ABSTRACT
VP26 is a herpes simplex virus 1 (HSV-1) small capsomere-interacting protein. In this study, we investigated the function of VP26 in HSV-1-infected cells with the following results. (i) The VP26 null mutation significantly impaired incorporation of minor capsid protein UL25 into nucleocapsids (type C capsids) in the nucleus. (ii) The VP26 mutation caused improper localization of UL25 in discrete punctate domains containing multiple capsid proteins (e.g., the VP5 major capsid protein) in the nucleus; these domains corresponded to capsid aggregates. (iii) The VP26 mutation significantly impaired packaging of replicated viral DNA genomes into capsids but had no effect on viral DNA concatemer cleavage. (iv) The VP26 mutation reduced the frequency of type C capsids, which contain viral DNA but not scaffolding proteins, and produced an accumulation of type A capsids, which lack both viral DNA and scaffold proteins, and had no effect on accumulation of type B capsids, which lack viral DNA but retain cleaved scaffold proteins. Collectively, these results indicated that VP26 was required for efficient viral DNA packaging and proper localization of nuclear capsids. The phenotype of the VP26 null mutation was similar to that reported previously of the UL25 null mutation and of UL25 mutations that preclude UL25 binding to capsids. Thus, VP26 appeared to regulate nucleocapsid maturation by promoting incorporation of UL25 into capsids, which is likely to be required for proper capsid nuclear localization.
IMPORTANCE HSV-1 VP26 has been reported to be important for viral replication and virulence in cell cultures and/or mouse models. However, little is known about the function of VP26 during HSV-1 replication, in particular, in viral nucleocapsid maturation although HSV-1 nucleocapsids are estimated to contain 900 copies of VP26. In this study, we present data suggesting that VP26 promoted packaging of HSV-1 DNA genomes into capsids by regulating incorporation of capsid protein UL25 into capsids, which was reported to increase stability of the capsid structure. We also showed that VP26 was required for proper localization of capsids in the infected cell nucleus. This is the first report showing that HSV-1 VP26 is a regulator for nucleocapsid maturation.
INTRODUCTION
Herpesviruses are classified into three subfamilies: Alphaherpesvirinae, Betaherpesvirinae, and Gammaherpesvirinae (1). Herpes simplex virus 1 (HSV-1), the subject of this study, is a member of the Alphaherpesvirinae subfamily and is one of the best-studied herpesviruses, causing a variety of human diseases, e.g., mucocutaneous diseases, keratitis, skin diseases, and encephalitis (2).
The genomes of viruses in the Herpesviridae family (herpesviruses) are encased and protected by icosahedral capsids (1). These capsids are formed by 161 capsomeres (150 hexons and 11 pentons), a portal complex that has an axial channel through which viral genome DNA enters and exits capsids, 320 triplexes that connect the capsomeres and the portal complex, small capsomere-interacting proteins (SCPs), and capsid vertex-specific complexes (CVSCs) that are rod-shaped with five rods located near each capsid vertex (3–5). In HSV-1 capsids, both pentons and hexons are composed of 5 and 6 VP5 molecules, respectively; the CVSCs are composed of 1 molecule of UL17 and 1 molecule of UL25, the triplexes are composed of 1 molecule of VP19C and 2 molecules of VP23, the portal complex is composed of 12 molecules of UL6, and HSV-1 VP26 SCPs form a hexameric ring on the outer surface of each hexon (3–5).
Herpesvirus capsid formation takes place in the infected cell nucleus (3–5). In HSV-1-infected cells, complexes of VP5 and scaffolding proteins UL26.5 and UL26, in which UL26 is less abundant than UL26.5, associate with each other to form a spherical intermediate capsid, designated the procapsid, with binding promoted by scaffold protein-scaffold protein interactions and by the triplexes that link VP5 molecules (3–5). UL26 is the VP24 maturation protease fused to the N terminus of UL26.5 and is located on the inside of the scaffold shell (3–5). After the procapsid is formed, UL26 proteolytic activity is activated, and the scaffolding proteins detach from the capsid shell, a process mediated by proteolytic cleavage of UL26 and UL26.5 near their C-terminal ends. The viral DNA genome is then packaged, with DNA genome transport into the capsid mediated by the viral terminase, a three-component ATPase complex composed of UL15, UL28, and UL33 (3–5). The HSV-1 terminase cleaves nascent viral concatemeric DNA into unit-length viral genomes, docks at the capsid portal vertex, and packages a cleaved progeny virus genome into the capsid (3–5). In addition, the UL25 and UL17 components of CVSCs have been reported to be required for cleavage and/or packaging of nascent HSV-1 DNA genomes (6, 7).
In HSV-1-infected cells, three types of capsids (A, B, and C capsids) have been detected (3–5). Type A and B capsids are incomplete structures resulting from problems in viral genome packaging: type B capsids do not contain viral genome DNA but do contain cleaved scaffold proteins, and type A capsids do not contain either viral genome DNA or scaffold proteins (3–5). The type C capsid is a mature capsid and contains a viral genome but no longer contains scaffold proteins (3–5). The C capsid also has CVSCs on its surface (8). Although it has been reported that components of the CVSCs could be detected on type A and B capsids, the amount was lower than that found on type C capsids (8–13).
The HSV-1 VP26 SCP is conserved in herpesviruses (1). In vitro reconstitution analyses of herpesvirus capsids showed that VP26 was not essential for assembly of HSV-1 capsids in vitro (14, 15), unlike its homologs in Epstein-Barr virus (EBV) and Kaposi's sarcoma-associated herpesvirus (KSHV) (16, 17). Although there has been a lack of information on the involvement of HSV-1 VP26 in capsid assembly and maturation in infected cells, the report that VP26 was not essential for assembly of HSV-1 capsids in vitro is in agreement with the observation that VP26 was not essential for viral replication in cell cultures (18). But HSV-1 VP26 has been reported to be required for efficient viral replication and virulence in vivo and in cell cultures (18–20). However, the mechanism(s) by which VP26 acts in HSV-1 replication and virulence remains largely unknown. We recently reported that, in cells infected with HSV-1 with the VP26 null mutation, VP5 was improperly localized in discrete punctate structures in the nucleus (19). This improper localization of VP5 was also observed in cells infected with HSV-1 carrying null mutations in UL25 and UL17 (21, 22), both of which were shown to be required for cleavage and/or packaging of nascent HSV-1 genomes into capsids (6, 7), as described above. These results suggested that HSV-1 VP26 acted in the same pathway as UL17 and UL25 and led us to hypothesize that HSV-1 VP26, like these HSV-1 capsid proteins, was also a regulator of nucleocapsid maturation. We tested this hypothesis in this study.
RESULTS
Characterization of the recombinant viruses generated in this study.To investigate the effects of VP26 in HSV-1-infected cells, especially effects on UL17 and UL25, we constructed and characterized the following recombinant viruses: YK495 (ΔP26-repair) in which the VP26 null mutation in YK490 (ΔVP26) (19) was repaired; YK497 (UL17-Myc/Flag-UL25) expressing UL17-Myc and Flag-UL25; YK498 (UL17-Myc/Flag-UL25/ΔVP26) expressing UL17-Myc and Flag-UL25 and carrying a null mutation in VP26; and YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) in which the VP26 null mutation in YK498 (UL17-Myc/Flag-UL25/ΔVP26) was repaired (Fig. 1). As described above, the antibodies against UL17 and UL25 generated in this study could not be used in immunofluorescence studies. Therefore, we generated recombinant viruses expressing UL17 and UL25 that were fused to different tags to enable the spatial analyses of these capsid proteins in HSV-1-infected cells.
Schematic diagrams of the genome structures of wild-type HSV-1(F) and the relevant domains of the recombinant viruses used in this study. Line 1, wild-type HSV-1(F) genome; lines 2 and 6, domain of the UL34 gene to UL36 gene; lines 3, 4, 7, and 8, recombinant viruses with mutations in the UL35 (VP26) gene; line 5, recombinant virus encoding Myc-tagged UL17 and Flag-tagged UL25. UL35 and UL36 encode VP26 and VP1/2, respectively.
As expected, Vero cells infected with YK490 (ΔVP26) or YK498 (UL17-Myc/Flag-UL25/ΔVP26) did not express VP26, but cells infected with wild-type HSV-1(F), YK495 (ΔVP26-repair), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) did (Fig. 2A and B). Vero cells infected with YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) expressed the tagged UL17 and UL25 proteins, but cells infected with wild-type HSV-1(F) did not (Fig. 2B). Vero cells infected with YK490 (ΔVP26) or YK498 (UL17-Myc/Flag-UL25/ΔVP26) at a multiplicity of infection (MOI) of 5 for 18 h accumulated capsid proteins, including VP23, UL17, UL25, UL17-Myc, and Flag-UL25, at levels similar to those in cells infected with wild-type HSV-1(F), YK495 (ΔP26-repair), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) (Fig. 2A and B). These results indicated that the VP26 null mutation had no effect on accumulation of the other capsid proteins in HSV-1-infected cells.
Characterization of the recombinant viruses generated in this study. (A and B) Vero cells were mock infected or infected with wild-type HSV-1(F), YK490 (ΔVP26), YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 5 for 18 h and then analyzed by immunoblotting with the indicated antibodies. (C) Vero cells were mock infected or infected with wild-type HSV-1(F) or YK497 (UL17-Myc/Flag-UL25) at an MOI of 5 for 18 h and then analyzed by immunoblotting with the indicated antibodies. (D) Vero cells were infected with wild-type HSV-1(F) or YK497 (UL17-Myc/Flag-UL25) at an MOI of 0.01. At 48 h postinfection, the cell culture supernatants were harvested, and virions were purified from the supernatants and analyzed by immunoblotting with the indicated antibodies. (E to H) Vero cells were infected with wild-type HSV-1(F), YK490 (ΔVP26), YK495 (ΔVP26-repair), YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), and YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair), as indicated, at an at an MOI of 5 or 0.01. Total virus from the cell culture supernatants and infected cells was harvested at the indicated times and assayed on Vero cells. Each data point is the mean ± standard error of the results of three independent experiments.
Vero cells infected with YK497 (UL17-Myc/Flag-UL25) at an MOI of 5 for 18 h accumulated VP5, VP26, UL17, and UL25 at levels similar to those of cells infected with wild-type HSV-1(F) (Fig. 2C). UL17, UL25, and VP26 were incorporated into virions purified from the supernatants of Vero cells infected with YK497 (UL17-Myc/Flag-UL25) at levels similar to those of virions purified from supernatants of wild-type HSV-1(F)-infected cells (Fig. 2D). Viral production of YK490 (ΔVP26) and YK498 (UL17-Myc/Flag-UL25/ΔVP26) in Vero cells infected at an MOI of 5 or 0.01 was less than that of wild-type HSV-1(F), YK495 (ΔP26-repair), and YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) (Fig. 2E to H). The decrease in progeny virus production by cells infected with the VP26 null mutant viruses [YK490 (ΔVP26) and YK498 (UL17-Myc/Flag-UL25/ΔVP26)] in this study was in agreement with previous reports of other VP26 deletion mutant viruses (18–20). YK497 (UL17-Myc/Flag-UL25) and YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) grew in Vero cells almost as efficiently as wild-type HSV-1(F) (Fig. 2G and H). These results indicated that the tags on UL17 and UL25 had no apparent effect on (i) accumulation of capsid proteins (i.e., UL17, UL25, VP5, VP23, and VP26) in HSV-1-infected cells, (ii) incorporation of UL17 and UL25 into extracellular progeny virions, and (iii) viral replication in cell cultures.
Effect of VP26 on incorporation of UL17 and UL25 into intranuclear capsids and extracellular virions.As the first step to investigate whether VP26 is involved in nucleocapsid maturation, we examined the effect of the VP26 null mutation on incorporation of UL17 and UL25 into capsids. Intranuclear capsids were isolated from lysates of the nuclear fractions of Vero cells infected with YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 3 for 18 h and then purified on sucrose gradients. Capsids were detected as visible light-scattering bands containing type A, B, or C capsids (Fig. 3A). The gradients were fractionated, and the proteins in each fraction were analyzed by denaturing gel electrophoresis followed by silver staining. Fractions 3, 5, and 11 contained type A, B, and C capsids, respectively (Fig. 3B). These fractions were also analyzed by immunoblotting (Fig. 3C), and the amount of protein in the VP5, UL17-Myc, and Flag-UL25 immunoblot bands was quantitated. The incorporation of UL17-Myc and Flag-UL25 into each type of capsid was calculated relative to the amount of protein in VP5 and normalized to the relative value for wild-type virus. As shown in Fig. 3D and E, the amount of UL17-Myc and Flag-UL25 incorporated in type A, B, and C capsids from cells infected with YK498 (UL17-Myc/Flag-UL25/ΔVP26) was lower than the amount in capsids from cells infected with YK497 (UL17-Myc/Flag-UL25) (Fig. 3D and E, wild type) or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair). Thus, the VP26 null mutation significantly impaired the incorporation of UL17-Myc (Fig. 3D) and Flag-UL25 (Fig. 3E) into type C capsids. In contrast, the VP26 null mutation appeared to produce a small decrease in the incorporation of UL17-Myc (Fig. 3D) and Flag-UL25 (Fig. 3E) into type A and B capsids, but these were not statistically significant decreases (Fig. 3D and E).
Effect of the VP26 null mutation on the incorporation of UL17 and UL25 into intranuclear capsids. (A) Vero cells were infected with YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 3 and harvested at 18 h postinfection. Nuclear lysates were isolated and layered onto 20 to 50% sucrose gradients and ultracentrifuged at 110,000 × g for 1 h. The positions of type A, B, and C capsid bands are indicated. (B) Proteins in the gradient fractions shown in panel A were analyzed by denaturing gel electrophoresis and silver stained. The positions of capsid proteins VP5, VP19C, VP21, VP22a, VP23, and VP26 are indicated. (C) The gradient fractions in shown in panel A containing type A, B, or C capsids were analyzed by immunoblotting with the indicated antibodies. These data are representative of five independent experiments. (D and E) Quantitation of the amount of UL17-Myc and Flag-UL25, as indicated, in the immunoblot bands shown in panel C relative to the amount in the VP5 band. Each value is the mean ± standard error of five independent experiments and was normalized relative to the mean value of YK497 (UL17-Myc/Flag-UL25) capsids, which are designated wild type in panels D and E. *, P < 0.05 (by Tukey's test); n.s., not significant.
We also examined the effect of the VP26 null mutation on incorporation of UL17 and UL25 into mature virions by analyzing virions purified from the supernatants of Vero cells infected with YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 0.01 for 48 h by immunoblotting (Fig. 4). In agreement with the results for type C capsids purified from the nuclei of infected cells (Fig. 3), the levels of incorporation of UL17-Myc and Flag-UL25 into YK498 (UL17-Myc/Flag-UL25/ΔVP26) virions were significantly lower than the amounts incorporated into YK497 (UL17-Myc/Flag-UL25) and YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) virions (Fig. 4). Together, these results suggested that VP26 was required for efficient incorporation of UL17 and UL25 into HSV-1 nucleocapsids.
Effect of the VP26 null mutation on incorporation of UL17 and UL25 into extracellular virions. (A) Vero cells were infected with YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 0.01. Cell culture supernatants were then harvested at 48 h postinfection, and virions were purified and analyzed by immunoblotting with the indicated antibodies. Data are representative of five independent experiments. (B and C) Quantitation of the amounts of UL17-Myc and Flag-UL25, as indicated, in the immunoblot bands shown in panel A relative to the amount of VP5. Each value is the mean ± standard error of five independent experiments and was normalized relative to the mean value of YK497 (UL17-Myc/Flag-UL25) capsids. *, P < 0.05 (by Tukey's test); n.s., not significant.
Effect of VP26 on subcellular localization of capsid proteins in HSV-1-infected cells.As described above, the absence of VP26 led to improper localization of VP5 in discrete punctate structures in the nucleus of HSV-1-infected cells (19). To investigate whether VP26 affects subcellular localization of UL17 and UL25 in HSV-1-infected cells, as it does VP5 (19), Vero cells mock infected or infected with wild-type HSV-1(F), YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 5 were fixed at 15 h postinfection and analyzed by confocal microscopy. In mock-infected cells or cells infected with wild-type HSV-1(F), anti-Myc and anti-Flag antibodies did not detect any specific fluorescence, but fluorescence was detected in cells infected with YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) (Fig. 5). The localization pattern of VP5 in cells infected with YK497 (UL17-Myc/Flag-UL25) or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) was similar to that in cells infected with wild-type HSV-1(F) (Fig. 5). In addition, localization of UL17-Myc and Flag-UL25 detected by anti-Myc and anti-Flag antibodies, respectively, and the relationship of these tagged capsid proteins with VP5 (Fig. 5) were in agreement with these aspects of the endogenous viral proteins reported in wild-type HSV-1-infected cells (22). Thus, UL17-Myc and Flag-UL25 were distributed throughout the nucleus of cells infected with YK497 (UL17-Myc/Flag-UL25) as numerous punctate foci and partially colocalized with VP5 in these nuclear punctate structures (Fig. 5). These results strongly suggested that the fluorescence detected with anti-Myc and anti-Flag antibodies by confocal microscopy of cells infected with the recombinant viruses expressing tagged UL17 and UL25 was due to specific localization of Myc-UL17 and Flag-UL25, respectively, and that tagging these capsid proteins had no apparent effect on wild-type localization of UL17, UL25, and VP5 in HSV-1-infected cells.
Effect of the VP26 null mutation on subcellular localization of UL17, UL25, and VP5 in HSV-1-infected cells. Vero cells were mock infected or infected with wild-type HSV-1(F), YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 5, fixed at 15 h postinfection, permeabilized, stained with antibodies to VP5 (green) and Myc (red) (A) or antibodies to VP5 (green) and Flag (red) (B), and examined by confocal microscopy. The panels in the fourth and seventh rows are magnified images of the boxed areas in the panels in the third and sixth rows, respectively.
As described above, in cells infected with YK497 (UL17-Myc/Flag-UL25) or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair), VP5, UL17-Myc, and Flag-UL25 were dispersed throughout the nucleus as numerous punctate dots, whereas in cells infected with YK498 (UL17-Myc/Flag-UL25/ΔVP26), UL17-Myc and Flag-U25 were detected in large discrete punctate structures in the nucleus and colocalized strongly with VP5 in these nuclear structures (Fig. 5). These results indicated that VP26 was required for proper localization of UL17 and UL25 in HSV-1-infected cells and, as previously reported (19), that VP26 regulated proper localization of VP5 in HSV-1-infected cells.
Effect of VP26 on subcellular localization of ICP8 and UL25 in HSV-1-infected cells.It has been proposed that HSV-1 capsid proteins are recruited to and assembled in nuclear domains where viral genome replication takes place, designated replication compartments (23, 24). The observation reported above that, in the absence of VP26, capsid proteins VP5, UL17, and UL25 were improperly localized in aberrant discrete domains in the nucleus led us to investigate the relationship between the sites for accumulation of capsid proteins and the replication compartments in the presence or absence VP26. Therefore, Vero cells were mock infected or infected with wild-type HSV-1(F), YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 5, fixed at 15 h postinfection, and examined by confocal microscopy. As shown in Fig. 6, the localization pattern of ICP8 in cells infected with YK497 (UL17-Myc/Flag-UL25) or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) was similar to that in wild-type HSV-1(F)-infected cells, strongly suggesting that tagging UL17 and UL25 had no apparent effect on ICP8 localization. In agreement with previous reports (22), Flag-UL25 and ICP8 were distributed throughout the nucleus as numerous punctate foci and were partially colocalized in these punctate structures in cells infected with YK497 (UL17-Myc/Flag-UL25) or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) (Fig. 6). In the nuclei of cells infected with YK498 (UL17-Myc/Flag-UL25/ΔVP26), UL25 was improperly localized in large discrete punctate domains, as shown in Fig. 5, but ICP8 localization in these cells was similar to that in cells infected with YK497 (UL17-Myc/Flag-UL25) or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) (Fig. 6). Three-dimensional reconstitution images of ICP8 and Flag-UL25 localization in cells infected with YK498 (UL17-Myc/Flag-UL25/ΔVP26) confirmed that the discrete punctate structures of Flag-UL25 were apparently dissociated from the ICP8 foci, indicating that ICP8 was not colocalized with Flag-UL25 in cells YK498 infected with (UL17-Myc/Flag-UL25/ΔVP26) (see Movie S1 in the supplemental material). Taken together with the result that VP5 strongly colocalized with UL17-Myc and Flag-UL25 in discrete nuclear domains in the absence of VP26 in HSV-1-infected cells (Fig. 5), these results suggested that VP26 was required for proper recruitment of multiple capsid proteins, including VP5, UL17, and UL25, to the replication compartments.
Effect of the VP26 null mutation on subcellular localization of ICP8 and UL25 in HSV-1-infected cells. Vero cells were mock infected or infected with wild-type HSV-1(F), YK497 (UL17-Myc/Flag-UL25), YK498 (UL17-Myc/Flag-UL25/ΔVP26), or YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair) at an MOI of 5, fixed at 15 h postinfection, permeabilized, stained with antibodies to Flag (green) and ICP8 (red), and examined by confocal microscopy. Boxed areas are magnified in the images immediately below them.
Effect of VP26 on localization intranuclear capsids in HSV-1-infected cells.The observations noted above (Fig. 5 and 6) that, in the absence of VP26, multiple capsid proteins were improperly localized in the discrete nuclear domains in HSV-1-infected cells led us to examine whether VP26 affected localization of nuclear capsids in HSV-1-infected cells. Therefore, Vero cells were infected with wild-type HSV-1(F), YK490 (ΔVP26), or YK495 (ΔVP26-repair) at an MOI of 5 for 18 h and then analyzed by electron microscopy. As shown in Fig. 7, capsids were dispersed throughout the nucleus in cells infected with wild-type HSV-1(F) or YK495 (ΔVP26-repair). In contrast, capsids were aggregated in discrete nuclear domains in cells infected with YK490 (ΔVP26) (Fig. 7), which was in agreement with the fluorescence analyses of capsid proteins in these infected cells (Fig. 5 and 6). These results indicated that VP26 was required for proper localization of nuclear capsids in HSV-1-infected cells. Correlative light and electron microscopy (CLEM) analyses of Vero cells infected with YK490 (ΔVP26) showed that the discrete nuclear domains with VP5 detected by confocal microscopy appeared to correspond to the capsid aggregates detected in the nucleus of these infected cells by electron microscopy (Fig. 8). We should note that most HSV-1 capsids detected in our CLEM analyses appeared to be empty (Fig. 8) since the cores of the HSV-1 capsids were difficult to visualize due to inefficient fixation of the samples in these analyses.
Ultrastructural analysis of the effects of the VP26 null mutation on morphogenesis of intranuclear capsids. Vero cells were infected with wild-type HSV-1(F), YK490 (ΔVP26), or YK495 (ΔVP26-repair). At 18 h postinfection, the cells were embedded, sectioned, stained, and examined by transmission electron microscopy. Each image in the right column is the magnified image of the boxed area in the image to its left. Arrows indicate capsid aggregates. White, yellow, and black arrowheads indicate type A, B, and C capsids, respectively. Nuc, nucleus; Cy, cytoplasm. Scale bars, 1 μm (left panels) and 500 nm (right panels).
CLEM analysis of YK490 (ΔVP26)-infected Vero cells. (A) Vero cells were infected with YK490 (ΔVP26) at an MOI of 5, fixed at 18 h postinfection, permeabilized, stained with antibody to VP5 (green), and examined by confocal microscopy. (B) The cell analyzed in the experiment shown in panel A was embedded, sectioned, stained, and examined by transmission electron microscopy. (C to E) Magnified images of the indicated boxed areas in panel B. Nuc, nucleus; Cy, cytoplasm. Scale bars, 2 μm (A and B) and 500 nm (C to E).
Effect of VP26 on cleavage of HSV-1 DNA concatemers and DNA packaging into capsids in HSV-1-infected cells.We next investigated whether VP26, like UL25 and UL17, was involved in regulation of cleavage of HSV-1 DNA concatemers and/or DNA packaging into capsids. Total DNA and DNase I-resistant DNA, which was considered to be packaged viral DNA, were purified from the nuclear fraction of Vero cells infected with wild-type HSV-1(F), YK490 (ΔVP26), or YK495 (ΔVP26-repair) at an MOI of 3 for 18 h and was then digested with BamHI, analyzed by Southern blotting, and probed with the BamHI S-P fragment. BamHI cleaved HSV-1 viral DNA to produce the terminal BamHI S and P fragments, which are the termini of the unique long (UL) and unique short (US) components of the HSV-1 genome, respectively, and the junction-spanning BamHI S-P fragment (25) (Fig. 9A). In contrast, BamHI digestion of concatemeric viral DNA gave rise to only the BamHI S-P fragment (25) (Fig. 9A). The amount of total DNA in the BamHI S, P, and S-P fragments in the Southern blot bands was quantitated, and the amount of DNA in the S and P fragments was calculated relative to the amount of DNA in the BamHI S-P fragment. As shown in Fig. 9B and C, the amounts of BamHI S and P fragments relative to that of the BamHI S-P fragment in cells infected with YK490 (ΔVP26) were similar to the amounts in cells infected with wild-type HSV-1(F) or YK495 (ΔVP26-repair), indicating that the VP26 null mutation had no apparent effect on HSV-1 DNA concatemer cleavage in infected cells.
Effect of the VP26 null mutation on HSV-1 DNA genome cleavage and packaging. (A) Schematic diagram of the HSV-1 genome showing the positions of the BamHI S, P, and S-P DNA fragments. (B to E) Vero cells infected with wild-type HSV-1(F), YK490 (ΔVP26), or YK495 (ΔVP26-repair) at an MOI of 5 were harvested at 18 h postinfection. Nuclear fractions of infected cells were lysed and incubated in the presence or absence of DNase I, and the DNAs were then purified. Total DNA and DNase I-resistant DNA (i.e., packaged viral DNA) were digested with BamHI and analyzed by Southern blotting with the BamHI S-P fragment as a probe (B). The amounts of DNA in BamHI S and P fragments in total DNA were calculated relative to the amount of DNA in the BamHI S-P fragment (C and D). The amount of DNA in the BamHI S-P fragment in DNase I-resistant DNA was calculated relative to the amount of DNA in the BamHI S-P fragment in total DNA (E). Each value is the mean ± SEM of three independent experiments. *, P < 0.05 (by Tukey's test); n.s., not significant.
To investigate the effect of the VP26 null mutation on HSV-1 DNA packaging, the amount of BamHI S-P fragment from DNase I-resistant DNA was determined relative to the amount of BamHI S-P fragment without DNase I treatment. As shown in Fig. 9D, the amount of DNase I-resistant BamHI S-P DNA relative to the amount of total BamHI S-P DNA in cells infected with YK490 (ΔVP26) was significantly lower than that in cells infected with wild-type HSV-1(F) or YK495 (ΔVP26-repair), indicating that the VP26 null mutation significantly impaired HSV-1 DNA genome packaging into capsids in HSV-1-infected cells. Collectively, these results indicated that VP26 was required for efficient HSV-1 DNA genome packaging but not for its cleavage from concatemers in HSV-1-infected cells.
Effect of VP26 on the distribution of types of intranuclear capsids in HSV-1-infected cells.In the electron micrographs shown in Fig. 7, we noted that the frequency of type A and C capsids in the nuclei of cells infected with YK490 (ΔVP26) appeared to be different than that in cells infected with wild-type HSV-1(F) or YK495 (ΔVP26-repair). Therefore, we investigated the effect of VP26 on the morphogenesis of HSV-1 nuclear capsids by quantitating the number of each type of nuclear capsid in the electron micrographs shown in Fig. 7. As shown in Table 1, the fraction of capsids that were type A was higher in cells infected by YK490 (ΔVP26) than in cells infected by wild-type HSV-1(F); this difference was statistically significant. The wild-type level of type A capsids was restored in cells infected by YK495 (ΔVP26-repair) (Table 1). The fraction of capsids that were type C was lower in cells infected by YK490 (ΔVP26) than in cells infected by wild-type HSV-1(F); this difference was statistically significant (Table 1). The wild-type level of type C capsids was restored in cells infected by YK495 (ΔVP26-repair) (Table 1). The fraction of capsids that were type B was the same, at a statistically significant level, in cells infected with wild-type HSV-1(F), YK490 (ΔVP26), or YK495 (ΔVP26-repair) (Table 1). Therefore, infection with HSV-1 carrying the VP26 null mutation led to an increase in the fraction of type A capsids, no change in the fraction of type B capsids, and a decrease in the fraction of type C capsids. These results indicated that VP26 was required for efficient production of HSV-1 nucleocapsids.
Effect of the VP26 null mutation on the types of nuclear capsids in HSV-1-infected Vero cells
DISCUSSION
Homologs of HSV-1 SCP VP26 in alphaherpesviruses have often been utilized as targets for tagging by fluorescent proteins to visualize viral capsids by image analysis since they are dispensable for capsid assembly (26–31). Tagging capsid proteins that are essential for capsid assembly with fluorescent proteins generally prevents capsid assembly, and, therefore, recombinant viruses expressing these capsid proteins tagged with fluorescent proteins cannot be used to study capsid assembly. On the other hand, HSV-1 VP26 and its alphaherpesvirus homologs have been reported to be important for viral virulence in mice and for viral replication in cell cultures, trigeminal ganglia, and human skin xenografts in mice (18–20, 32, 33). However, there is a lack of information on the molecular mechanism(s) by which VP26 and its homologs act in viral replication and pathogenicity, especially in viral nucleocapsid maturation, even though alphaherpesvirus SCPs are the second most abundant protein in viral capsid structures (3, 4). In this study, we present data demonstrating that HSV-1 VP26 is a novel regulator for nucleocapsid maturation. To our knowledge, this is the first report clarifying a role for an alphaherpesvirus SCP in nucleocapsid maturation in infected cells.
In this study, we showed that VP26 was required for efficient incorporation of UL25 into HSV-1 nucleocapsids and virions. The phenotype of the VP26 null mutation in nucleocapsid maturation observed in this study was similar to that of the UL25 null mutation, as previously reported (7, 34). The UL25 null mutation (i) had no obvious effect on cleavage of replicated HSV-1 DNA concatemers, (ii) impaired packaging of viral DNA genomes, and (iii) accumulated type A capsids and reduced type C capsid formation in the nuclei of infected cells (7, 34). Interestingly, of the HSV-1 regulators for viral DNA genome packaging, this phenotype is unique to UL25. Mutations in other HSV-1 regulators (including UL6, UL15, UL17, UL28, UL32, and UL33) were reported to completely eliminate viral DNA concatemer cleavage and genome packaging so that the DNA concatemers were not cleaved, and only type B capsids were formed (6, 35–40). Also, mutations in UL25 that preclude its binding to capsids were shown to exhibit the same phenotype as the UL25 null mutation (9), suggesting that UL25 functions in HSV-1 DNA genome packaging in capsids. Collectively, these observations suggested that VP26 mediated recruitment of UL25 to the capsid surface, thereby promoting packaging of viral DNA genomes into capsids.
At present, it is not known how VP26 recruits UL25 to the capsid surface. It has been reported that VP26 interacted with UL25 in yeast two-hybrid assays and that VP26 tagged with a fluorescent protein coprecipitated with UL25 from lysates of HSV-1-infected cells (41, 42). Furthermore, reconstitution of cryo-electron microscopy images of HSV-1 capsids showed that VP26 was located close to UL25 on the capsid surface (8). Therefore, VP26 may recruit UL25 to the capsid surface and/or bind UL25 to aid in its proper positioning on the capsid surface. It has also been reported that UL17 was required for efficient incorporation of UL25 into capsids (43). In addition, UL17 was shown to interact with VP26 (42). These results suggest another possibility: VP26 may interact with UL17 and aid the incorporation of UL17 into capsids, thereby indirectly promoting the recruitment of UL25 on the capsid surface. Alternatively, VP26 may contribute indirectly to recruiting UL25 to and/or retaining UL25 on the capsid surface by regulating the conformation of the capsid. It has been reported (i) that CVSC components (i.e., UL25 and UL17) were incorporated into type A, B, and C capsids, with the greatest amount in type C capsids (8–13, 43), (ii) that cryo-electron microscopic analyses showed that CVSC structures were easily visible in type C capsids but not in type A and B capsids (8), (iii) that the presence of CVSCs on type C capsids appeared to depend on the capsids containing viral genome DNA since removal of viral genome DNA from type C capsids appeared to release UL25 and UL17 from the capsids (8), and (iv) that internal pressure from encapsidated viral DNA in type C capsids is considerably high (∼18 atmospheres) (44). Collectively, these observations suggested that the high pressure from encapsidated viral DNA may induce conformational changes of the capsid surface that affect CVSC binding sites, resulting in enhanced CVSC affinity with the capsid. In support of this hypothesis, internal pressure from encapsidated viral DNA was recently reported to directly influence the stability of viral particles (45). Thus, proper retention of intact CVSCs on the capsid surface appeared to be dependent on the conformation of the capsid surface. Since 900 copies of VP26 are estimated to be located on the capsid surface (3, 46), VP26 alone may regulate the optimal conformation of the capsid surface for efficient recruitment of CVSCs. Such a VP26 effect and a conformational change of the capsid surface mediated by the pressure of its encapsidated DNA genome on CVSC recruitment would likely be synergistic, based on the observations in this study that the VP26 null mutation reduced the incorporation of UL25 and UL17 into type C capsids more than into type A and B capsids.
It has been suggested that auxiliary capsid proteins of double-stranded DNA (dsDNA) phages and HSV-1 UL25 reinforce capsids to retain the pressurized viral DNA in capsids (34, 47). Recent atomic force microscopy nanoindentation analysis showed that capsids of HSV-1 and bacteriophage λ become stiffer upon binding auxiliary capsid proteins UL25 and gpD, respectively, due to increased structural stability (48). Therefore, the absence of UL25 on HSV-1 capsids and the decrease in UL25 incorporation into capsids mediated by the VP26 null mutation may lead to an imbalance between capsid structural stability and the high pressure due to encapsidated viral DNA. This might result in the loss of viral DNA from capsids, thereby leading to an accumulation of type A capsids in cells infected with a UL25 null or VP26 null mutant virus (34, 47). Structural instability of capsids due to absent or reduced UL25 on capsids also appeared to change the property(ies) of capsids. In support of this suggestion, the absence or reduction in incorporation of UL25 into capsids, which can be caused by the UL25, UL17, and VP26 null mutations, produced improper localization of multiple capsid proteins (e.g., UL17, UL25, VP5, and VP22a) in discrete nuclear domains and appeared to induce aggregation of nuclear capsids (6, 19, 21, 22, 43). Previous studies were not able to determine whether the discrete nuclear domains with accumulations of multiple capsid proteins detected by immunofluorescence represent capsid aggregates or are aberrant accumulations of capsid proteins that have not formed capsid structures (6, 19, 21, 22, 43). In this study, CLEM analysis indicated that the accumulations of capsid proteins were probably capsid aggregates. Thus, the absence of or reduction in UL25 incorporation into capsids may induce a change in capsid structural stability, causing the capsids to be “sticky,” thereby resulting in formation of capsid aggregates in the nucleus. We noted that mature nucleocapsids (type C capsids) were easily observed in the discrete nuclear domains with accumulations of multiple capsid proteins and capsids, suggesting that the aggregates of multiple capsid proteins were functional. We also showed in this study that the discrete nuclear domains with accumulations of multiple capsid proteins induced by the VP26 null mutation appeared to be spatially separate from the replication compartments in which viral genome DNA replication takes place. As described above, it has been proposed that HSV-1 capsid proteins may be recruited to and assembled in replication compartments to facilitate efficient packaging of replicated viral DNA genomes into capsids (23, 24). Therefore, VP26 and perhaps UL25 and UL17 may facilitate packaging of replicated viral DNA genomes into capsids by regulating proper localization of capsids in the replication compartments in the nuclei of HSV-1-infected cells. Further studies to clarify the mechanism of the effect(s) of VP26 on the conformation and/or the stability of capsids are of interest and under way in this laboratory.
MATERIALS AND METHODS
Cells and viruses.Vero and rabbit skin cells have been described previously (49). The HSV-1(F) wild-type strain and the VP26 null mutant virus YK490 (ΔVP26) (Fig. 1) have been reported previously (19).
Plasmids.Transfer plasmid pBS-VP26-kpn+, used for generating recombinant viruses YK495 (ΔVP26-repair) and YK499 (UL17-Myc/Flag-UL25/ΔVP26-repair), in which the VP26 null mutations in YK490 (ΔVP26) and YK497 (UL17-Myc/Flag-UL25/ΔVP26), respectively, were repaired, have been described previously (19). To generate a fusion protein of the maltose binding protein (MBP) and a domain of UL17 or UL25, pMAL-UL17:154–703 and pMAL-UL25:300–581 were constructed by cloning the UL17 domain encoding codons 154 to 703 and the UL25 domain encoding codons 300 to 581, respectively, that had been amplified by PCR from the HSV-1(F) genome, into pMAL-c (New England BioLabs) in frame with MBP.
Mutagenesis of viral genomes and generation of recombinant viruses.Recombinant virus YK496 encoding Flag-tagged UL25 in which the Flag epitope was inserted in frame between UL25 amino acids 50 and 51 (Fig. 1) was constructed by a two-step red-mediated mutagenesis procedure using Escherichia coli GS1783 carrying pYEbac102, a full-length infectious HSV-1(F) clone (49), as described previously (50), except with the following primers: 5′-GTCGCCCGTCTTTAACCTCCCCCGGGAGACGGCGGCGGAGGACTACAAAGACGATGACGACAAGCAGGTGGTCGTCCTACAGGCAGGATGACGACGATAAGTAGGG-3′ and 5′-CGGCAGCCGCTGTGCGCTGGGCCTGTAGGACGACCACCTGCTTGTCGTCATCGTCTTTGTAGTCCTCCGCCGCCGTCTCCCGGGCAACCAATTAACCAATTCTGATTAG-3′. Recombinant virus YK497 (UL17-Myc/Flag-UL25) encoding Flag-tagged UL25 and UL17 tagged with the Myc epitope at its C terminus (Fig. 1) was constructed as described previously (50) except using E. coli GS1783 carrying the YK496 genome and primers 5′-GTTTTGTCGCAAGGTGTCGTCCGGGAACGGCCGGTCTCGCGAACAAAAACTCATCTCAGAAGAGGATCTGGCGGGCGCCTTCCCCCGGCCAGGATGACGACGATAAGTAGGG-3′ and 5′-GGAGGAGTGGATGGGCGAGGTGGCCGGGGGAAGGCGCCCGCTACAGATCCTCTTCTGAGATGAGTTTTTGTTCGCGAGAACGGCCGTTCCCGGCAACCAATTAACCAATTCTGATTAG-3′. Recombinant virus YK498 (UL17-Myc/Flag-UL25/ΔVP26), in which UL25 and UL17 were tagged with Flag and Myc, respectively, and the UL35 gene encoding VP26 was disrupted by replacing UL35 codons 28 to 306 with a foreign gene cassette containing a kanamycin resistance gene (Fig. 1), was generated by the red-mediated mutagenesis procedure as described previously (50), except using E. coli GS1783 carrying the YK497 (UL17-Myc/Flag-UL25) genome and primers 5′-ACCTCCGGTCCCGATGGCCGTCCCGCAATTTCACCGCCCCAGGATGACGACGATAAGTAGGG-3′ and 5′-CTGGGCCTCACGGGGTCCCGGGCGTCGAAGGTTCTCGAACCAACCAATTAACCAATTCTGATTAG-3′. Recombinant viruses YK495 (ΔVP26-repair) and YK499 (UL17-Myc/Flag-UL25-/ΔVP26-repair) (Fig. 1), in which the VP26 null mutations in YK490 (ΔVP26) and YK498 (UL17-Myc/Flag-UL25/ΔVP26), respectively, were repaired, were generated by cotransfection of rabbit skin cells with pBS-VP26-kpn+ and the YK490 (ΔVP26) or YK498 (UL17-Myc/Flag-UL25/ΔVP26) genome as described previously (51). Plaques of progeny viruses from the transfected cells were isolated three times on Vero cells, and restoration of the original VP26 sequence was confirmed by sequence analysis.
Production and purification of MBP fusion proteins expressed in E. coli.MBP fusion proteins MBP-UL17:154–703 and MBP-UL25:300–581 were expressed in E. coli that had been transformed with pMAL-UL17:154–703 and pMAL-UL25:300–581, respectively, and purified using amylose resin (New England BioLabs) as described previously (50).
Antibodies.Commercial mouse monoclonal antibodies to Flag (M2; Sigma), Myc (PL14; MBL), α-tubulin (DM1A; Sigma), VP5 (3B6; Virusys), and HSV-1 single-stranded DNA (ssDNA) binding protein ICP8 (HB-8180; ATCC) and commercial rabbit polyclonal antibodies to VP23 (CAC-CT-HSV-UL18; CosmoBio) and Myc (562; MBL) were used. Rabbit polyclonal antibody to VP26 has been described previously (28). To generate mouse polyclonal antibodies to HSV-1 UL17 or UL25, BALB/c mice were immunized with the purified MBP fusion protein MBP-UL17:154–703 or MBP-UL25:300–581 as described previously (52). These antibodies were used for immunoblotting, but did not work in immunofluorescence studies (data not shown).
Immunoblotting and immunofluorescence.Immunoblotting was performed as described previously (53). The amount of protein in immunoblot bands was quantified using the ImageQuant LAS 4000 system with ImageQuant TL7.0 analysis software (GE Healthcare Life Sciences). Immunofluorescence was performed as described previously (54), except that the samples were examined with an LSM 800 laser scanning confocal microscope with ZEN software, version 2.3 (Zeiss).
Purification of virions.Vero cells were infected with each of the indicated viruses at an MOI of 0.01 for 48 h. Cell culture supernatants containing extracellular virus were harvested, and virions were purified as described previously (28).
Purification of nuclear capsids.Nuclei of infected cells were isolated and fractionated, and capsids were purified from specific fractions as described previously, but with minor modifications (55, 56). Briefly, Vero cells were infected with each of the viruses at an MOI of 3 for 18 h. The cells were then harvested, washed with phosphate-buffered saline (PBS), resuspended in hypotonic buffer (10 mM Tris-HCl pH 7.5, 10 mM KCl, 3 mM MgCl2, 0.05% Nonidet P-40 [NP-40], 1 mM EDTA, 1 mM dithiothreitol [DTT], 10 mM NaF, 1% protease inhibitor cocktail [Nakarai Tesque]), and incubated on ice for 15 min. The nuclei were pelleted using low-speed centrifugation (250 × g, 8 min at 4°C), washed with the hypotonic buffer, resuspended in TNE buffer (500 mM NaCl, 1 mM EDTA, 20 mM Tris-HCl [pH 7.5]), and sonicated on ice three times for 15 s each. Cell debris was cleared by centrifugation (2,200 × g, 5 min at 4°C), and nuclear capsids in the supernatants were pelleted through a 35% (wt/wt) sucrose cushion (in TNE buffer) in a P40ST rotor (Hitachi) with centrifugation at 110,000 × g for 1 h at 4°C. Pellets were resuspended in TNE buffer by sonication on ice three times for 15 s each, layered onto a 20 to 50% (wt/wt) sucrose gradient (in TNE buffer), and centrifuged at 110,000 × g for 1 h at 4°C in a P40ST rotor. After centrifugation, 0.3-ml fractions were collected from the top of each tube. Trichloroacetic acid (TCA) was added to each fraction to a final concentration of 10% TCA, and the fractions were incubated overnight at 4°C. Precipitated proteins were pelleted by centrifugation at 21,000 × g for 20 min at 4°C, washed two times with ethanol, dried, and analyzed by immunoblotting.
DNase I protection assays.DNase I protection assays were performed as described previously but with minor modifications (25). Briefly, Vero cells were infected with each virus at an MOI of 3 for 18 h. The cells were then harvested, and nuclei were prepared as described above, resuspended in 10 mM Tris-HCl (pH 7.5), and incubated on ice for 15 min. NP-40 was added to each sample to a final concentration of 1.0%, and each sample was rotated at 4°C for 30 min. Each sample was then centrifuged for 5 min at 2,200 × g, and the supernatant was removed and split into two equal parts. One part was incubated on ice, and the other was digested with 20 U of DNase I (TaKaRa) for 3 h at 37°C. EDTA was added to both samples to a final concentration of 20 mM, and the samples were then treated with proteinase K for 1 h at 50°C. Viral DNA was extracted with phenol-chloroform and precipitated with ethanol.
Southern blotting.Viral DNAs were digested with BamHI, analyzed by electrophoresis on 0.8% agarose gels, and transferred to Hybond-N+ nylon membranes (GE Healthcare). The bands were hybridized with the appropriate DNA probe labeled with horseradish peroxidase using a Direct Nucleic Acid Labeling System (GE Healthcare) as instructed by the manufacturer. The amount of DNA in the Southern blot bands was quantified using the ImageQuant LAS 4000 system with ImageQuant TL7.0 analysis software (GE Healthcare Life Sciences).
Electron microscopic analysis.Vero cells infected with each virus at an MOI of 5 for 18 h were examined by ultrathin-section electron microscopy as described previously (57).
Correlative light and electron microscopy (CLEM) analysis.Vero cells were grown to 1.0 × 105 cells in 35-mm glass-bottom grid dishes (Matsunami) to aid the localization of individual cells and then infected with each virus at an MOI of 5 for 18 h. The cells were subsequently fixed with 4% paraformaldehyde and 0.1% glutaraldehyde in PBS on ice for 1 h and permeabilized with 0.1% Triton X-100 in PBS at room temperature for 15 min. The glutaraldehyde was quenched by the addition of l-lysine monohydrochloride to a final concentration of 0.2 M for 30 min at room temperature. The cells were blocked with 3% bovine serum albumin and 10% calf serum in PBS and reacted with anti-VP5 antibody and then with anti-mouse IgG-Alexa Fluor 488 (Invitrogen). Fluorescence signals in the cells were observed with an LSM 800 laser scanning confocal microscope. The cells were then washed with PBS and fixed further with 2.5% glutaraldehyde in PBS overnight at 4°C and subsequently with 2.5% glutaraldehyde in 0.1 M sodium phosphate buffer (pH 7.4) overnight at 4°C. After fixation, the cells were stained with 2% OSO4 in cacodylate buffer (0.05 M cacodylate, 2 mM CaCl2) on ice for 2 h, washed twice with H2O, incubated in 1% uranyl acetate on ice for 1 h, washed with H2O overnight at 4°C, dehydrated with an ethanol gradient series, and flat-embedded in an Epon mixture. Ultrathin sections cut parallel to the substrate were used to identify cells characterized by the nuclear aggregate morphology of VP5 that had been observed by immunofluorescence. Such ultrathin sections were stained with uranyl acetate and lead citrate and examined with a Hitachi H7500 electron microscope.
ACKNOWLEDGMENTS
We thank Tomoko Ando and Yoshie Asakura for their excellent technical assistance.
This study was supported by Grants for Scientific Research from the Japan Society for the Promotion of Science, grants for Scientific Research on Innovative Areas from the Ministry of Education, Culture, Science, Sports and Technology of Japan (16H06433, 16H06429, and 16K21723), a contract research fund for the Program of Japan Initiative for Global Research Network on Infectious Diseases from the Japan Agency for Medical Research and Development, and grants from the Takeda Science Foundation and the Mitsubishi Foundation.
FOOTNOTES
- Received 26 June 2017.
- Accepted 27 June 2017.
- Accepted manuscript posted online 5 July 2017.
Supplemental material for this article may be found at https://doi.org/10.1128/JVI.01068-17 .
- Copyright © 2017 American Society for Microbiology.
