Varicella-zoster virus (VZV) is a member of the Alphaherpesvirinae and the etiological agent of the ubiquitous childhood viral infection commonly known as “chickenpox”. While VZ virions share many physical and biochemical properties that distinguish herpesviruses, VZV possesses unique pathogenic features that are not shared by the other human neurotropic alphaherpesviruses, herpes simplex virus type 1 and type 2 (HSV) (Cohen et al. 2007).

Primary VZV infection is initiated upon inhalation of respiratory droplets containing infectious particles (Arvin et al. 2010). Robust replication facilitates infection of susceptible T lymphocytes in mucosa-associated lymphoid tissues. VZV-infected T-lymphocytes with skin homing phenotypes infiltrate and infect dermal sites leading to development of the characteristic pruritic rash on the skin surface within 10–21 days (Ku et al. 2002, 2004). The capacity to infect and replicate within T-lymphocytes, which is not shared by HSV, enables two different mechanisms for VZV neurotropism. Sensory nerve ganglia may acquire virus via retrograde axonal transport in neurons innervating cutaneous and mucosal sites of replication or through circulating T cells that enter neuronal ganglia and transfer virus to sensory neurons. Similar to HSV, VZV persists within the infected host as a lifelong latent infection in sensory nerve ganglia from which it may reactivate, resulting in anterograde transport of infectious particles that re-infect dermal sites causing a painful rash known as herpes zoster. Whereas HSV is associated with many reactivation events over the lifetime of the infected individual, VZV reactivation is usually limited to a single event (Cohen et al. 2007). The capacity to cause significant neuropathic pain following an episode of herpes zoster (post-herpetic neuralgia) is another clinical feature that distinguishes VZV from HSV. Differences in the route of neuronal infection and in the frequency and severity of reactivation suggest that VZV and HSV may have evolved different strategies for neurovirulence in the human host.

The pathogenic mechanisms through which VZV causes neurovirulence are difficult to investigate because, unlike HSV, VZV replicates poorly across a species barrier. Human cadaveric tissues from individuals who died during primary varicella infection or reactivation are rare. Examination of latently infected cadaver ganglia cannot provide information concerning early events in VZV neuronal infection. Our approach to investigating VZV neurotropism and neurovirulence has been to develop a mouse–human xenograft model in which human dorsal root ganglia (DRG) are maintained in severe compromised immunodeficient (SCID) mice (Zerboni et al. 2010). Xenotransplanted DRG maintains an organotypic microenvironment so that VZV infection of human neurons can be examined under “in vivo” conditions. In this brief review, we will describe the SCID DRG model and discuss our key findings using this model with respect to initial events in VZV replication, spread, satellite glial cell tropism, polykaryon formation between neurons and encapsulating satellite cells, latency, and the molecular requirements for VZV glycoprotein E (gE) and glycoprotein I (gI).

Methods for implantation and VZV infection of DRG xenografts in SCID mice

DRG xenograft implantation

Our procedures for implantation of human dorsal root ganglia into immunodeficient SCID mice have been previously described, but we will review them briefly (Zerboni et al. 2005, 2007). All of our animal protocols and procedures have been reviewed and approved by the Stanford University Administrative Panel on Laboratory Animal Care. Human tissues are procured by Advanced Bioscience Resources (Alameda, CA) in accordance with state and federal regulations. Tissues are used within 24 h after extraction. Individual dorsal root ganglions, including a portion of the dorsal root, are dissected from fetal spinal tissues and kept on ice until transplanted. A single ganglion (1.5–2 mm3) is loaded into a 16-gauge lubricated transplantation needle. Six-week-old male scid/scid homozygous C.B.17 mice are anesthetized, and a small incision is made along the abdominal wall to expose the left kidney. The kidney capsule is perforated, and the DRG is inserted under the renal capsule. After the transplantation needle is withdrawn, the DRG can be easily seen through the kidney capsule. The abdominal wall and skin incision are closed, and mice are given a post-surgical analgesic and monitored on a warming pad until awakening.

At 4 weeks after transplantation, DRG xenografts can be assessed for viability and engraftment. An incision is made in the abdominal wall to expose the xenograft. At that time, the graft can be macroscopically analyzed for indicators of good engraftment and viability such as pink shiny color and neo-vascularization, indicating anastomosis to the host vasculature (Fig. 1a, arrow denotes implanted DRG). Nonviable xenografts have a dull pale appearance. In general, good engraftment occurs in >95% of surgeries.

Fig. 1
figure 1

a DRG xenograft in SCID mouse 4 weeks after xenotransplantation. b Hematoxylin and eosin (H&E)-stained tissue section of uninfected SCID DRG xenograft. c H&E-stained tissue section; VZV-infected SCID DRG xenograft 14 days after infection with neurons exhibiting cytopathic effect. d DAB (brown)-immunohistochemistry of VZV-infected SCID DRG xenograft 14 days after infection using a rabbit polyclonal antibody to IE63. The majority of neurons express VZV proteins at this time point (thin arrow), but some neurons are observed that do not express VZV proteins (thick arrow). e DAB (brown)-immunohistochemistry of VZV-infected SCID DRG xenograft 56 days after infection using a rabbit polyclonal antibody to IE63, counterstained with hematoxylin. No IE63-positive neurons are observed. f H&E-stained tissue section 56 days after infection; microsatellite cell proliferations indicate neuronal cell loss (dashed arrow), but many neurons persist following resolution of VZV infection. g–i Confocal immunofluorescent staining of DRG infected with rOka at day 14 (g), rOka gEΔCys at day 44 (h), and rOkaΔgI at day 70 (i). Sections are stained with antibody to neural cell adhesion molecule (Alexa-594, red), anti-IE63 (Alexa-488, green), and counterstained with Hoechst. The short arrows in (g–i) indicate neurons with intact NCAM staining of neuronal membranes, which indicates lack of polykaryon formation. The long arrow in (g) indicates polykaryon formation in rOka-infected tissue sections, which occurs in up to 50% of neuron–satellite cell complexes. Some panels in this composite image were adapted from panels that appeared in previous publications (see references): Zerboni et al. 2005, 2007, 2011a

Full size image

Histological evaluation of DRG xenografts

Histological evaluation of DRG xenografts at 4 weeks after transplantation reveals an architecture that resembles normal human DRG, with abundant neuronal cell bodies encapsulated by satellite glial cells, within a network of axons and fibroblasts (Fig. 1b) (Zerboni et al. 2005). Endothelial cells within the microvasculature express the human-specific cell marker PECAM-1 (Zerboni et al. 2005). In the developing human, peripheral sensory neurons arise from three waves of neural crest cell (NCC) migration to DRG (Marmigere and Ernfors 2007; Montelius et al. 2007). Fetal tissues are used at 18–21 gestational weeks, by which time major events in sensory neurogenesis (NCC migration and DRG condensation) have occurred (Marmigere and Ernfors 2007; Montelius et al. 2007). The major subdivisions of DRG neuronal populations, large-diameter RT97+ neurons and small-diameter TrkA+ nociceptors, are present in DRG xenografts. DRG neurons express human-specific neuronal cell markers with the expected subcellular localization patterns such as neural cell adhesion molecule (NCAM), which is distributed along the cell surfaces of neurons and satellite cells, herpes viral entry mediator (HVEM/HveA), and synaptophysin, a major synaptic vesicle protein. DRG xenografts can be maintained in SCID mice for an extended period of time (>5 months).

VZV infection of DRG xenografts

At 4 weeks after transplantation, DRG xenografts are ready for infection experiments (Zerboni et al. 2005). DRG xenografts are inoculated by direct injection of VZV-infected fibroblasts using a 30-gauge needle. The needle is inserted through the kidney capsule, and a small volume (10 μl, ~1,000 PFU, ~3 × 104 cells) is slowly injected into the DRG xenograft. The needle is held in place for 15 s to prevent leakage from the needle track. The incision is closed, and mice are monitored until they awaken.

At specific timepoints after infection, mice are euthanized and the DRG xenograft is removed for analysis. Methods for assessment of VZV-infected DRG xenografts have been described in detail (Oliver et al. 2008; Reichelt et al. 2008; Zerboni et al. 2005, 2007, 2010, 2011a). For histological assessment, DRG are immersion fixed in 4% paraformaldehyde and paraffin-embedded (Zerboni et al. 2005). Serial sections (10 μm) are made through the entire implant, and every 20th section is stained with hematoxylin and eosin to identify regions of interest for further analysis by immunostaining or in situ hybridization. Quantitative DNA/RNA real-time polymerase chain reaction (PCR) is performed using the 5′-exonuclease method. For imaging experiments, bioluminescence is measured at 3–5-day intervals following intraperitoneal injection with d-luciferin. Mice are imaged using an IVIS Spectra imaging system (Xenogen), and bioluminescence (photons per meter per second squared) is calculated using Living Image Software (Caliper Life Sciences).

VZV infection of DRG xenografts

Acute VZV replication in DRG xenografts

We have used VZV–luciferase reporter viruses to sequentially assess viral replication within the DRG xenograft in living animals. In the experiment shown in Fig. 2a, b and c, DRG xenografts were infected with a recombinant VZV that expresses firefly luciferase driven by the immediate early regulatory gene ORF62 promoter, as a marker for lytic replication (Jones et al. 2006). As expected for a human restricted virus, the bioluminescent signal was restricted to the region of the DRG xenograft (Fig. 2a). As a negative control, SCID DRG mice were infected in parallel with a virus containing a promoterless firefly luciferase cassette; no signal was detected (Fig. 2b). As shown in Fig. 2c, bioluminescent values rose rapidly in the first 14 days following VZV–luciferase inoculation and then slowly declined reaching levels equivalent to measurements in mice infected with non-luciferase-expressing VZV by day 33. Live animal imaging using a recombinant VZV that expresses firefly luciferase as a fusion with the immediate early protein 63 (IE63) demonstrated a similar pattern (Oliver et al. 2008).

Fig. 2
figure 2

a–c The kinetics of VZV DRG infection can be monitored using VZV-expressing luciferase. a SCID DRG mouse infected with VZV–luciferase at 14 days after infection; the bioluminescent signal is restricted to the region of the left kidney. b SCID DRG mouse infected with VZV contains a promoterless luciferase cassette. c Course of VZV DRG infection using VZV expressing luciferase under control of the IE62 promoter. The dotted line indicates average bioluminescence in the promoterless control virus. d VZV genome copies in rOka, rOka gEΔCys, and ΔgI viruses at 14, 28, and 56 days after infection. e Quantitative real-time PCR for VZV mRNAs corresponding to ORFs 31 (gB), 62, and 63 at 56 days after infection. Circles represent individual DRG. f Standard PCR of DNA extracted from SCID xenografts using primers to amplify a 239-bp fragment of VZV ORF9a (top) and a 187-bp fragment of cellular β-globin (bottom). Lane 1, VZV-infected SCID mouse skin xenograft (day 21 post-infection); lane 2, uninfected skin xenograft; lane 3, human fetal DRG (prior to implantation); lane 4, VZV-infected DRG at day 14 after infection and day 56 (lane 5) after infection. g Immunoblot of DRG extracts using (top blot) human polyclonal anti-VZV antibody; lane 1: uninfected DRG, lanes 2–4: rOka-infected DRG, day 14; lanes 5–7: rOka-infected DRG, day 56. Immunoblot of DRG extracts (bottom two blots) using human polyclonal anti-VZV antibody (top) or rabbit polyclonal anti-IE63 IgG (bottom); lane 1: uninfected DRG; lanes 2–5: rOka-infected DRG, day 70; lanes 6–8: rOkaΔgI-infected DRG, day 70. Some panels in this composite image were adapted from panels that appeared in previous publications (see references): Zerboni et al. 2005, 2007, 2011a

Full size image

In a separate set of experiments, we demonstrated recovery of infectious virus from homogenized VZV-infected DRG at 14 days after infection (~70% of DRG), only rarely at day 28 (~12%), but not at day 56 and beyond (0%) (Zerboni et al. 2005, 2007). Taken together, these data indicate that the first 14-day interval following VZV ganglionic infection is characterized by a robust replicative phase within the infected ganglion that resolves, i.e., no infectious virus production, within 8 weeks post-inoculation.

VZV cytopathic effect, satellite cell infection, and polykaryon formation in DRG xenografts

Histological analysis of VZV-infected DRG at 14 days after infection reveals that most neurons express VZV proteins (Fig. 1d, thin arrow). However, neurons are observed that do not express VZV proteins (Fig. 1d, thick arrow). Within neurons that express VZV proteins, significant cytopathic changes are observed (Fig. 1c) (Zerboni et al. 2005). Evaluation of acutely infected DRG by ultrathin sections examined by electron microscopy (EM) revealed that satellite cells as well as neurons participate in virion production (Reichelt et al. 2008). Cytoplasmic and intranuclear viral capsids were observed within satellite cells surrounding some infected neurons (Reichelt et al. 2008). In a portion (~50%) of those VZV-infected neuron–satellite cell complexes, neuronal cell membranes had reorganized and fused with the membranes of the encapsulating satellite cells, forming multinuclear structures that resemble polykaryons that are characteristic of VZV skin infection (Reichelt et al. 2008). By confocal immunofluorescence, using the neuronal cell marker NCAM, examples of satellite cell nuclei with no apparent membrane separating them from the neuronal cell body are readily observed (Fig. 1g) (Zerboni et al. 2011a). The capacity for satellite cell infection and polykaryon formation may enhance VZV spread by providing a mechanism to amplify infectious virus progeny; however, in that satellite glial cells are critical regulators of neuronal homeostasis, disruption of the boundaries between neurons and satellite cells is likely to result in neuronal cell death. The consequences of VZV satellite cell infection and polykaryon formation may explain the increased severity of the herpes zoster rash compared with HSV reactivation. These virulence features may also contribute to neuropathic consequences of VZV reactivation, such as post-herpetic neuralgia.

Characteristics of VZV latency in DRG xenografts

As shown in Fig. 1d, at the peak of VZV replication (14 days after infection), neurons are observed that do not express VZV proteins, suggesting that some neurons resist VZV replication. Histological analysis of VZV-infected DRG at 56 days after infection reveals that neurons are still maintained in DRG xenografts (Fig. 1e). At the tissue level, satellite cell microproliferations indicate regions of neuronal cell loss (Fig. 1f, dashed arrow). However, clusters of neurons persist that express neural-specific cell markers (not shown) and do not express VZV proteins (Fig. 1e). The persistence of neurons in DRG xenografts several weeks after VZV DRG inoculation, in the absence of infectious virus production, suggests that VZV ganglionic infection may be intrinsically self-limiting. The resolution of lytic replication within the DRG xenograft occurs without any requirement for adaptive immune responses, which SCID mice lack.

Detection of VZV DNA by PCR (Fig. 1f) and quantitative real-time PCR (Fig. 1d) within DRG neurons that persist following lytic replication indicates that those neurons that were not permissive for lytic infection harbor latent genomes. At 14 days after VZV DRG infection, VZV genome copies are typically detected in the range of 108–1010 copies per 100,000 cells; by day 56, VZV genome copy numbers are significantly reduced but are consistently detected above background values in uninfected DRG, at 106–108 copies per 100,000 cells (Fig. 2d) (Zerboni et al. 2005, 2007, 2011a). In latently infected human cadaver ganglia, VZV DNA persists in 1.0–6.9% of neurons, and VZV genome copies have been widely estimated at 9–6,000 copies per 100,000 cells (Pevenstein et al. 1999; Wang et al. 2005). Our estimates in neurons that persist in DRG after cessation of VZV replication are higher, but this is expected as autopsied human cadaver ganglia are usually examined many decades after primary VZV infection.

Unlike HSV, VZV latency is not characterized by transcription of a region associated with latency, i.e., latency-associated transcripts (Cohen et al. 2007). In analyses of the VZV transcriptome in cadaver ganglia from individuals who died without evidence of herpes zoster, transcripts corresponding to VZV genes of all kinetic classes have been detected; RNAs corresponding to ORFs 63, 62, 21, and 29 are the most consistently reported (Cohrs et al. 1996, 2000; Nagel et al. 2011). The consistent detection of ORF63 mRNA suggests that IE63 transcription may be a characteristic of VZV latency (Cohrs et al. 2000). In an analysis of the presence of VZV mRNA in SCID DRG xenografts at day 56 after infection, transcripts corresponding to IE63 were most abundant with some IE62 transcripts, but no mRNA corresponding to the late viral glycoprotein gB were detected (Fig. 2e) (Zerboni et al. 2005). These quantitative real-time RNA PCR data demonstrate that, similar to observations in latently infected human cadaver ganglia, VZV genomes that persist in SCID DRG xenografts exhibit limited viral gene transcription.

Our experiments using the SCID DRG xenograft model indicate that VZV protein synthesis is not a characteristic of VZV latency in human neurons. In VZV-infected SCID DRG xenografts at 56 days after infection, VZV proteins are not detected by immunostaining (Fig. 1e), or when DRG cell lysates are examined by immunoblot (Fig. 2g) (Zerboni et al. 2007). Analyses of VZV protein synthesis during neuronal latency in human cadaver ganglia have been discordant, and all have relied exclusively on immunostaining methods. Several studies have reported abundant expression of IE63 and other “latency-associated proteins”, such as IE62 and proteins corresponding to ORFs 4, 21, 29, and 66 (Grinfeld and Kennedy 2004; Hufner et al. 2006; Lungu et al. 1998; Theil et al. 2003; Cohrs et al. 1996); whereas others (Mahalingam et al. 1996; Kennedy et al. 1998, 1999) have described IE63 protein expression in cadaver DRG as rare.

We conducted an extensive examination of cadaver ganglia and determined that IE63 protein expression in latently infected sensory neurons is an extremely rare occurrence (Zerboni et al. 2011a). In those rare IE63-positive neurons, co-expression of VZV glycoprotein E suggests an abortive or subclinical reactivation event, rather than bona fide latency (Zerboni et al. 2011a). In further support of only rare expression of VZV proteins during latency, we recently identified an immunostaining artifact that occurs in sensory neurons from individuals with blood type A (Zerboni et al. 2011b). This staining artifact may explain apparent positive staining attributed to VZV rabbit antisera and some ascites-derived anti-VZV monoclonal antibodies that contain endogenous anti-A antibodies (Zerboni et al. 2011b).

Summary of results using intact recombinant VZV

Our investigations using intact recombinant VZV and VZV–luciferase reporter viruses have demonstrated that initial events in VZV infection of SCID DRG xenografts are characterized by efficient spread in neurons, enabled by satellite cell infection and polykaryon formation, resulting in robust viral replication and release of infectious virus. In neurons that persist following this acute replicative phase, VZV genomes are present at low frequency with limited gene transcription and no protein synthesis, in a state that resembles latency in the natural human host. In the next part of this review, we will discuss our progress defining determinants of VZV neurovirulence using recombinant viruses with targeted mutations in VZV glycoprotein E and gI.

Contributions of VZV gE and gI to neurovirulence

VZV gE and gI are type I membrane proteins encoded by open reading frames 68 and 67, respectively. Both VZV gE and gI share features with orthologous herpesviral species, such as the formation of gE/gI heterodimers that participate in cell–cell spread (Cohen et al. 2007). HSV and pseudorabies virus (PRV) gE and gI proteins have been shown to participate in anterograde axonal transport and neuron-to-neuron spread in neuronal cultures and small animal models (Ch’ng and Enquist 2005; Dingwell et al. 1995; Polcicova et al. 2005; Saldanha et al. 2000). The cytoplasmic domain of PRV gI also contributes to neurovirulence in a manner that is independent of gE/gI interaction (Card et al. 1992; Tirabassi and Enquist 2000). We examined the capacity for recombinant Oka viruses containing deletions or mutations in gE and gI, or in gI promoter regulatory domains, to replicate in SCID DRG xenografts (Zerboni et al. 2007, 2011a). Acute replication and persistence in a latent state were assessed by quantitative real-time DNA PCR, infectious virus release assay, and evaluation of tissue sections for cytopathic effect, satellite cell tropism, and polykaryon formation (Zerboni et al. 2007, 2011a).

VZV glycoprotein E

A portion of the VZV gE N-terminal ectodomain is not conserved in other alphaherpesviruses, and this portion contains regions that are critical for replication in skin and T cell (thy/liv) xenografts in SCID mice (Berarducci et al. 2006, Berarducci et al. 2010). We examined the requirement for a portion of this region (amino acids 27–90) for VZV neuronal infection. The recombinant virus gEΔ27-90 lacks amino acids 27–90, and this abolishes gE interaction with insulin-degrading enzyme, a putative VZV entry receptor (Berarducci et al. 2010; Li et al. 2006, 2007). Although gEΔ27-90 exhibited decreased cell–cell spread in skin xenografts, it was not impaired in DRG xenografts (Berarducci et al. 2010; Zerboni et al. 2011a). Compared with recombinant Oka, the gEΔ27-90 mutant had equivalent VZV genome copies at day 14 and day 56, infectious virus release was similar, and there were no significant differences in spread, satellite cell infection, or polykaryon formation. Mutation of two endodomain motifs, the AYRV motif that mediates gE trafficking to the trans-Golgi network and a SSTT acidic cluster motif, had little or no effect on DRG infection or persistence (Moffat et al. 2004; Zerboni et al. 2011a; Zhu et al. 1996).

Mutation of the cysteine-rich region in the VZV gE ectodomain that abolishes interaction between VZV gE and gI had a significant effect on DRG infection (Zerboni et al. 2011a). The gEΔCys mutant was significantly impaired at early times after infection as demonstrated by reduced cytopathic effect and spread, indicating that gE/gI interaction is critical for efficient neuron–neuron transmission in DRG (Zerboni et al. 2011a). Unlike the expected pattern of decreasing VZV genome copies over time, VZV genomes increased between days 14 and 28 after infection (Fig. 2d). As shown in Fig. 1h, IE63 protein (green) was prominently expressed at 44 days after infection, which was equivalent to rOka-infected DRG at day 14. Dingwell and colleagues noted a similar requirement for HSV gE/gI interaction for spread within synaptically linked neurons (Dingwell et al. 1994, 1995).

Although the capacity to infect satellite cells was intact, the distribution of NCAM on neuronal cell surfaces demonstrated that the gEΔCys mutant had a reduced capacity for polykaryon formation compared with intact rOka (Fig. 1h, short arrow). Deficiencies in cytopathic effect, neuron–neuron transmission, and polykaryon formation would be expected to enhance neuronal survival; however, as infection progressed, we observed a marked fibrosis and microsatellite cell proliferations that indicate areas of neuronal cell loss. By day 56, large areas of gEΔCys-infected DRG xenografts contained acellular fields. These unexpected pathological changes suggest that mislocalization of gE that occurs in the absence of gE/gI interaction may have toxic effects in ganglionic cells. Alternatively, the slow spread of the gEΔCys mutant in DRG xenografts may have overstimulated cellular pathways that mediate the fibrotic response to neuronal damage.

VZV glycoprotein I

Mutation of gI promoter elements through which cellular transcription factors Sp1 and USF bind had no effect on DRG infection, although replication of both rOka-gI-Sp1 and rOka-gI-Sp1/USF was diminished in skin and T cell xenografts (Ito et al. 2003; Zerboni et al. 2007). Complete deletion of gI had a dramatic effect on spread of VZV within DRG xenografts. Rather than the typical short replicative phase characterized by high VZV genome copies and infectious virus production that slowly transitions to long-term persistence by day 56 (i.e., latency), infection with the recombinant virus rOkaΔgI resulted in an extremely prolonged replicative phase with slowly increasing VZV genomes over an 8-week interval (Fig. 2d) (Zerboni et al. 2007). While not strictly required for replication, histopathic effect at day 70 was minimal with very few cells exhibiting cytopathology, although infectious virus was recovered from minced tissues and VZV proteins were detected by immunoblot (Fig. 2e). In histological sections, uninfected neurons were observed surrounded by VZV-infected satellite cells (Fig. 1i), which suggests that gI may be a critical component of VZV entry into sensory neurons. Similar to the gEΔCys mutant, the normal membrane localization of gE was impaired, and instead gE was only detected within a few foci within the neuronal cytoplasm (Zerboni et al. 2007). However, we did not observe the severe pathological changes that characterized infection with the gEΔCys mutant. Instead, unusual Golgi stacks and disorganized rough ER containing gE were observed by immuno-EM staining of ultrathin sections (Zerboni et al. 2007). Polykaryon formation was more severely effected than with the gEΔCys mutant. Taken together, these results indicate that VZV entry into sensory neurons may require gI and the capacity for polykaryon formation is a pathogenic mechanism that greatly facilitates neuron–neuron spread in DRG xenografts.

Conclusion

Many questions about VZV neuropathogenesis remain unanswered, in particular, concerning the state of VZV during neuronal latency. We have made progress in defining pathogenic features and molecular requirements for VZV neuronal infection that contribute to clinical manifestations of varicella and herpes zoster. We hope that further investigation using the SCID DRG xenograft model will contribute to our understanding of VZV molecular virology and pathogenesis that may inform and improve future approaches to treatment and prevention of VZV-related disease.