INTRODUCTION

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The mechanisms of envelopment and intracellular trafficking of herpes simplex virus (HSV) are poorly understood. A number of ultrastructural studies have shown that viral nucleocapsids containing packaged DNA are assembled in the infected cell nucleus and then bud through the inner nuclear membrane into the perinuclear space, acquiring an envelope (62, 63, 66). However, the interpretation of images representing subsequent stages in virus egress has been controversial. Electron microscopy reveals that the infected cell cytoplasm contains capsids partially wrapped by adjacent membranes, enveloped virions completely enclosed within membranous vesicles, and "naked" capsids entirely lacking envelopes (4, 14, 38, 41, 72).

Two principal HSV egress models have been proposed to account for these various cytoplasmic structures (27, 80). In one model, it has been suggested that perinuclear enveloped virions enter transport vesicles budding from the outer nuclear membrane or endoplasmic reticulum (ER) and then traffic as normal secretory pathway cargo through the Golgi apparatus and are eventually secreted from the cell (25, 38, 66, 72). In this model, naked cytoplasmic capsids arise due to nonproductive, erroneous deenvelopment, wherein the envelope of trafficking virions becomes fused with the surrounding lipid bilayer of the transport vesicle. Increased accumulation of naked capsids in the cytoplasm of cells infected by an HSV strain mutant in glycoprotein gD supports the view that premature fusion can generate nonenveloped capsids (14); however, it is not clear if such a phenomenon is the source of naked capsids during a wild-type viral infection.

In an alternative model (65), perinuclear virions undergo a programmed deenvelopment in which they fuse their envelopes with the outer nuclear membrane, releasing naked capsids into the cytoplasm. These capsids lie on a productive route of assembly and subsequently reenvelope by budding into the lumen of some organelle of the secretory or endocytic pathways (41). Consistent with this reenvelopment model are the recent observations that retention of either of the viral glycoproteins gH and gD in the ER prevents their incorporation into the envelopes of secreted virions (11, 79). Similarly, examination of the phospholipid composition of extracellular HSV particles revealed that the viral membrane is enriched in lipids which are most abundant in organelles other than the ER (76), and studies in explanted neurons have shown that the anterograde transport of HSV virions along axons appears to involve distinct pathways for nucleocapsid/tegument and glycoproteins (35, 36, 49, 55). Similar reenvelopment egress pathways have been proposed for other herpesviruses, based primarily on electron microscopic observations (15, 29, 30, 34, 39, 51, 71, 78, 84). Evidence for and against each of these models of egress has recently been comprehensively reviewed (27).

Whichever egress model is correct, the identity of those cytoplasmic organelles in which enveloped virions accumulate is unclear. Similarly, the kinetics with which capsids pass through the egress pathway and the relative abundance of naked and enveloped capsids in various cytoplasmic compartments are questions which have been difficult to address. One reason for this is that infected cells replicate HSV continuously for 18 to 24 h. They therefore contain viruses at every stage of assembly, making it impossible to determine the order of events during viral biogenesis. Furthermore, electron microscopic techniques cannot reveal whether a particular compartment contains infectious or defective particles.

We have described a model system to facilitate the study of late events in HSV assembly. The HSV-1 strain tsProt.A carries a reversible temperature-sensitive lesion in UL26, which encodes the maturational protease Pra (28, 48, 57, 58). At the nonpermissive temperature of 39°C, tsProt.A-infected cells accumulate immature nuclear procapsids (54, 59). Following downshift to the permissive temperature of 31°C, these procapsids recruit the capsid subunit VP26 (21, 54), package DNA (22, 24, 57), and give rise to exocytosing infectious particles in a synchronous wave (23). In the present study, we make use of this synchronized assembly system, in combination with biochemical assays for packaged capsids and enveloped viral particles, to examine virus-containing organelles by differential ultracentrifugation. This has enabled us to characterize which cytoplasmic organelles are associated with infectious and noninfectious virions, to determine the relative abundance of capsids and enveloped virions in various compartments, and to monitor the kinetics with which exiting virus traverses the cytoplasm. Our data show that enveloped infectious virions accumulate in organelles with biochemical properties similar to those of the trans-Golgi network (TGN) and endosomes and that nonenveloped capsids are also tethered to the cytoplasmic face of these organelles. No HSV appeared to accumulate in earlier compartments of the Golgi apparatus or in lysosomes. Chloroquine or bafilomycin A1, which neutralize the acidic pH of the endosomal/recycling network, dramatically decreased the yield of infectious virions, even though the number of enveloped particles remained similar to untreated controls. We conclude that, under our conditions, TGN and/or endosomes are key organelles in HSV exocytosis and that an acidic pH is critical for cytoplasmic viral particles to acquire infectivity.

	MATERIALS AND METHODS

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Cells, media, and viruses. Human hepatoma (HuH7) cells were maintained in RPMI supplemented with 1% penicillin-streptomycin (PS) (Gibco Laboratories) and 10% fetal calf serum (FCS). Vero cells were grown in Dulbecco's modified Eagle's medium (DMEM)-1% PS supplemented with 10% newborn calf serum (NCS). The HSV-1 strains tsProt.A and SC16 were prepared as previously described (23). Plaque assays were done by titration on preformed Vero cell monolayers. Chloroquine and brefeldin A (BFA) were obtained from Sigma, and bafilomycin A₁ was from Kamiya Biomedical, Seattle, Wash.

Western blotting and antibodies. Extracts were boiled in Laemmli buffer and then subjected to denaturing sodium dodecyl sulfate-polyacrylamide gel electrophoresis using 8 or 12% resolving gels. Proteins were transferred to a nitrocellulose membrane at 80 mA for 2 h with a Hoeffer semidry blotter, and then the membrane was blocked for 1 h at room temperature in Tris-buffered saline (150 mM NaCl, 10 mM Tris-HCl [pH 7.6]) supplemented with 0.5% Tween 20 and 5% nonfat dried milk. Membranes were then incubated overnight at 4°C with the appropriate antibody. Antibodies were obtained from the following sources: anti-COP monoclonal clone maD was from Sigma; affinity-purified anti-TGN46 antiserum was from Serotec; anti--adaptin, anti-EEA1, and anti-rab5 (clone C1) monoclonals were from Transduction Laboratories; polyclonal anticalnexin was from Stressgen Biotech. Corp.; and anti-PCNA monoclonal clone PC10 was from Santa Cruz Biotechnology. For anti-LAMP1 monoclonal clone H4A3, anti-rab7, and anti-p115, see the Acknowledgments. Secondary antibody detection was done by Pierce Supersignal chemiluminescence.

Cell labeling and total-extract and PNS preparation. HuH7 cells at 70 to 80% confluence were incubated for 2 h at 37°C in RPMI medium containing 1% dialyzed FCS. They were then infected at a multiplicity of infection (MOI) of 10 with HSV strain tsProt.A for 1 h at 37°C, and unpenetrated virus was inactivated by a wash with glycine-buffered saline (GBS; 136 mM NaCl, 5 mM KCl, 100 mM glycine [pH 2.8]), then overlaid with warmed RPMI-1% dialyzed FCS containing [³H]thymidine at a final concentration of 25 µCi/ml, and incubated for a further 7 h at 39°C (nonpermissive temperature) before downshift to 31°C (permissive temperature). At 30 min prior to downshift, cycloheximide (20 µg/ml) was added to ensure a single pulse of viral assembly. To prepare total extracts, cells were rinsed twice in TBS_B (130 mM NaCl, 20 mM KCl, 25 mM Tris-HCl [pH 7.6]) and then frozen, thawed, collected by scraping, and sonicated. Alternatively, cells were rinsed twice in homogenization buffer (HB_A, 250 mM sucrose, 2 mM MgCl₂, 10 mM Tris-HCl [pH 7.6]) collected by scraping, homogenized by eight passages through a 25^5/8-gauge needle, and then spun at 2,000 × g for 10 min at 4°C to remove unbroken cells and nuclei and yield a postnuclear supernatant (PNS).

Measurement of DNA packaging and capsid envelopment. The trichloroacetic acid (TCA) precipitation assay used to measure DNA packaging was modified from our earlier published study (24) as follows. Cell extracts or gradient fractions were incubated in the presence of 2 mM MgCl₂ and 280 U of DNase I (Sigma; type II) per ml for 90 min at 37°C. EDTA and SDS were then added to a final concentration of 10 mM and 0.3%, respectively, and incubation was continued for a further 15 min at 37°C before spotting onto individual GF/C Whatman filters. Each filter was subjected to one 4°C wash and two consecutive 65°C washes in TP buffer (5% TCA, 20 mM sodium pyrophosphate) before being rinsed in 70% ethanol at room temperature and dried. Levels of TCA-precipitable radioactivity were determined by liquid scintillation counting. To measure only that DNA present in enveloped capsids, samples were first incubated with 0.2 mg of proteinase K per ml for 90 min at 37°C to destroy nonenveloped capsids. The reaction was quenched by addition of 2 mM phenylmethylsulfonyl fluoride and then subjected to DNase I treatment and TCA precipitation as above.

Percoll density gradient centrifugation. Generally, four to five 15-cm dishes of HuH7 cells at 70 to 80% confluency were used for each gradient. Cells were washed twice with HB_A, and a PNS was prepared as described above. The PNS was mixed with stock Percoll solution to prepare 11 ml of a solution of 1.065 g of Percoll per ml in 250 mM sucrose, as per the manufacturer's instructions (Pharmacia Biotech). A self-forming gradient was produced by centrifugation for 45 min at 20,000 rpm (36,000 × g) in a Ti50 fixed-angle rotor at 4°C, and 22 0.5-ml fractions were collected from the top of the gradient.

Sucrose float-up gradients. Cells were washed twice with HB_A, and 1 ml of PNS was prepared as described above, of which 0.9 ml was adjusted to 1.4 M sucrose and loaded onto a step gradient as follows: 1 ml of 2 M sucrose overlaid with 2 ml of 1.4 M adjusted PNS, 7.5 ml of 1.2 M sucrose, and 1.5 ml of 0.8 M sucrose; all solutions contained 10 mM Tris-HCl (pH 7.6) and 2 mM MgCl₂. The gradient was centrifuged for 4 h at 39,000 rpm (261,000 × g) in an SW41 rotor at 4°C, and 12 1-ml fractions were collected from the top of the gradient.

HRP uptake to label endocytic compartments. After 16 h of HSV infection, HuH7 cells were washed three times with warmed serum-free RPMI. Serum-free RPMI containing 1.4 mg of horseradish peroxidase (HRP; obtained from Sigma) per ml was then added to each dish, and the cells were incubated at 37°C for 12 min. The cells were subsequently transferred to ice and washed five times (2 min per wash) with cold phosphate-buffered saline (PBS)-1 mM MgCl₂-1 mM CaCl₂-0.1% bovine serum albumin (BSA) and then washed twice with HB_A. The cells were scraped from the dish into HB_A, and a PNS was prepared and fractionated as described above. Gradient fractions were blotted onto a nitrocellulose membrane using a Hoeffer vacuum blotting apparatus, and bound HRP was detected directly by incubation of the membrane with Supersignal Chemiluminescence (Pierce).

Measurement of Golgi glycosyltransferase activities in gradient fractions. Galactosyltransferase activity was determined as described by Chaney and colleagues (16). Sialyltransferase activity was measured by incubating each gradient fraction with asialofetuin (20 mg/ml; Sigma) in 1.25% Triton X-100-50 mM MOPS (morpholinopropanesulfonic acid, pH 6.5)-5 mM MnCl₂-1 mM CMP-[³H]sialic acid (New England Nuclear; adjusted to a final specific activity of 2,000 cpm/nmol using unlabeled CMP-sialic acid from Sigma) for 1 h at 37°C. The reaction was stopped by addition of 1 ml of ice-cold water, and then samples were TCA precipitated onto GF/C filters, and precipitable cpm were determined by liquid scintillation counting.

Transmission electron microscopy. Fractions from sucrose float-up gradients were pelleted in an Airfuge at ~26 lb/in² for 20 min at 4°C. The pellet was fixed in 2.5% glutaraldehyde in SC (100 mM sodium cacodylate [pH 7.43]) at room temperature for 45 min, rinsed in SC, postfixed in 1% osmium tetroxide in SC followed by 1% uranyl acetate, dehydrated through a graded series of ethanols, and embedded in LX112 resin (LADD Research Industries, Burlington, Vt.). Ultrathin sections were cut on a Reichert Ultracut E, stained with uranyl acetate followed by lead citrate, and viewed on a Joel 1200EX transmission electron microscope at 80 kV.

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Development of rapid biochemical assays to detect packaged and enveloped capsids. We first wished to develop a rapid and quantitative means of assaying for the presence of naked and enveloped intracellular capsids. Our previously described TCA precipitation assay monitors the incorporation of [³H]thymidine-labeled DNA into capsids by measuring DNase I-resistant TCA-precipitable counts (22, 24) in whole-cell extracts and thus scores total packaged capsids. To measure the proportion of packaged capsids which are enveloped, we modified our conditions by digesting cell extracts with proteinase K prior to DNase I treatment and TCA precipitation. We reasoned that after proteinase K incubation, only enveloped virions would contain packaged, DNase I-resistant DNA.

View larger version (16K): [in this window] [in a new window]   FIG. 1.   Measuring DNA packaging and capsid envelopment using a TCA precipitation assay. HuH7 cells were infected with the HSV strain tsProt.A, acid washed, and then incubated at 39°C in the presence of [3H]thymidine. After 6.5 h, cycloheximide was added, and after a further 30 min the cells were downshifted to 31°C. Total cell extracts were prepared at particular times after downshift (indicated on the horizontal axes) and measured as follows. (A) DNase I-resistant (total packaged) DNA in the presence (squares) or absence (triangles) of 0.05% Triton X-100. (B) Proteinase K/DNase I-resistant (enveloped) DNA in the presence (solid squares) or absence (solid triangles) of 0.05% Triton X-100. (C) PFU present in each extract. In each case, plotted values represent the mean of two to four samples, and error bars indicate the range from the mean.

Biochemical detection of cytoplasmic virus particles. We next tested whether these assays could be used to examine those virus particles residing in the cytoplasm of HSV-infected cells. To do this, HuH7 cells were infected at an MOI of 10 with tsProt.A and then incubated at 31°C for 16 h in the presence of [³H]thymidine, and a PNS was prepared. Samples were subjected to density gradient ultracentrifugation on a 1.065-g/ml Percoll gradient, and fractions were collected from the top of the gradient. Each fraction was assayed for total packaged and enveloped DNA, and PFU were counted. As a control, we also subjected [³H]thymidine-labeled extracellular secreted virus to similar centrifugation.

View larger version (17K): [in this window] [in a new window]   FIG. 2.   Intracellular cytoplasmic virus is less dense than extracellular virus. [3H]thymidine-labeled extracellular virus (A and B) or PNS intracellular virus (C and D) from tsProt.A-infected HuH7 cells were subjected to isopycnic centrifugation on a 1.065-g/ml self-forming Percoll gradient. Gradients were fractionated from the top (corresponding to fraction 1), as indicated on the horizontal axes. (A and C) Fractions were assayed for total packaged DNA (open triangles) or proteinase K-protected, enveloped DNA (solid triangles). (B and D) Fractions were titrated for PFU (open circles). Plotted values represent the means of duplicate samples, and error bars indicate the range from the mean.

Cytoplasmic virus cofractionates with Golgi and endosomal markers. To further analyze the subcellular distribution of HSV, radiolabeled intracellular and extracellular virus were prepared as before and then subjected to a sucrose float-up equilibrium gradient (7). Gradients were fractionated from the top into 12 1-ml fractions, and each fraction was assayed for packaged and enveloped DNA, tested for infectivity by PFU titration, and subjected to Western blot analysis. As shown in Fig. 3A, under these conditions three populations of intracellular viral particles were observed by TCA precipitation assay: a buoyant, light population (peak I) lying at the top of the gradient at the interface between the 0.8 M and 1.2 M sucrose layers in fraction 2, and two denser populations of virus (peaks II and III) within the 1.4 M and 2.0 M sucrose layers, in fractions 9 and 11-12, respectively. As previously observed for intracellular virus, the cpm derived from packaged capsids is greater than that for enveloped capsids (Fig. 3A, peaks I and II). Interestingly, although all three peaks appear to contain enveloped virus (Fig. 3A), PFU were only present in peak I and peak II, not in peak III (Fig. 3C). When every gradient fraction was taken into account, 72% of the total capsid population was enveloped under these conditions.

View larger version (76K): [in this window] [in a new window]   FIG. 3.   Fractionation of intracellular virus by sucrose equilibrium density gradient centrifugation. Intracellular (A and C) or extracellular (D and E) virus was prepared as described for Fig. 2 and subjected to sucrose density gradient centrifugation. Fractions were collected from the top of the gradient, as indicated on the horizontal axes (fraction 1 is at the top of the gradient). (A) Top: Intracellular fractions were assayed for packaged and enveloped DNA (open and solid symbols, respectively). The three peaks of radioactivity are labeled I, II, and III. Bottom: Intracellular fractions were subjected to Western blot analysis for the organelle markers rab7 (late endosomes), rab5 and EEA1 (early endosomes), HRP (bulk-phase endocytic compartments), p115 and -COP (Golgi apparatus), -adaptin (TGN, clathrin-coated vesicles), TGN46 (trans-Golgi network), calnexin (microsomesER), and LAMP1 (lysosomes). (B) PNSs were subjected to a 100,000 × g spin, and the resulting pellet (P100) and supernatant (S100) were Western blotted for the antigens indicated at the left. (C) Intracellular fractions were titrated for PFU. (D) Extracellular virus gradient fractions were assayed for packaged and enveloped DNA (open and solid symbols, respectively). (E) Extracellular virus gradient fractions were titrated for PFU. All plotted values represent the mean of duplicate samples, and error bars indicate the range from the mean.

Ultrastructural features of gradient fractions from tsProt.A-infected cells. To further investigate the nature of virus particles in peaks I, II, and III, a sample of each was subjected to transmission electron microscopy. Consistent with our previous biochemical data, peak I was found to contain enveloped virions fully enclosed by a smooth membrane and also naked capsids in close proximity, and presumably attached, to these organelles (Fig. 4). Interestingly, an electron-dense tegument-like material can often be observed at the surface of these membranes and between the capsid and the bilayer. These structures are similar to those observed in infected cell cytoplasms and are interpreted to represent capsids either budding into or erroneously emerging from cellular organelles (14, 27). Analysis of peak II material revealed the presence of both naked capsids and singularly enveloped virus particles, as shown in Fig. 5. These structures are entirely consistent with the biochemical observation that mature extracellular virus fractionated at this position upon gradient centrifugation, and peak II virions therefore most likely represent enveloped capsids released from broken peak I organelles. We also observed, at lower frequency, aberrant multicapsid clusters surrounded by a single membrane bilayer (Fig. 5C). Analysis of peak III material revealed the presence of naked capsids (Fig. 6B) or large membrane-bound structures which contain chromatin-like electron-dense material and single or clustered capsids (Fig. 6A and C) and which appear to be fragments of cells and nuclei. The presence of capsids within nuclear fragments explains why our biochemical assays detected an apparent envelopment signal in peak III despite the absence of infectivity (Fig. 3A and C).

View larger version (207K): [in this window] [in a new window]   FIG. 4.   Ultrastructural analysis of peak I. Material from peak I was pelleted and fixed in glutaraldehyde and prepared for transmission electron microscopy as described in the text. (A) Enveloped virus particle enclosed within a smooth membrane compartment (indicated by the black arrow head) and naked capsids in close proximity to an organellar membrane (indicated by the white arrowhead). (B, C, and D) Additional representative images from peak I. Bar, 0.1 µm.

View larger version (178K): [in this window] [in a new window]   FIG. 5.   Ultrastructural analysis of peak II. Extracts were prepared as described for Fig. 4. (A) Representative section showing mainly naked capsids. (B and D) Presence of singularly enveloped virus particles and (C) a cluster of capsids enclosed within a single membrane, a minor population in this fraction. Bar, 0.1 µm.

View larger version (204K): [in this window] [in a new window]   FIG. 6.   Ultrastructural analysis of peak III material. Extracts were prepared as for Fig. 4. (A) Nuclear fragment containing clusters of naked capsids or single capsids (indicated by the white arrows). The region in the upper left, indicated by an arrow, is magnified in panel C. (B) Free naked capsids. Bar, 0.1 µm.

Kinetics of viral egress. To further test the relationship between peak I and peak II, we determined the kinetics of delivery of viral particles to cytoplasmic compartments during a synchronous wave of viral egress. The experimental approach is outlined in Fig. 7A. HuH7 cells were infected at an MOI of 10 for 1 h at 37°C with HSV strain tsProt.A, the residual input virus was removed by acid wash, and cells were incubated at 39°C for 7 h. At 30 min prior to downshift, cycloheximide was added, and the cells were downshifted to 31°C for 0, 2, 4, and 6 h or maintained at 39°C for a further 6 h. At each time point, PNSs were collected and subjected to sucrose gradient centrifugation. Each fraction was also titrated for infectivity.

View larger version (134K): [in this window] [in a new window]   FIG. 7.   Time course of delivery of virus to cytoplasmic organelles. (A) Schematic showing the time course and conditions for this experiment. HuH7 cells were infected with tsProt.A and incubated for 7 h at 39°C in the presence of [3H]thymidine (cycloheximide was added 30 min prior to downshift). Cells were downshifted to 31°C for 0, 2, 4, or 6 h or kept at 39°C for a further 6 h. For each time point, a PNS was prepared and subjected to sucrose gradient centrifugation, and fractions were titrated for PFU. BAF/CQ indicates the time of addition of the drugs bafilomycin A1 and chloroquine in the experiments shown in Fig. 9. (B to D) PFU yields 2, 4 and 6 h after downshift, respectively. Fractions were also assayed for total packaged DNA (light bars) and enveloped DNA (dark bars) at 2, 4, 6, and 0 h after downshift to 31°C (panels E to H, respectively) or after an additional 6 h at 39°C (panel I). Plotted values are means of duplicates, and error bars indicate the range from the mean. An immunoblot for a nuclear marker, proliferating cell nuclear antigen (PCNA), was performed on fractions from a representative 4-h-downshifted gradient (F).

Separation of Golgi from TGN and endosomes using brefeldin A. From the data in Fig. 3A, peak I appears to contain markers characteristic of endosomes (rab5 and rab7) and is the exclusive site of accumulation of a bulk-phase endocytic marker (exogenously added HRP). However, it also contains the TGN markers -adaptin and TGN46 and (as shown in Fig. 8 below) glycosyltransferase activities characteristic of the trans-Golgi cisternae. We attempted a number of density gradient centrifugation techniques to separate these organelles and to determine which compartments contain infectious HSV, but were unsuccessful. We therefore chose to exploit the properties of the drug brefeldin A (BFA). BFA addition is known to cause disassembly of the cis-, medial-, and trans-Golgi cisternae and their fusion with the ER. In contrast, the TGN appears to fuse and mix with the endocytic network (47, 81). This enables density gradient separation of the TGN and the endosomal apparatus from Golgi, since fusion of Golgi with the ribosome-studded ER greatly increases the density of Golgi-derived membranes.

View larger version (71K): [in this window] [in a new window]   FIG. 8.   Use of BFA to separate Golgi cisternae from TGN/endosomes. An experiment similar to that in Fig. 3 was performed, except that cells were incubated with BFA (5 µg/ml) prepared in ethanol (righthand panels, solid symbols) or with an equivalent volume of ethanol alone (lefthand panels, open symbols) for 1 h prior to PNS preparation and fractionation. Fractions were assayed for galactosyltransferase activity (A), sialyltransferase activity (B), or PFU (C) or Western blotted for rab5, rab7, and TGN46 (D). (E) Peak I was collected following fractionation of BFA-treated and mock-treated cells (as indicated) and then Western blotted as in panel D. Lanes contained 100, 60, or 40% (as indicated at the top of the figure) of the peak I material in panel D.

Neutralization of organellar pH. The TGN and endosomes are known to maintain a low internal pH (between approximately 5 and 6.5), which is required for their normal biological function. If these organelles truly play a critical role in HSV assembly and egress, then we reasoned that these processes might be interrupted by addition of the weak base chloroquine (17) or the drug bafilomycin A1, which inhibits the proton ATPase responsible for acidification (2, 6, 10, 82, 83).

View larger version (24K): [in this window] [in a new window]   FIG. 9.   Infectivity of virus particles within cytoplasmic organelles is dependent on an acidic environment. HuH7 cells were infected with the HSV-1 strain tsProt.A, acid washed, and then incubated at 39°C for 7 h. Cycloheximide (20 µg/ml) was added 30 min prior to downshift to 31°C. At the time of downshift, cells received 150 µM chloroquine or 200 nM bafilomycin A1 or no treatment as shown in Fig. 7A. Cells were then incubated at 31°C for 9 h. PNSs were subjected to sucrose gradient centrifugation and fractionated. (A) PFU titrations of gradient fractions from untreated cells (light gray bars) and cells treated with chloroquine (dark gray bars) or bafilomycin A1 (open bars). (B) Assay for enveloped virions present in untreated cells and cells treated with chloroquine or bafilomycin A1 (indicated as in A). All values plotted are means of duplicate samples, and error bars indicate the range from the mean.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

A major challenge in the study of HSV-1 egress from infected cells has been the interpretation of electron microscopic images showing cytoplasmic virus adjacent to or within membranous organelles. The relevance of these various structures to assembly and exocytosis of infectious HSV particles has been much discussed, and several hypotheses have been suggested to explain their role and origin (27). In this study we describe a biochemical approach to the analysis of HSV egress. We have developed rapid and quantitative assays which enable us to measure naked and enveloped capsids as they travel through the cytoplasm of infected cells. Using these assays and differential centrifugation techniques, we have isolated a gradient fraction which contains infectious HSV and markers characteristic of the Golgi apparatus, TGN, and endosomes. Addition of the drug BFA enabled further subdivision of this fraction, resulting in removal of the majority of the trans-Golgi (and, by implication, the entire Golgi stack) but leaving behind TGN, endosomes, and all of the PFU. The simplest interpretation of these data is that the HSV particles are present within the lumen of the TGN and/or endocytic network, which in the latter case includes early and late endosomes, and sorting endosomes (50). We cannot discount the possibility that BFA caused disassembly of the trans-Golgi cisternae without affecting the cis or medial compartments and that the PFU in peak I reside exclusively in these BFA-resistant cisternae, bypassing the trans cisternae as they traffic out of the cell. However, selective BFA resistance of cisternae and bypass of the trans-Golgi during trafficking are without precedent, and it is difficult to imagine a mechanism by which such phenomena could occur.

Consistent with a role for TGN and/or the endocytic apparatus in HSV assembly is the effect of organellar pH neutralization. Previous work has shown that the weak bases NH₄Cl and chloroquine inhibit production of infectious HSV particles (5, 42, 43, 45, 46), but in those studies the bases were usually present throughout the infection and could have affected any step in replication. We showed that chloroquine and bafilomycin A1 (a specific drug which blocks endosomal acidification) abolish infectivity in peak I when added after accumulation of synchronized tsProt.A procapsids. These compounds must therefore inhibit some step after procapsid assembly. Consistent with this notion, normal numbers of enveloped though noninfectious particles were found within neutralized organelles. Interestingly, addition of monensin to HSV-infected cells, which would also result in pH neutralization (8, 26), led to accumulation of infectious virions (38), though at diminished levels compared to controls. However, those studies considered PFU in total cell extracts (presumably including any infectious virions in the perinuclear space) rather than the neutralized organelle itself.

Peak I was the only detectable destination for exocytosing virions during the earliest stages of a controlled wave of tsProt.A assembly. PFU reached the organelles in this region of the gradient within 2 h of the start of procapsid maturation; however, these studies were conducted at 31°C, and the rate at 37°C might well be higher (and this 2-h time period includes the time required for capsid maturation and DNA packaging, shown to be 40 to 60 min in our earlier studies [23]). Under these conditions, PFU were not detectable in the medium until 4 h after it was first observed in peak I (data not shown). This suggests that exit of HSV from this compartment is slow at 31°C and also shows that TGN/endosomes are unlikely to contain HSV as a result of having endocytosed it from the medium (consistent with the fact that exogenously added radiolabeled HSV was not detectable in peak I under our conditions). Our results are in agreement with a recent study which showed that gD and the capsid protein VP5 colocalize with the 275-kDa mannose-6-phosphate receptor and the transferrin receptor (12). However, the significance of that finding for productive virus assembly was not addressed.

Since we failed to detect HSV particles in the Golgi apparatus, they may have reached TGN and/or endosomal compartments by a route other than the classical secretory pathway. The simplest interpretation of our data is that enveloped capsids accumulate within these organelles by budding into them from the cytoplasm. Consistent with this notion, biochemical analysis and electron microscopic inspection revealed that peak I organelles contained both lumenal enveloped virions and peripherally attached capsids, resembling structures previously observed by thin-section analysis of herpesvirus-infected cells (29, 39, 71, 72, 78, 84). That the nonenveloped peripherally attached capsids are on a productive pathway is consistent with their abundance (biochemical assays indicated that membrane-attached capsids were similar in abundance to enveloped ones) and the kinetics of their accumulation (during a synchronized wave of assembly they could be detected as early as infectious virions). It is also worth noting that these capsids were biochemically defined by their ability to protect the viral genome from DNase I digestion. They are therefore unlikely to be damaged or degraded, as pointed out for some populations of cytoplasmic capsid (72). Envelopment of HSV at the TGN was predicted by Griffiths and Rottier (32), but cytomegalovirus has been reported to envelope at endosomal membranes (71).

Envelopment at TGN and/or endosomes is consistent with the finding that ER-retained gH and gD are not incorporated into the viral envelope (11, 79) and also with envelope incorporation of TGN-targeted gD (79), which cycles between TGN and plasma membrane in endosomes (9, 37). This model for envelopment predicts that all of the membrane proteins found in the mature HSV particle must traffic through TGN/endosomes too, but there is only limited information concerning recycling of HSV envelope proteins (1, 12). For pseudorabies virus, it is clear that removal of the endocytosis motif from gE does not prevent its incorporation into viral particles (69, 70).

Direct envelopment of capsids at the TGN or endosomes is at odds with data showing enveloped virions in transit through the Golgi (72), but this study did not report what proportion of the total viral population was Golgi associated. TGN/endosomal envelopment would also appear to contradict the finding that gD, when retained in the trans-Golgi using the membrane span of sialyltransferase, still became assembled into the HSV envelope (53, 79). These data can, however, be reconciled in a number of ways. First, the distribution of the gD-sialyltransferase fusion protein within the Golgi stack was not tested by Whiteley and colleagues (79). It is possible that, in the context of an HSV-infected cell, not all of the fusion protein is tightly localized to the trans cisternae; some may have reached later compartments of the Golgi or endosomes. Furthermore, sialyltransferase has been reported to be an enzyme of both the trans-Golgi and TGN in some cell types (19, 64).

An alternative interpretation of our data is to suppose that enveloped HSV does pass through the ER and Golgi en route to TGN/endosomes but traverses these organelles so rapidly under our conditions that it never accumulates within them. We are currently testing this hypothesis by examining synchronized egress at lower temperatures to reduce the rate of trafficking. In this case, membrane-bound cytoplasmic capsids could well be the products of erroneous fusion between the HSV envelope and the bounding membrane (14, 27). Alternatively, it may be important to consider that our synchronized assembly studies were conducted at relatively early times postinfection: perhaps HSV preferentially accumulates within TGN/endosomes at early times, but at later time points occupies other organelles such as the earlier cisternae of the Golgi. Furthermore, HSV may utilize Golgi and TGN/endosomes to various extents in different cell lines; some of the effects of HSV infection on cytoplasmic organelles are clearly cell type dependent (3, 13).

In summary, we believe our data are most consistent with the notion that TGN or endosomes are the principal destination for trafficking HSV during a single wave of egress and that an acidic pH is important for the acquisition or maintenance of infectivity in these compartments. It remains unclear whether this requirement reflects some acid-dependent processing event critical for maturation or the absence from the organelle of essential proteins or lipids. Experiments are currently under way to distinguish between these possibilities and to determine the reason for the defect in particle infectivity.