the "QuikChange" mutagenesis protocol (Stratagene). The use of this method is described in detail by Walters et al. (36). Positive clones carrying the mutation were confirmed by sequence analysis. The mutant VP5 coding sequence was then cloned into the yeast two-hybrid vector containing the VP5 coding sequence (pGBT9-VP5) as an EcoR1-BspE1 fragment and into the genomic plasmid pKKI as an Age1-HindIII fragment (36), that is, the wild-type sequences were replaced with the sequence containing the site-directed mutation.

Yeast two-hybrid assays. The protocol for yeast two-hybrid assays is described by Desai and Person (9). Sequences encoding VP5 were fused to the yeast Gal4 DNA-binding domain (pGBT9-VP5), and the gene encoding p22a was fused to the Gal4 transactivation domain (pGAD424-22a) (9). Both plasmids were cotransformed into SFY526 cells, and the transformants were allowed to grow on nitrocellulose membranes in order to assay each mutation using the X-Gal (5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside) filter assay (5).

Marker transfer and marker rescue. Marker rescue/marker transfer assays were performed as described by Person and Desai (23). C32 cells (1 x 10⁶) in 60-mm dishes were cotransfected with K18-19G purified DNA (2 µg) and linearized pKKI mutant plasmids (0.1 to 1 µg). The transfection progeny harvested 3 days later were titered on Vero, C32, and 31 cell monolayers to determine the growth properties of each mutant virus.

Radiolabeling and SDS-PAGE. Procedures for radiolabeling of infected cells and sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis are described by Person and Desai (23).

Sedimentation analysis of capsids. Sedimentation analysis of capsids from infected cells was performed as described by Desai et al. (6), Person and Desai (23), and Walters et al. (36).

Electron microscopy. Vero cells (1 x 10⁷) in 100-mm dishes were infected at a multiplicity of infection (MOI) of 10 PFU/cell. Samples for transmission EM (TEM) were prepared essentially as described previously (15). Samples were fixed (3.0% formaldehyde, 1.5% glutaraldehyde, 5 mM CaCl₂, 5 mM MgCl₂, and 2.5% sucrose in 0.1 M sodium cacodylate at pH 7.4) for 1 hour, followed by three 15-min washes in 0.1 M sodium cacodylate. They were then postfixed in Palade's 1% osmium tetroxide for 1 hour on ice, rinsed in deionized water, and en bloc stained in Kellenberger's uranyl acetate overnight. Samples were rinsed in deionized water and dehydrated with a graded series of cold ethanol washes (75%, 95%, 100%) and three washes in 100% ethanol at room temperature for 15 min each. Dehydration was followed by two 5-min exchanges in propylene oxide, followed by an overnight exchange with 50% propylene oxide-50% Epon under vacuum. The samples were infiltrated with 100% Epon under vacuum overnight, embedded in fresh Epon, and polymerized at 60°C for 24 to 48 h. The embedded samples were sectioned with a diamond knife on a Leica Ultracut UCT ultramicrotome, collected on carbon-Formvar-coated grids, poststained with uranyl acetate and lead citrate, and analyzed using a Philips EM 410 or EM 420 transmission electron microscope (FEI Co., Hilsboro, OR) operated at 100 kV. Images were captured on a Soft Imaging Systems Megaview III digital camera (Olympus Soft Imaging Solutions Corp., Lakewood, CO).

For negative-stain analysis, 400-mesh nickel grids coated with Formvar were floated on sucrose gradient fractions, floated on 5 drops of double-distilled H₂O, and then stained with 2% uranyl acetate. The samples were then air dried, and images were captured on a Philips EM 410 or EM 420 transmission electron microscope (FEI Co., Hilsboro, OR) operated at 100 kV.

Data and figure preparation. Autoradiographs were scanned at 600 dots per inch in Adobe Photoshop, and figures were compiled in Photoshop. Electron micrographs were captured as 12-bit images (16-bit tiff files) and exported as 8-bit tiff files, which were adjusted for brightness/contrast and resized in Adobe Photoshop. TEM and negative-stain EM figures were done in Adobe Illustrator.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Mutagenesis of residues in the N terminus of VP5 that are important for scaffold interaction. Previous studies showed that residues in two small domains in the N terminus of VP5 were important for interaction with the scaffold protein (36). In this study the amino acids in the first domain, I27 and L35, as well as the four residues in the second domain L58, L65, L67, and L71, were replaced with another hydrophobic side chain to determine the importance of a specific hydrophobic residue and also with a proline side chain to probe the structure of the interactive domain. Substitutions at F39 were made to identify again the importance of the hydrophobic residue and the contribution of a large aromatic side chain to this interaction. A highly conserved glycine at position 59 was also changed to alanine, valine, or proline. Finally, a lysine walk was performed on residues L24, V29, F36, V42, V68, L73, and V80, which are embedded in these hydrophobic domains; this was done to confirm the previous mutagenesis study as well as to uncover additional hydrophobic contacts. All substitution mutations were made using the "QuikChange" mutagenesis protocol (Stratagene). Each mutation was transferred into the yeast two-hybrid vector, pGBT9 (12), and also into the genomic clone pKKI for introduction into the virus genome. The residues targeted and the amino acid changes are shown in Fig. 1, as are the two predicted alpha helices, helix 1 and helix 2. In all of the tables and figures, the VP5 residue number and two amino acids designate the mutant viruses. The amino acid preceding the number is the wild-type residue, and the amino acid following the number is the altered amino acid.

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FIG. 1. The amino acid sequence of the N terminus of VP5. The amino acid sequence of VP5 (strain KOS) from residue 20 to 83 is shown. The amino acid changes are shown below the targeted residues. The two predicted alpha helices are shown above (black coil). Helix 1 spans from amino acid 22 to 42, and helix 2 spans from 58 to 72 (29). Residues that are in boldface are conserved in all herpesviruses examined (HSV-1 and HSV-2, pseudorabies virus [PRV], equine herpesvirus 1 [EHV-1], bovine herpesvirus 1 [BHV1], varicella-zoster virus [VZV], turkey herpesvirus 2 [THV-2], human herpesvirus 8 (HHV8), EHV-2, Epstein-Barr virus, murine herpesvirus 68, alcelaphine herpesvirus 1, HHV6, HHV7, HCMV, simian cytomegalovirus, and murine cytomegalovirus). The residues that are underlined are conserved in the alphaherpesviruses (HSV-1, HSV-2, PRV, EHV-1, BHV1, THV-2, and VZV).

Yeast two-hybrid analysis of the VP5 mutations. All of the mutations were analyzed using the yeast two-hybrid assay (12) to determine the effect of these changes on the interaction with the scaffold protein. The X-Gal filter assay was used to analyze the potential of each VP5 mutant for interaction with the scaffold protein (Tables 1 and 2), as judged by colonies that turn blue.

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TABLE 1. Interaction of the VP5 mutants with the scaffold proteina

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TABLE 2. Interaction of the VP5 mutants with the scaffold proteina

As shown in Table 1, the mutant proteins produce three different results. There are those that interact with p22a with wild-type kinetics of blue color development; most of the conservative hydrophobic substitution mutations produced this phenotype (Table 1), as well as the G59A mutation (Table 2). There are those that have a decreased interaction with p22a, as judged by the time it took for blue color development; the valine substitutions at I27, L35, L58, and L67 as well as the F39W mutation produced these phenotypes (Table 1). Finally, there are those that fail to interact with p22a, as determined by the absence of blue color upon prolonged incubation; all the proline substitution mutations as well as mutations F39H, F39Y, and G59V produced this phenotype (Tables 1 and 2). Lysine substitution at residues L24, V29, V42, L73, and V80 did not alter the interaction with the scaffold protein; however, lysines at positions F36 and V68 abolished the interaction with p22a in this assay (Table 2). These two amino acids reside in the previously identified SID.

Introduction of the VP5 mutations into the virus genome and replication properties of the mutant viruses. We used the marker rescue/marker transfer assay (36) to facilitate the introduction of all 38 VP5 mutations into the virus genome. For each mutant, the transfection progeny were plated onto C32, Vero, and 31 cell lines. The virus that predominantly plaqued on C32 cells (UL18/UL19 transformed) (23) was the parental virus, K18-19G. Viruses that plaqued on 31 cells (UL19 transformed) (36) were viruses in which the UL18 mutation was rescued and the UL19 mutation transferred into the UL19 locus. In many cases viruses were also observed to plaque on Vero cells, indicating that the mutation was not lethal. All the mutant viruses were plaque purified prior to characterization. Most of the VP5 mutants were capable of growth on noncomplementing Vero cells (Table 3). Mutations that were lethal for virus replication, as judged by the requirement of the 31 cell line (VP5-expressing cells) for virus propagation, are also shown in the Table 3. For these replication-defective viruses, a PCR product, which spanned from the start of the VP5-encoding gene to codon 153, was amplified using the virus DNA and then sequenced. This confirmed the introduction of each lethal mutation into the virus genome. The data show that there was a good correlation between the ability of a mutant protein to interact with the scaffold protein and its ability to replicate on Vero cells (Tables 1 to 3). There were two exceptions to this general observation, the G59A and V80K VP5 mutants. Both interacted with p22a in the yeast two-hybrid assay; however, the mutant viruses encoding these changes failed to replicate or plaque efficiently on Vero cells. In the yeast two-hybrid assay, VP5 molecules with these mutations (G59A and V80K), like wild-type VP5, did not interact with a scaffold protein that did not express the C-terminal 25 amino acids, as judged by the absence of blue color of the yeast transformants (data not shown). This result indicated that these proteins specifically interact with the C terminus of p22a. The virus encoding VP5 with the G59A mutation (G59A virus) was observed to form very small plaques on Vero cell monolayers upon prolonged incubation. These data indicated that, in these viruses (G59A and V80K), VP5 function was somehow compromised, a phenotype, which was not detected in a bimolecular assay but was revealed in the infected cell during the assembly of a complex structure.

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TABLE 3. Replication properties of the VP5 mutant viruses

ICP5 (VP5) synthesis and accumulation in mutant infected cells. The expression and accumulation of the ICP5 polypeptide by all the different mutants were examined by SDS-PAGE of radiolabeled infected-cell lysates (Fig. 2). Only the mutants that encode lethal mutations in VP5 were examined. Proline substitution mutants, especially those with mutations in the first interaction domain (top panel), express and accumulate levels of ICP5 comparable to K5R and KOS (Fig. 2). K5R was the marker-rescued virus of K18-19G and has been previously described (36). Proline substitution mutations in the second domain at positions G59, L65, and L67 abolished accumulation of ICP5 (Fig. 2, second panel). Proline substitution at L58 also resulted in a significant decrease in the levels of ICP5 accumulation. An alanine substitution at this position also resulted in reduced levels of ICP5 accumulation (36). Substitutions at F39 (H/Y) and G59 (A/V) did not affect ICP5 accumulation (third panel). The shutoff of host protein synthesis was more pronounced for KOS then the other viruses in the second and fourth panels due to the differences in the working stock titers. Pulse-chase experiments were carried out to determine the stability of the G59P, L65P, and L67P mutant proteins; however, ICP5 was not detected in pulse-labeled cells either (data not shown). Mutant viruses encoding a lethal lysine substitution in the VP5 N terminus all accumulate wild-type levels of ICP5 (bottom panel).

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FIG. 2. Accumulation of ICP5 in the mutant-infected cells. Vero cell monolayers (2.5 x 105 cells) were either mock infected (MI) or infected with KOS (wild-type), K5R (rescued-wild-type), and the mutant viruses at an MOI of 10 PFU/cell. Infected cells were radiolabeled with [35S]methionine from 9 to 24 h postinfection. The cell pellet was solubilized in 2x Laemmli sample buffer, and the proteins were analyzed by SDS-PAGE (9% acrylamide). The autoradiographs obtained following exposure of the dried gels to X-ray film are shown. The VP5 residue number and two amino acids designate the mutant viruses. The position of the ICP5 polypeptide is shown on the right of the panels. The first and third panels are from the same experiment, and the second and fourth panels are from the same experiment.

Phenotypes of the mutant viruses. In order to determine the nature of the mutation and its effect on capsid assembly in infected cells, we carried out three procedures for each virus. Infected cells were harvested 18 h after infection and processed for TEM. Infected cells were also metabolically labeled with [³⁵S]methionine and the lysates sedimented through sucrose gradients. The fractions from the sucrose gradients were analyzed by SDS-PAGE. Similar gradients were also performed and the fractions analyzed by EM following negative staining (see Materials and Methods for more details). The following sections address the different mutant classes separately for clarity.

(i) Mutations at I27, L35, and F39. Mutation of I27 and L35 to P abolished replication of virus on Vero cells. This was also seen with viruses encoding mutations F39H and F39Y. In I27P and F39H virus-infected cells icosahedral capsid structures that were evident for wild-type-infected cells (Fig. 3D) were not observed in the nuclei of these infected cells by TEM (Fig. 3). Rather, open capsid shell structures were detected in the nucleus (A and C). These structures are the same as the shells we have observed in cells infected with the scaffold null mutant virus or in cells with mutations in the VP5 N terminus (11, 36). The features of the shell structures were evident in the negatively stained image for I27P virus (Fig. 3B); the structures were determined by EM analysis of sucrose gradient fractions of infected-cell lysates. These shells have an open configuration unlike that of KOS capsids analyzed in a similar manner, which have a closed icosahedral configuration (E). Similar observations were made for L35P and F39Y virus-infected cells by TEM analysis as well as EM of sucrose gradient fractions (data not shown). SDS-PAGE analysis of sucrose gradient fractions showed the cosedimentation of shell proteins VP5, VP19C, VP23, and VP26 (Fig. 4). Data for lysates derived from L35P and F39Y virus-infected cells are shown here (Fig. 4), and similar observations were made for I27P and F39H viruses (data not shown). This is the typical profile of open shells; the absence of the scaffold protein in the gradient fractions was the most striking observation (compare fraction 7 of KOS with similar fractions of the mutant viruses). These cosedimenting capsid shell proteins were observed in a large number of the fractions, probably because of the different sizes and shapes of these shells. In fractions of sucrose gradients derived from wild-type (KOS)-infected cells, cosedimentation of the capsid proteins that comprise C (fraction 4), B (fraction 7), and A (fraction 9) capsids was observed (Fig. 4).

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FIG. 3. Mutations at amino acids I27 and F39. Vero cells were infected with KOS (D and E) and the I27P (A and B) and F39H (C) mutant viruses. Infected cells were harvested 18 h following infection and processed for TEM (A, C, and D). Open capsid shells (arrowheads) were seen in the nuclei of the mutant-infected cells. In KOS-infected cells, a large array of capsids containing a variety of internal structures was seen (the whole array is indicated by arrowheads). In addition, DNA-containing enveloped capsids were evident in panel D. In panel B sucrose gradient fractions of I27P virus-infected cell lysates were negatively stained to reveal the detailed structure of the open shells. Gradient fractions of KOS (B capsids) were similarly examined to reveal closed structures (E). Bars, 0.5 µm. The plasma membrane (pm) and nuclear envelope (ne) are indicated.

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FIG. 4. Sedimentation analysis of the L35P and F39Y mutant viruses. Vero cell monolayers (1 x 107 cells) were infected at an MOI of 10 PFU/cell and labeled with [35S]methionine from 8 to 24 h postinfection. Total cell extracts were prepared and layered onto 20 to 50% sucrose gradients. Following sedimentation, fractions were analyzed by SDS-PAGE (15% acrylamide). The autoradiographs obtained following exposure of the dried gels to X-ray film are shown. Fraction 1 represents the bottom of the gradient. The positions of the capsid proteins are indicated on the right of each panel. Cosedimentation of proteins that make up the three intranuclear capsids (C, B, and A) was observed in fractions of KOS lysates (marked at the bottom of the top panel), whereas only cosedimentation of the shell proteins was seen in the gradients of the L35P and F39Y mutant viruses. Molecular mass standards were visible only for KOS and are in lane M (220, 97.4, 66, 46, 30, and 14.3 kDa).

(ii) Mutations at amino acids L58, L65, L67, and L71. Conservative substitutions at these residues allowed the viruses encoding these mutations to replicate on Vero cells. The proline substitutions were severely impaired. Whereas viruses encoding L58P and L71P accumulated ICP5, L65P and L67P viruses did not. When cells infected with the L58P virus were analyzed, no evidence of capsid shells was found, as judged by all three assays described above (data not shown). Similarly, in L71P virus-infected cells no capsid shells in the nucleus or in sucrose gradients of infected-cell lysates were detected (data not shown).

(iii) Mutations at the conserved glycine at position 59. This is one of the few highly conserved amino acids in this N-terminal domain. This amino acid was not analyzed in the previous study, but its conservation and location (in helix 2^58-72) made it an important target for mutagenesis. A change to alanine enables VP5 to interact with the scaffold protein, but virus replication was impaired, in that this virus formed very tiny plaques on Vero cells. When cells infected with this virus were examined by TEM, closed capsids were seen, as well as DNA-filled enveloped capsids (Fig. 5A). Some of the scaffold-containing capsids were also enveloped (Fig. 5A). In sucrose gradients of G59A virus-infected cell lysates the SDS-PAGE data (Fig. 6, top panel) revealed all three intranuclear capsids, that is, the radioactivity observed in fractions 3, 7, and 9 correspond to the polypeptide compositions of C, B, and A capsids, respectively. This profile was similar to that of the marker-rescued virus K5R (Fig. 6, bottom panel), although the levels of capsids, as judged by radioactivity amounts, were lower in G59A gradients. This was also visualized by EM analysis of the fractions obtained following sedimentation of G59A virus-infected cell lysates. Complete closed structures, as well as partial shells and deformed capsids, were observed (Fig. 5B). The different capsid structures observed in negatively stained material are shown in the four insets of Fig. 5B. The presence of these aberrant capsid structures explains the growth defect of the virus expressing VP5 G59A. Mutant G59V virus failed to replicate, and the defect in this virus was the inability of the mutant VP5 to bind with the scaffold protein since we observed numerous open shells in the nucleus by TEM (Fig. 5C and D). This was also confirmed by SDS-PAGE and EM analysis of sucrose gradient fractions of G59V virus-infected cell lysates, which revealed large open shells (data not shown). The virus encoding G59P did not accumulate ICP5 and thus was not analyzed further.

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FIG. 5. Mutations at conserved glycine 59. Ultrastructural analysis of cells infected with G59A mutant virus (A) revealed several enveloped particles, many that contained DNA (black arrowheads) and some that lacked DNA (white arrowheads). Naked nuclear capsids are indicated by the white arrow in panel A. Fractions derived from sucrose gradient analysis of G59A virus-infected cell lysates (B) revealed closed icosahedral capsids (white arrowheads) as well as aberrant shells (black arrowheads) that did not attain closure. Higher magnification of the shell structures is shown in the four insets of panel B. All images were obtained from material in fraction 8. Capsid shells (arrowheads) that have the characteristic 6 and 9 configuration were observed in the nuclei of cells infected with the G59V mutant virus (C and D) using TEM analysis. The nuclear envelope (ne), where visible, is indicated. Bars, 1 µm (A and B), 0.5 µm (C and D), and 100 nm (panel B insets).

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FIG. 6. Sedimentation analysis of the virus encoding an alanine substitution for the conserved glycine at position 59. Vero cells were infected and radiolabeled, and the lysates were sedimented through sucrose gradients as described in the legend to Fig. 4. The direction of sedimentation was from right to left (arrow). The positions where cosedimenting proteins of C, B, and A capsids were observed are shown at the bottom of the panel for G59A and K5R. The positions of the capsid proteins are indicated on the right. Molecular mass standards are shown in lane M and, where visible, are 220, 97.4, 66, 46, 30, and 14.3 kDa.

Lysine walk through the N terminus of VP5. The goal was to confirm previously identified hydrophobic binding domains and possibly uncover additional hydrophobic contacts between VP5 and the scaffold protein. Introduction of a lysine that does not directly target an interactive amino acid could add a hydrophilic side chain in a hydrophobic surface. Introduction of the lysine at 24, 29, 42, and 73 did not affect virus replication. This confirmed previous data from alanine scanning mutagenesis that showed these sites are not important (36); thus, no additional contact points exist in these regions. Lysine substitution at F36 (helix 1^22-42) and V68 (helix 2^58-72) abolished the interaction with the scaffold protein. This was visually seen by TEM analysis that revealed open shells in the nuclei of infected cells (Fig. 7A and B) and by sedimentation of infected-cell lysates followed by electron microscopy (Fig. 7A and B, insets) or SDS-PAGE analysis of the sucrose gradient fractions (Fig. 8, F36K). Only cosedimentation of the shell proteins was observed. The shells observed in V68K virus-infected cells and gradients were generally smaller and less abundant than those seen with the F36K virus. The lysine substitution at V80 gave an interesting result. The virus encoding this mutation failed to replicate on Vero cells (Table 3), even though it interacted with p22a in the yeast two-hybrid assay (Table 2). Sedimentation analysis revealed a peak of radioactivity in the region of the gradients at which B and A capsids sediment (Fig. 8, fractions 7 to 9 of V80K panel). However, the radioactivity corresponding to the scaffold protein was absent in the peak fractions (Fig. 8, fractions 7 to 9; compare to similar fractions of K5R in Fig. 6). TEM analysis of infected cells revealed many closed structures (Fig. 7C), although the shapes and internal compositions were unlike those of wild-type capsids (Fig. 3D). This was further illustrated when fractions from sucrose gradients were examined by EM (Fig. 7C, inset). The structures were aberrant in shape and staining pattern, even though there are some that are icosahedral in shape. The distribution of the aberrant capsid sizes was small enough so that the capsids sediment as a tight band in sucrose gradients (Fig. 8, fractions 7 to 9 in V80K). The V80K mutant particles in the nucleus in Fig. 8C were measured. The size range of the capsids was 71.95 nm to 109.65 nm (n = 22), with a mean of 94.05 nm. Wild-type capsids measured in a similar way ranged from 101.94 to 121.3 nm (n = 31), with a mean of 112.09 nm.

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FIG. 7. Electron micrographs of thin sections of infected cells and negatively stained structures of F36K, V68K, and V80K viruses. Cells were infected and processed for TEM or sucrose gradient fraction analysis as described in the legends to Fig. 3 and 4. Open shells were evident in F36K (A) and V68K (B) virus-infected cells (white arrowheads), whereas closed structures were evident in V80K virus (C)-infected cells (white arrowheads). The material in sucrose gradients of F36K (fraction 5), V68K, and V80K (fractions 7 to 9) virus-infected cell lysates was imaged following negative staining and is shown in the inset of each panel. The V68K virus structures observed were consistently smaller (inset to panel B). The nuclear envelope (ne) is indicated. Bars, 0.5 µm (A and C), 1 µm (B), and 100 nm (insets).

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FIG. 8. Polypeptide composition of F36K and V80K shell structures. Vero cell monolayers were infected, labeled, and processed for sucrose gradient analysis as described in the legend to Fig. 4. The direction of sedimentation, is indicated by the arrow. The positions of the capsid proteins are indicated on the right. Molecular mass standards are shown in lane M and, where visible, are 220, 97.4, 66, 46, 30, and 14.3 kDa.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
A mutant virus blocked at the maturation cleavage site in the scaffold protein (23) gave rise to second-site revertants that mapped to the gene encoding VP5 (10), the binding partner of the scaffold protein (p22a). Many of these second-site mutations mapped to the N terminus of this large polypeptide (37). Mutagenesis efforts were focused on a small specific region, the N-terminal 100 amino acids of this large polypeptide, to identity the SIDs in the major capsid protein. Two small scaffold protein binding domains were identified; the first comprises amino acids I27, L35, and F39, and the second includes amino acids L58, L65, L67, and L71 (36). We now designate these two SIDs SID1 and SID2, and they reside in two predicted alpha helices: helix 1^22-42 and helix 2^58-72 (1, 29).

The outcome of the current investigation has been a further elucidation and understanding of the hydrophobic nature and structural features of these interactive domains (Table 4). This has significant implication for developing an antiviral compound that targets an essential protein-protein interaction. It is clear both from the yeast two-hybrid assay and the replication properties of the mutant viruses that there is not a requirement for a specific hydrophobic side chain at the seven positions studied. Thus, leucine, isoleucine, methionine, and valine are interchangeable. This correlates with the bioinformatic analysis of these seven amino acids between the different herpesvirus families. In an alignment of these amino acids we saw at these positions substitutions of the hydrophobic group similar to those seen in our mutagenesis studies on HSV-1 VP5. At amino acid 39 (HSV numbering), for example, in all the members of the alpha- and gammaherpesvirus families there is a phenylalanine while in the betaherpesvirus family there is in most cases a leucine. At amino acid 67 there is a W for all the betaherpesviruses and in some gammaherpesviruses there is a Y. Overall, the conservation of the hydrophobic nature of these seven amino acids was evident. We changed the HSV binding domains to be more like those of human cytomegalovirus (HCMV) or Kaposi's sarcoma-associated herpesvirus (KSHV) and tested these constructs in the yeast two-hybrid assay for interaction with HSV-1 p22a. If the first domain was changed to that of HCMV (I27V, L35M, F39L), the VP5 protein expressed from this construct did bind to p22a but with slower kinetics than the wild type (data not shown). This was most likely due to the change to valine at 27 (Table 1). However, if the second domain was changed to that of HCMV (L58F, L67W), the altered protein reacted with p22a with wild-type kinetics (data not shown). The second domain was also altered to make it like KSHV (L65V, L67F); this protein failed to react with HSV-1 p22a (data not shown). The valine substitution in the SID regions affected the interaction between the two HSV-1 proteins in the yeast two-hybrid assay (I27, L35, L58, and L67), which was not so surprising considering that valine would add some rigidity in a region that may need to be flexible; however, there was minimal effect on virus replication in cell culture. Substitutions at F39 were made to determine the contribution of the aromatic side chain to this interaction. At this site the hydrophobic nature is still more important than the size of the side chain because replacement with leucine (nonaromatic) did not affect the interaction.

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TABLE 4. Summary of the phenotypes of the viruses encoding lethal VP5 mutations

The results of the replacement of the seven interactive amino acids with proline were most informative (Table 4). These seven amino acids are embedded in two alpha-helical sections of the protein. Proline residues are important for the conformation of the polypeptide backbone; they are generally well tolerated in turns and coils and at the beginning of alpha helices or beta sheets (25). The pyrrolidine ring of proline in a protein imposes significant constraints on the rotation of the polypeptide backbone at this site. Proline thus can affect protein stabilization (24), and insertion of this residue in an alpha helix has been shown to decrease protein stability (14). In the first domain, introduction of a proline abolished interaction with p22a but did not affect capsid shell formation. Proline substitutions in the second domain were deleterious both to protein accumulation and shell synthesis. Thus, the second domain is highly structured and very sensitive to perturbations of the helix. This correlates with previous studies that showed that the mutation one of the revertant viruses that mapped to amino acid R69 was temperature sensitive (37) and that alanine substitutions at F70 abolished the accumulation of ICP5 (36). In addition, an important observation that hinted at the role of this second domain was the observation that alanine substitution at L58 reduced capsomere-capsomere interactions, in that smaller open shells were assembled in L58A virus-infected cells (36). In this study the changes to proline in helix 2^58-72 significantly disrupted the structure of this domain and none of the viruses with proline substitutions were able to synthesize capsid shells (Table 4). Hence SID2 or helix 2^58-72 is important for shell accretion and stabilization by capsomere-capsomere interactions and/or capsomere-triplex interactions.

The lysine walk through this region was performed to uncover additional hydrophobic contacts. Of the seven lysines introduced into this region, two, at F36 and V68, abolished the interaction of VP5 with p22a both in vitro and as judged by virus replication. Both of these amino acids are close (F36 close to L35 and V68 close to L67) to the interactive residues previously identified. The introduction of the lysine in SID2 (V68K) not only abolished the interaction with p22a but also affected shell accretion; smaller open shells were consistently observed in V68K virus-infected cells. The phenotype produced by the V80K mutation was interesting. The mutation did not appear to affect the interaction with p22a in a bimolecular assay; however, in the virus the mutation impaired replication. We detected closed structures in V80K virus-infected cells, but these lacked the scaffold protein and were not the typical wild-type capsid structures. It appears that the change at that position may allow closure of the capsid shell (presumably via interactions with p22a), but as the structure nears closure the scaffold proteins may be eliminated by some unknown mechanism. In the absence of scaffold protein in these structures it was not possible to determine whether it (p22a) was cleaved at the maturation site prior to their loss.

Previously, we had not changed the conserved glycine at position 59. The flexibility of the glycine side chain and its location in SID2/helix 2^58-72 made it a candidate for investigation. Mutation to alanine does not affect its interaction with p22a, indicating that it does not directly participate in this process; however, virus replication was compromised by this change. The mutant virus was able to accumulate all intranuclear capsids, albeit at reduced levels, but there was still a significant population of capsid shells that were aberrant. A change to a valine that imparts a more rigid side chain abolished the VP5-p22a interaction; thus, flexibility in this region may be important for positioning the domains in the correct configuration for interaction. Since glycine is the simplest side chain (hydrogen), it imparts to the polypeptide chain much greater flexibility. Addition of progressively larger side chains at this site, alanine (CH₃) or valine (CH₃CH₂), similarly causes increased defects in capsid assembly: closed shells (G59A) and open shells (G59V). The proline substitution abolished accumulation of the polypeptide. Hence glycine 59, which resides in helix 2^58-72, although not directly participating in the interaction, may be important for the conformation or positioning of the SID. Since this amino acid is conserved in all three herpesvirus families, it may be a key residue for facilitating scaffold interaction in the herpesvirus family.

The structure of the HSV-1 capsid has been visualized by cryo-EM (39). These high-resolution analyses also predicted that the alpha-helical segments of the protein may also participate in capsomeric and intracapsomeric contacts (1). This appears to be validated by the mutagenesis studies that show that residues in helix 2^58-72 are involved in shell accretion. Helix 2^58-72 and the residues that reside in it may be important for accretion of the capsomeres to form large elaborate capsid shells for this large virus. Thus, we have identified a domain that is important for capsomere-capsomere interactions, either by VP5 interactions or by VP5-triplex interactions, that allow the formation of a large capsid shell for this virus.

The crystal structure of the major capsid protein has been obtained for a 65-kDa trypsin cleavage fragment (amino acids 451 to 1054) (4). This region of VP5 has been referred to as the upper domain. The floor domain, or the N terminus of VP5, is an important structural and functional motif. The structural fold in the floor domain is conserved in the Caudovirales bacteriophage (Hong Kong 97 and phage T4) family (2). It is believed that structural rearrangements of the floor domain that occur in phage during capsid maturation also occur in the HSV-1 VP5 protein (2, 16). Therefore, these viruses have conserved a protein fold that is an important regulator of capsid shell assembly and stabilization. Elucidation of the structure of the HSV-1 floor domain is important not only for developing a new antiviral strategy but also in the context of evolution of animal viruses and their structure.

 
This work was supported by National Institutes of Health grant AI 033077.

We thank Gerry Sexton for his help with some of the initial TEM analysis and Marieta Sole for help with the sucrose gradient analysis shown in Fig. 4. We also thank Stanley Person for his support, discussions of the data and critical review of the manuscript, and infectious enthusiasm for this research.

 
* Corresponding author. Mailing address: Viral Oncology Program, The Sidney Kimmel Comprehensive Cancer Center, Rm. 353 CRB 1, 1650 Orleans Street, The Johns Hopkins University, Baltimore, MD 21231. Phone: (410) 614-1581. Fax: (410) 955-0840. E-mail: pdesai{at}jhmi.edu

Published ahead of print on 20 June 2007.

Present address: New York University, College of Dentistry, 345 East 24^th Street, New York, NY 10010.

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Journal of Virology, September 2007, p. 9396-9407, Vol. 81, No. 17
0022-538X/07/$08.00+0     doi:10.1128/JVI.00986-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

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