Early Transcriptomic Response of Human Iris Stromal Cells During Herpes Simplex Virus Entry Reveals Interplay Between Cell Glycocalyx and Viral Exploitation

TL;DR

This study explores how human iris cells respond early during herpes simplex virus infection, revealing key inflammatory pathways and potential targets for treatment.

Contribution

The study identifies early transcriptomic changes in human iris stromal cells infected with HSV-1, highlighting conserved responses and potential therapeutic targets.

Findings

HSV-1 rapidly activates IL-17, TNFα, MAPK, and NF-κB pathways in human iris stromal cells.

Later stages show upregulation of epithelial–mesenchymal transition and G2/M checkpoint pathways.

Modulation of HS3ST enzymes and loss of heparan sulfate and syndecans were observed during infection.

Abstract

Herpes simplex virus type 1 (HSV-1) initiates infection through sequential interactions with host receptors, yet the early transcriptional responses driving HSV-mediated iritis remain poorly understood. Given the clinical burden of HSV-induced anterior uveitis and the lack of targeted therapies, we sought to define the initial host response to infection. We performed temporal transcriptomic profiling of primary human iris stromal (HIS) cells at 1, 3, and 6 h post-infection. HSV-1 triggered rapid and extensive gene expression changes, with early activation of IL-17, TNFα, MAPK, and NF-κB signaling pathways, all associated with inflammation and stress responses. At later time points, pathways related to epithelial–mesenchymal transition and the G2/M checkpoint were upregulated, alongside sustained inflammatory signaling, suggesting a balance between stromal integrity and stress adaptation. Imaging studies, together with transcriptomic data, revealed modulation of HS3ST enzymes and a corresponding loss of heparan sulfate and syndecans. These transcriptional dynamics mirror those observed in HSV-1-induced keratitis, indicating a conserved ocular response across cell types. By mapping these early events, this study identifies potential molecular targets for therapies aimed at mitigating inflammation during HSV-induced iritis.

Figures

Full text

1. Introduction

Herpes simplex virus type 1 (HSV-1) is a ubiquitous human pathogen responsible for a wide spectrum of diseases involving the oral mucosa, genital tract, central nervous system, and ocular tissues [1,2]. Infection may occur as a primary event or through periodic reactivation from latency established within sensory ganglia [3]. HSV-1 entry is a complex, multistep process initiated by virion attachment to cell-surface heparan sulfate (HS) proteoglycans, followed by fusion of the viral envelope with the host cell plasma membrane [4,5]. Current models indicate that viral entry requires the concerted action of four essential viral glycoproteins, gB, gD, gH, and gL [6,7], in coordination with at least one cellular receptor specific for gD [8,9]. Established gD receptors include herpesvirus entry mediator (HVEM), a member of the tumor necrosis factor receptor (TNFR) superfamily [10]; nectin-1, an immunoglobulin superfamily protein [11]; and 3-O-sulfated heparan sulfate (3-OS HS) [12]. The latter is synthesized by the enzymatic activity of the D-glucosaminyl 3-O-sulfotransferase (3-OST) family, among which all isoforms except 3-OST-1 can facilitate HSV-1 entry [12] and promote virus-induced cell-to-cell fusion [13]. Following corneal infection, HSV-1 undergoes retrograde axonal transport along the ophthalmic branch (V1) of the trigeminal nerve to establish lifelong latency within the trigeminal ganglion [14]. Upon reactivation, the virus travels anterogradely to the cornea, eliciting localized inflammatory responses. Ocular HSV-1 infection primarily targets the cornea, leading to herpetic keratitis, the most common infectious cause of blindness in developed nations [15,16,17]. In addition to keratitis, HSV-1 can cause acute uveitis with or without corneal involvement, characterized by inflammation of the iris stroma and pigmented epithelium, sectoral iris atrophy, and central keratic precipitates, frequently accompanied by elevated intraocular pressure (Figure 1A–C) [18]. If untreated, HSV uveitis can result in secondary glaucoma, cataract formation, corneal perforation, and retinal detachment [18,19,20,21,22]. The condition typically presents unilaterally, more commonly in middle-aged women, and may occur with or without concurrent corneal involvement [18,19]. The pathogenesis of HSV-1 induced uveitis is multifactorial, involving direct viral replication within the iris, ischemic vasculitis, infiltration of lymphocytes into the iris stroma and intraocular nerves, and immune-mediated tissue injury driven by recurrent infection [18,19,20]. To mechanistically dissect these processes, we established primary human iris stromal (HIS) cell cultures as a physiologically relevant in vitro model for studying host susceptibility and viral receptor utilization [23,24,25,26]. HIS cells maintain donor-specific HLA haplotypes and native extracellular matrix (ECM) organization, exhibit innate immune competence through pattern recognition receptors such as Toll-like receptors, and secrete proinflammatory mediators—including IL-6, IL-8, MCP-1, and RANTES following HSV-1 exposure [23]. Their genetic and phenotypic stability offers superior translational fidelity relative to animal models, making them a valuable platform for evaluating antiviral efficacy, inflammatory signaling modulation, and cytotoxicity in a human-relevant context [24].

The aim of this study was to perform temporal transcriptomic profiling of HSV-1–infected HIS cells compared to mock infected cells with a goal of identifying potential therapeutic targets for the development of antiviral and anti-inflammatory interventions capable of protecting ocular structures in the face of HSV-1 infection.

2. Results

2.1. Establishment of Primary HIS Cell Cultures and HSV-1 Infection for Transcriptomic Analysis

To investigate HSV-1 induced transcriptomic responses, primary human iris stromal (HIS) cells were isolated and cultured from healthy donor tissues. The purity of HIS cultures was confirmed by flow cytometry and immunofluorescence using the fibroblast marker vimentin (Figure 1D,E). To assess productive viral entry, HIS cells were exposed to the reporter β-galactosidase HSV-1 gL86 virus in a dose-dependent manner. As shown in Figure 2, HIS cells infected with the reporter virus exhibited a clear dosage-dependent response, as quantitively measured by colorimetric ONPG (Figure 2, panel A) and visually by X-gal assays (Figure 2, panel B). Subsequently, HIS cells were challenged with replication-competent HSV-1 (KOS 804 strain) at a multiplicity of infection (MOI) of 2.0 in a time-dependent manner, confirming productive infection via lytic gene expression with viral transcriptional activator protein (VP16) (Figure 2, panel C). Because VP16 can originate from both incoming tegument-associated protein and early de novo viral gene expression, protein-based detection alone cannot distinguish between these sources; accordingly, VP16 detection was interpreted as an indicator of early HSV-1 infection rather than definitive evidence of viral replication. Three biologically independent replicates were collected at 1, 3, and 6 h post-infection (hpi), alongside time-matched mock-infected controls, for RNA sequencing. Transcriptomic responses were analyzed over time to delineate early and late host responses. Principal component analysis (PCA) of normalized counts using DESeq2 confirmed high sample quality, with clear clustering of infected versus mock-infected samples at each time point (Supplementary Figure S1), indicating robust and distinct transcriptional changes induced by HSV-1 infection.

2.2. HSV-1 Infection Induces Progressive Transcriptional Changes in HIS Cells

Prolonged HSV-1 exposure elicited increasingly extensive transcriptional remodeling in HIS cells. Principal component analysis (PCA) demonstrated clear segregation between mock and HSV-1–infected HIS cells (Supplementary Figure S1). Transcriptomic analyses revealed that 1 h post-infection, 4074 transcripts were differentially expressed (1915 upregulated, 2159 downregulated; FDR < 0.05; Figure 3A), of which 32 exhibited ≥2-fold changes relative to time-matched mock-infected controls (Supplementary Table S1). By 3 h post-infection, the number of differentially expressed transcripts increased substantially to 9525 (5018 upregulated, 4507 downregulated), with 790 transcripts demonstrating ≥2-fold changes (Figure 3B). At 6 h post-infection, 11,714 transcripts were altered (6445 upregulated, 5269 downregulated), with 2708 transcripts showing ≥2-fold changes compared to controls (Figure 3C). Notably, 20 transcripts were differentially expressed (≥2-fold) across all three time points, while 625 transcripts that changed at 3 h remained significantly altered at 6 h (Figure 3D). The consistently altered transcripts included ANGPTL7, CCDC80, EGR3, FOS, FOSB, GALNT13, GDF5, IER3, JUNB, LRRC17, LRRN1, MAB21L1, MAMDC2, NF_K_BIA, NF_K_BIZ, NTNG1, NUAK2, TSPAN13, ZC3H12A, and ZFP36 (Figure 3E). Protein–protein interaction (PPI) analysis revealed a highly significant network connectivity among these proteins (p < 1.0 × 10^−16^). KEGG pathway enrichment analysis identified transcripts involved in osteoclast differentiation (FDR = 0.002), IL-17 signaling (FDR = 0.019), TNF signaling (FDR = 0.022), and hepatitis B-related pathways (FDR = 0.046) (Figure 3F), highlighting the activation of immune and inflammatory pathways in HIS cells during early HSV-1 infection.

2.3. Transcriptional Changes Following One Hour of HSV-1 Infection in HIS Cells

Analysis of the 32 transcripts exhibiting ≥2-fold differential expression at 1 h post-HSV-1 infection (Figure 4A) revealed protein–protein interactions (PPI) associated with canonical immune signaling pathways (Supplementary File S1), including TNF (FDR = 2.16 × 10^−9^), IL-17 (FDR = 9.64 × 10^−10^), and NF-κB signaling (FDR = 2.00 × 10^−6^) (Figure 4B). Gene set enrichment analysis (GSEA) demonstrated robust positive enrichment of the hallmark TNFα signaling via NF-κB gene set (NES = 2.44, padj = 0.005), indicating early activation of proinflammatory transcriptional programs. Upregulated NF-κB regulators included NF_K_BIA (2.7-fold, padj = 0.001), NF_K_BIZ (6.2-fold, padj = 4.57 × 10^−47^), and TNFAIP3 (4.2-fold, padj = 2.04 × 10^−51^), consistent with a tightly regulated negative feedback mechanism restraining NF-κB activity. Chemokines CXCL1 (4.2-fold, padj = 1.90 × 10^−16^), CXCL2 (9.5-fold, padj = 0.048), and CXCL3 (9.8-fold, padj = 5.06 × 10^−8^) were concomitantly upregulated, indicative of early chemotactic signaling known to facilitate neutrophil and monocyte recruitment. Immediate–early transcription factors FOS (7.5-fold), FOSB (3.9-fold), JUNB (3.3-fold), EGR1 (5.0-fold), EGR2 (4.1-fold), and EGR3 (5.6-fold) exhibited pronounced induction, underscoring their roles in orchestrating cellular survival, proliferation, and inflammatory responses (Figure 4C). Conversely, CCDC80, a fibronectin- and heparin-binding protein, was markedly downregulated (2.6-fold, padj = 2.94 × 10^−16^), suggesting early modulation of extracellular matrix interactions. Notably, JUNB, EGR1, EGR3, and PTGS2/COX-2 are established regulators of VEGF signaling, a key mediator of angiogenesis and cytoskeletal dynamics (Figure 4D). The early growth response (EGR) family comprises C2H2-type zinc-finger transcription factors [27], with EGR-1 central to signaling cascades regulating inflammation, proliferation, and apoptosis [28]. EGR-1 regulates HSV-1 ICP22 and ICP4 promoters [29] and is rapidly induced in SIRC and Vero cells during infection [30]. In murine models, EGR-1 exacerbates herpetic stromal keratitis, while its inhibition reduces corneal damage [31]. EGR-1 also promotes replication in bovine herpesvirus, HCMV, and KSHV but suppresses EBV lytic replication, indicating virus-specific functions [32]. Given its roles in ocular development, neovascularization, corneal injury, and heparanase regulation [33], EGR-1 may drive glycocalyx remodeling and heparan sulfate turnover during late HSV-1 infection, potentially enhancing viral spread in ocular tissues (Figure 4D,E) [34]. We also observed elevated ZC3H12A (MCPIP1/Regnase-1) transcripts (Figure 4C), a critical regulator of immune homeostasis that degrades pro-inflammatory mRNAs [35,36,37]. In the eye, ZC3H12A limits cytokines like IL-6 and IL-2, which are involved in keratitis and pterygium [38], and reduces inflammation in fungal keratitis by suppressing TNF-α, IL-6, and IL-1β [39]. In this context, MCPIP1 also enhances mTOR-mediated autophagy, facilitating pathogen clearance while concurrently dampening inflammation [39]. We also observed a robust upregulation of PTGS2 (COX-2) transcripts at both 1 and 3 h post-infection (Figure 4C), indicative of an early prostaglandin E_2_ (PGE_2_)-driven modulation of the inflammatory microenvironment that may facilitate HSV replication. Notably, COX-2 induction has been previously reported during HSV infection of the cornea, where it plays a pivotal role in orchestrating inflammatory and angiogenic cascades underlying herpetic stromal keratitis (HSK) pathogenesis [40]. Future investigations employing the human iris stroma (HIS) cell model will be instrumental in determining whether pharmacological inhibition of COX-2 can mitigate virus-induced inflammation and tissue damage. Moreover, the marked upregulation of CXCL1 transcripts (Figure 4C) further supports the hypothesis of virus-mediated iritis in our HIS cells, consistent with prior studies demonstrating the critical role of CXCL1 in neutrophil chemotaxis to the cornea and its contribution to recurrent HSK disease [41]. Previous studies have demonstrated that HSVs exploit MAPK–ERK signaling to regulate the cell cycle, block apoptosis, suppress innate immunity, and enhance viral replication [42]. In our time course of HSV-1 infection, MAPK pathway genes, including SPRY4, DUSP6, IER2, and IER3 (Figure 4C), were also induced by 1 h, indicative of cellular stress-response activation and signaling cascades that may be co-opted by HSV-1 to enhance replication. DUSP6, a negative regulator of ERK signaling [43], may serve as a checkpoint modulating HSV-1 induced transcriptional reprogramming. In the course of profiling cellular RNAs induced after HSV-1 infection, we also noted that a significant number of the upregulated NF-κB modulators NF_K_BIA, NF_K_BIZ, and TNFAIP3 (encoding the ubiquitin-editing enzyme A20) (Figure 4C) suggests a coordinated host response to restrain proinflammatory and antiviral signaling while simultaneously providing a potential mechanism for viral evasion of NF-κB-mediated apoptosis and interferon responses [44]. Collectively, these data delineate a highly orchestrated early transcriptional program in HIS cells upon HSV-1 infection, encompassing immune activation, chemokine-mediated leukocyte recruitment, MAPK pathway engagement, and dynamic regulation of host factors that influence viral replication and inflammatory responses.

2.4. Transcriptional Changes Following Three Hours of HSV-1 Infection in HIS Cells

At three hours post-HSV-1 infection, 26 of the 32 transcripts differentially expressed at 1 h remained similarly altered (Figure 5A). An additional 867 transcripts were differentially expressed (Supplementary File S2), with protein–protein interaction network analysis identifying enrichment of KEGG pathways associated with transcriptional mis-regulation in cancer (FDR = 0.0172). GSEA revealed negative enrichment of hallmark gene sets for epithelial–mesenchymal transition (NES = −2.049, padj = 0.102) (Figure 5B), G2M checkpoint (NES = −1.711, padj = 0.240), and inflammatory response (NES = −1.729, padj = 0.240) (Figure 5C). Several of the transcripts differentially expressed at 3 h have also been implicated in regulating the extracellular matrix (ECM). Specifically, HSV-1 infection caused a marked downregulation of MMP14 and DCN (decorin), indicating significant perturbations in ECM homeostasis. MMP14 (MT1-MMP) is a key membrane-bound metalloproteinase involved in ECM remodeling via type I collagen cleavage, pro-MMP2 activation, and basement membrane modification. It also influences cytoskeletal dynamics and angiogenesis through VEGF signaling [45]. Its downregulation suggests impaired ECM turnover, collagen buildup, and fibrosis, contributing to abnormal wound healing and iris scarring (Figure 5D). HSV-1 infection also reduced DCN, encoding decorin, a major corneal proteoglycan crucial for collagen fibrillogenesis, tissue integrity, and TGF-β regulation [46,47]. Decorin loss is linked to elevated intraocular pressure (IOP) in mice and observed in glaucomatous human tissue [48,49], hinting that its suppression during HSV may worsen IOP dysregulation. Conversely, HSV-1 rapidly upregulated CCL2 (MCP-1) within three hours (Figure 5E), driving monocyte recruitment to the cornea [50]. Stromal cells produce CCL2 and CCL5 to attract CD4^+^ T cells, intensifying inflammation [51]. Type I IFNs boost CCL2, promoting leukocyte infiltration [50]. Despite reduced macrophages in MCP-1 knockout mice, increased MIP-2 and MIP-1α worsened lesions and neovascularization, reflecting chemokine redundancy [52]. Similarly, HSV-1 induced CCL2 in human iris stroma cells, suggesting recruitment of T cells and inflammation in iritis (Figure 5E). These results underscore the coordinated chemokine–immune cell interactions driving HSV-1 ocular inflammation.

2.5. Transcriptional Changes Following Six Hours of HSV-1 Infection in HIS Cells

At six hours post-HSV-1 infection, 698 of the 893 transcripts differentially expressed at three hours remained differentially expressed, with an additional 2336 transcripts also exhibiting differential expression compared to mock-infected cells (Supplementary File S3; Figure 6A–C). Protein–protein interaction networks revealed KEGG pathway enrichment of transcripts associated with the pathway’s transcriptional misregulation in cancer (FDR = 0.0093), TGF-β signaling (FDR = 0.0014), Hippo signaling (FDR = 0.0093), MAPK signaling (FDR = 0.0093), and multiple other oncogenic and stress-response pathways (Figure 6A). These findings highlight HSV-1’s capacity to hijack host signaling networks. Transcriptional reprogramming in HIS cells underscores the central role of TGF-β, a pleiotropic cytokine involved in immune regulation, cell proliferation, and tissue repair [53], with evidence showing its upregulation increases HSV-1 susceptibility in murine models [54]. Canonical TGF-β signaling involves SMAD2/3/4 activation downstream of TβR-I and TβR-II [55], and it is constitutively expressed in the cornea to mediate cell migration and wound healing [56,57]. HSV-1 may exploit this pathway to promote replication and modulate ECM dynamics in iris stroma. At 6 h post infection, HIS cells upregulated JMJD6, a multifunctional nuclear demethylase (Supplementary Figure S2) implicated in chromatin and immune regulation [58,59], as well as RASD1 and RRAD, small GTPases tied to antiviral signaling (Supplementary Figure S2), whose knockdown enhances HSV-1 replication [60]. HIS cells thus serve dual roles supporting viral replication while activating antiviral responses. Notably, SGK1, a PI3K-regulated kinase involved in inflammation and cell survival [61,62,63,64,65], was time-dependently induced (Supplementary Figure S2) and shown to promote HSV-1 replication and apoptosis in corneal models via Wnt/β-catenin signaling [66]. Early infection also triggered upregulation of chromatin regulators BRD2 and CBX4 (Supplementary Figure S2) [67,68,69], pro-apoptotic BCL2L11 (Supplementary Figure S2), and anti-apoptotic PIM3 (Supplementary Figure S2) [70], reflecting a balanced interplay of host defenses and viral subversion. Collectively, HSV-1 reshapes host transcriptional and epigenetic landscapes to enhance replication while modulating immune and apoptotic pathways, revealing molecular targets for antiviral intervention.

2.6. HSV Infection Modulates Host Cell Glycocalyx

RNA sequencing analysis revealed that HSV-1 rapidly alters the fine structural organization of heparan sulfate (HS), with these changes appearing early during infection. We observed differential expressions of several key heparan sulfotransferases, including HS3ST1, HS3ST2, and HS3ST3A1 (Figure 7A). At 6 h post-infection, HS3ST1 and HS3ST3A1 were downregulated by 3.4-fold and 3.3-fold, respectively, whereas HS3ST2 was markedly upregulated (7.1-fold). These enzymes regulate the biosynthesis of 3-O-sulfated HS (3-OS-HS) on the cell surface (HS3ST1 [71,72,73]; HS3ST2 [74]; HS3ST3A1 [73,75,76]). Because most 3-OST-generated HS isoforms function as entry receptors for gD (except HS3ST1), modulation of these enzymes is expected to have a profound impact on HSV-1 entry. One possible model is that during initial viral attachment and entry, HS3ST3A1-derived 3-OS-HS is rapidly consumed, prompting compensatory upregulation of HS3ST2, another gD-compatible 3-OST to sustain infection. In this scenario, the loss of HS3ST3A1 transcripts may be offset by increased HS3ST2 expression, ensuring that HSV-1 continues to encounter a sufficient supply of gD-preferred 3-OS-HS to facilitate efficient entry. Because HS3ST3A1 and HS3ST2 also function as growth factors signaling hubs [77] the selective reduction of particular 3-O-sulfated HS species may trigger redistribution of FGF/FGFR interactions with HS, contributing to homeostatic responses during early virus invasion. We further hypothesized that these early transcriptional changes may reduce total HS abundance to protect target cells from subsequent rounds of infection. Immunofluorescence (IF) staining of HIS cells at 1, 3, 6, and 12 h post-infection supported this prediction. As shown in Figure 7B,C, HS levels were significantly reduced at all measured time points compared with mock-infected controls.

Consistent with the decline in total HS, transcriptomic data further showed that the HS core proteins syndecan-1 (SDC1) and syndecan-2 (SDC2) were also downregulated by 1.5-fold and 1.8-fold, respectively, at 6 h post-infection (Figure 8). IF staining further confirmed a robust decrease in syndecan-1 expression. Because syndecan-1 expression is influenced by membrane tension and actin cytoskeletal remodeling, these findings suggest that virus–host interactions may induce broader reorganization of the cortical actin network. Modulation of syndecans expression is consistent with observations from other microbial pathogens: altered syndecan-1 levels have been reported during infections with Pneumocystis jirovecii and Neisseria gonorrhoeae [78,79], and Epstein–Barr virus infection similarly downregulates syndecan-1 [80]. Collectively, our transcriptomic and immunofluorescence data demonstrate that HSV-1 extensively remodels the cellular glycocalyx early during infection. These coordinated changes likely optimize viral entry and prime the cellular environment for downstream events such as viral gene expression and replication.

3. Discussion

HSV-1 displays exceptional cellular tropism, efficiently infecting a broad spectrum of host cells, including multiple ocular cell types [34], underscoring its adaptability and clinical relevance. This broad host tropism is largely mediated by the pleotropic viral glycoprotein gD, which recognizes multiple host cell surface receptors [6,7]. These include broadly expressed molecules such as heparan sulfate (HS), integrins, epidermal growth factor receptor (EGFR), and fibronectin, as well as more specific entry mediators like nectin-1, herpesvirus entry mediator (HVEM), 3-O-sulfated heparan sulfate, and paired immunoglobulin-like type 2 receptor alpha (PILRα) [4,5,34]. Such receptor flexibility enables HSV-1 to exploit diverse cellular environments and may explain its capacity to establish infection in different cells and tissues. Moreover, the virus besides using direct host cell membrane fusion can also utilize several entry routes including phagocytic-like, and endocytic entry, by modulating host actin cytoskeletal dynamics [34,81]. This plasticity in entry mechanisms underscores the evolutionary advantage of HSV-1 and poses challenges for developing targeted antiviral strategies. Given the established role of HSV-1 as a major etiologic agent in anterior uveitis [17,18,19,20,21,22], our previous studies elucidated the molecular basis of viral entry into human iris stromal (HIS) cells, providing a physiologically relevant model to investigate ocular infection [23,24,25,26]. Notably, we also previously observed a robust pro-inflammatory response (IL-6, IL-8, MCP-1, RANTES, MIP-1β, GM-CSF) in HSV-1 infected HIS cells, offering deeper mechanistic insights into HSV-1–induced iritis [23]. The transcriptomic analyses presented herein delineate host cellular programs governing HSV-1 infection, immune activation, and cellular remodeling (Figure 9A). An extensive modulation of genes across multiple functional categories, including proviral and antiviral responses, apoptotic and anti-apoptotic pathways, inflammatory signaling, cellular remodeling, epigenetic regulation, and heparan sulfate synthesis/remodeling was noted (Figure 9B,C). Proviral factors such as EGR1, JMJD6, DUSP6, and SGK1 were upregulated, suggesting enhanced viral permissiveness, while antiviral effectors including RASD1 and RAAD were concurrently activated, indicating the initiation of host defense mechanisms. Interestingly, transcripts associated with apoptosis and anti-apoptosis were both modulated, with BCL2 and PIM3 promoting cell survival, while TNFSR10 supported cell death, reflecting a delicate balance between viral cytotoxicity and cellular resilience. Inflammatory pathways were prominently engaged, as shown by upregulation of COX-2 and MCP1, while anti-inflammatory regulation through ZC3H12A suggests feedback control of excessive immune activation. Additionally, epigenetic regulators CBX4 and BRD2 were induced, highlighting potential chromatin-mediated modulation of these host–virus interactions. Collectively, these findings underscore a dynamic interplay between viral exploitation and host defense orchestrated at multiple transcriptional and epigenetic levels. The observed upregulation of CXCL1, CXCL2, and CXCL3 transcripts during HSV-1 HIS cell infection suggests a robust activation of innate immune signaling pathways [82]. This chemokine-driven inflammatory response is consistent with previous reports linking CXCR2-mediated neutrophil trafficking to early antiviral defense. However, sustained or excessive expression of these chemokines may exacerbate cellular inflammation, contributing to corneal stromal damage and opacity commonly associated with chronic HSV-1-induced immunopathology. These findings underscore the dual role of chemokine-mediated inflammation in both host protection and disease progression during ocular HSV infection. Activation of specific genes can significantly modulate the structure and function of the glycocalyx, with implications for viral entry and tissue remodeling. The gel-forming mucin genes MUC2 and MUC5B encode major components of the mucinous glycocalyx [83] (Supplementary File S3; Figure 9C), which can act as either a physical barrier to pathogens or, paradoxically, as a decoy receptor facilitating viral attachment, including HSV-1 [84], depending on the context of glycosylation and mucin density. Concurrently, activation of C9 can lead to glycocalyx disruption under pathological conditions through the formation of the membrane attack complex (MAC), contributing to endothelial or epithelial damage [85]. Transcriptional activation of HOXA5 (Supplementary File S3), which binds the TAATGARAT motif in promoters of HSV immediate–early genes, may directly enhance viral replication by promoting early viral gene expression in host cells [86]. The activation of FGF22, a known mediator of signaling pathways that regulate synapse formation during post-injury remodeling [87], which may indirectly influence glycocalyx composition or function by altering local tissue microenvironments and extracellular matrix interactions, was also noticed. Similarly, we also observed that HSV-1 entry into HIS cells modulates FGF18 and FGF8 (Supplementary File S3; Figure 9C), promoting cytoskeletal remodeling and developmental signaling pathways, respectively, which may facilitate viral entry or replication [88]. In contrast, SULF1 and EXT2 (Supplementary File S3; Figure 7A), key regulators of HS modification and biosynthesis, were downregulated, potentially reducing HS availability on the cell surface and extracellular matrix [89,90]. This coordinated up- and downregulation underscores HSV-1’s strategic manipulation of HS-related pathways to optimize infection while reshaping the host cellular environment. Collectively, these gene activations illustrate a complex interplay where host glycocalyx dynamics, viral exploitation, and HIS cell remodeling converge. In our HIS-HSV entry model, we also noticed a selective transcriptional modulation of human 3-O-sulfotransferase (HS3ST) isoforms 1, 2, and 3 (Figure 7A), which certainly reflects a host–pathogen interface centered on the structural remodeling of heparan sulfate (HS) glycosaminoglycans [4,12]. The 3-OST family (encoded by HS3ST1–HS3ST6) catalyzes the transfer of sulfate groups to the 3-O position of glucosamine residues, generating distinct sulfation motifs that define the fine structure and biological specificity of HS chains [4]. Upregulation of specific HS3ST2 and HS3ST3 isoforms during HSV infection can enhance the biosynthesis of 3-O-sulfated HS domains, which in turn provides pool of high-affinity attachment and entry receptors for HSV-1 [12]. Such induction may represent a viral strategy to hijack the host sulfotransferase machinery, thereby optimizing cellular tropism and enhancing viral fusion and spread [9]. In contrast, downregulation of certain 3-OST transcripts could serve as an antiviral host defense, limiting the formation of permissive HS structures and attenuating viral entry. Beyond direct receptor modulation, changes in 3-OST expression have broader implications for cellular signaling, extracellular matrix (ECM) organization, and immune regulation, as 3-O-sulfated HS motifs influence the binding and bioavailability of cytokines, chemokines, and growth factors [91]. Our imaging data supported the loss of HS and major proteoglycan core protein syndecan-1, suggesting virus host cell interactions downregulate HS (Figure 7) and syndecan-1 (Figure 8) reprogramming glycocalyx and a protective mechanism for the virus from further invasion (Figure 7 and Figure 8). This transcriptional reprogramming likely occurs through cytokine-mediated or epigenetic mechanisms, reflecting a coordinated host response to infection-associated stress and inflammation. Taken together, these findings also suggest that differential regulation of the 3-OST gene network constitutes a critical molecular axis linking glycocalyx remodeling, innate immune signaling, and viral pathogenesis.

We observed a significant change in the transcript encoding decorin, a critical extracellular matrix proteoglycan involved in maintaining cellular integrity and orchestrating wound repair processes [47,48,49]. The pivotal role of decorin in modulating corneal wound healing and mitigating fibrotic scarring has been well established. In addition, decorin deficiency associated with elevated IOP [47] suggests that its HSV-induced suppression may worsen IOP dysregulation and ocular pathology. Our HIS cell model thus represents a robust platform to elucidate mechanisms regulating decorin expression and to assess the functional consequences of its modulation on wound healing dynamics, including potential attenuation of iritis. Overall, our findings highlight a significant role for IL-17 in the HIS HSV-1 infection model. It is well established that HSV-1 infection of the cornea activates both innate and adaptive immune responses, leading to a cascade of inflammatory cytokine production, including interleukin-17 (IL-17) [92]. IL-17 has also been shown to contribute to pathological angiogenesis in the cornea and is implicated in other ocular diseases, such as proliferative diabetic retinopathy and age-related macular degeneration [93,94]. In addition, the observed modulation of HS3ST genes is particularly noteworthy, as enzymatic modifications mediated by HS3ST generate specific heparan sulfate structures that function as HSV-1 glycoprotein D (gD) receptors and are critical for viral entry. These above findings highlight the intricate balance between viral replication and host defense mechanisms during HSV infection in HIS cell models. Identification of key differentially expressed genes and pathways provides critical insights into molecular nodes that HSV exploits to promote its survival and propagation, as well as host factors that restrict viral replication. Such comprehensive profiling offers a foundation for rational design of targeted interventions, including therapeutics that inhibit proviral pathways, enhance innate antiviral responses, or modulate epigenetic regulators to limit viral latency and reactivation.

Interestingly, several genes implicated in the pathophysiology of HSV-mediated keratitis also overlap with those involved in infection of the iris stroma. This observation is particularly significant, as HSV-induced keratitis can sometimes extend to the uvea/iris, presenting as keratouveitis [18,19,20]. Therefore, targeting viral infection in the iris may provide therapeutic benefits to the cornea, and conversely, strategies aimed at the cornea could help control viral activity in the iris. These overlapping molecular signatures highlight the interconnected nature of ocular HSV infection and suggest that interventions in one part of the eye may have protective effects in others. Overall, these results provide a roadmap for prioritizing molecular targets for the development of more effective HSV therapies and host-directed antiviral strategies, particularly in the context of ocular infection where multiple tissues are involved. Finally, we acknowledge certain limitations of our study. A more extensive dosage-dependent analysis in the presence and absence of acyclovir or anti-HSV sulfated glycans could have provided additional depth to the transcriptomic landscape. In addition, the use of iris epithelium versus iris stroma could have revealed intrinsic differences related to cellular permissiveness, antiviral signaling, and innate immune responses. Epithelial cells are generally highly permissive to HSV-1 replication and support robust lytic infection, whereas ocular-derived primary cells, including iris cells, may exhibit enhanced intrinsic antiviral defenses, altered interferon signaling, differential receptor expression, and cell-type-specific restriction factors that limit viral replication and spread. Nevertheless, the use of an MOI of 2.0 is well aligned with established HSV-1 transcriptomic studies aimed at achieving near-complete and synchronized infection, which is critical for robust host–virus transcriptional profiling. Comparable MOIs have been widely employed in literature. For example, Benxia Hu et al. used an MOI of 5.0 [69], while Hui-Lan Hu et al. employed an MOI of 1.5 per cell for single-cell transcriptomic analyses [95]. Moreover, even higher MOIs have been reported in independent studies, including those published in Nature Communications [60] and iScience [96], where HSV-1 infections were performed at an MOI of 10.0 at early time points.

4. Material and Methods

4.1. Primary Culture of Iris Stromal Cells

HIS cell cultures were prepared in accordance with institutional review board-approved protocols and were isolated from anonymously donated human eyes (provided by the Illinois Eye Bank, Chicago, IL, USA) via sterile dissection of the iris and pigmented epithelial layer and subsequent removal with a sterile cotton swab. The tissue was then digested with 0.2% type II collagenase (Sigma-Aldrich, St. Louis, MO, USA) in cell culture medium, MCDB-131 (Sigma-Aldrich, St. Louis, MO, USA) at 37 °C with gentle stirring for 20 to 30 min. Digested tissues were next centrifuged to remove tissue debris, and HIS cells were cultured in MCDB-131 containing 10% fetal bovine serum (FBS) and antibiotics.

4.2. Purity of Iris Stromal Cells

Human iris stromal cell cultures were cultured in 75 mm^2^ flasks to roughly 80% confluency before using for flow cytometry or immunofluorescent (IF) staining. IF staining was performed on cells that were seeded at 60,000 cells per well in a 24-well plate on size 12 circular glass coverslips. Cells were fixed in 2% paraformaldehyde for 30 min at 4 °C and then permeabilized with PBS containing 0.2% Tween-20 for 15 min at 37 °C. The cells were blocked with 5% donkey serum and then incubated with 0.4 ug of Vimentin antibody (Proteintech, Rosemont, IL, USA, cat# PTG10366-1-AP) or non-specific rabbit IgG (Vector Laboratories, Newark, CA, USA, cat# I-2000) for 30 min at 37 °C. Next, the cells were incubated for 30 min at 37 °C with 1.875 μg/mL of F(ab′)2 fragment donkey anti-rabbit IgG (H + L) conjugated to Alexa Fluor 594 (Jackson ImmunoResearch, West Grove, PA, USA, cat# 711-586-152). Coverslips were mounted with Vectashield Vibrance anti-fade mounting media with DAPI (Vector Laboratories, cat# H-1800-10) and IF images were collected using the 40× objective of a Nikon AXR confocal microscope system (Nikon Instruments Inc. Melville, NY, USA). Flow cytometry was performed using approximately 840,000 cells per sample of trypsin-detached cells. Cells were fixed in 2% paraformaldehyde for 30 min at 4 °C and then permeabilized with PBS containing 0.2% Tween-20 for 15 min at 37 °C. The cells were blocked with 5% donkey serum and then incubated with 0.4 ug of Vimentin antibody (Proteintech, Rosemont, IL, USA, cat# PTG10366-1-AP) or non-specific rabbit IgG (Vector Laboratories, Newark, CA, USA, cat# I-2000) for 30 min at 37 °C. Next, the cells were incubated for 30 min at 37 °C with 1.875 μg/mL of F(ab′)2 fragment donkey anti-rabbit IgG (H + L) conjugated to Alexa Fluor 488 (Jackson ImmunoResearch, West Grove, PA, USA, cat# 711-546-152). Cells were resuspended in PBS and read on a Beckman Coulter Cytoflex flow cytometer (Beckman Coulter, Brea, CA, USA).

4.3. Virus Titration

African green monkey kidney cells (Vero, ATCC, CCL-881) were grown to confluence overnight in 6-well plates. HSV-1 KOS (ATCC, VR-1493) dilutions were prepared in tubes by first adding 10 μL of virus stock to 1 mL serum-free media, making the 10^−2^ dilution. Subsequent dilutions were prepared by taking 100 μL of the previous dilution and adding it to 1 mL of serum-free media, continuing until the 10^−9^ dilution was reached. The growth media was aspirated from the plates, virus dilutions were added in duplicate, and the plates were incubated for 2 h with gentle rocking. After absorption, the wells were washed 3× with PBS to remove unbound virus particles. After washing, the cells were incubated for 48 h in 1.5 mL overlay media containing DMEM supplemented with 1% heat-inactivated FBS and 0.5% methyl cellulose. After 48 h, cells were washed 3× with PBS and fixed for 30 min in 4% paraformaldehyde. Cell monolayers were then stained with Giemsa (Sigma-Aldrich, St. Louis, MO, USA, cat# 48900) for 20 min, and plaques were counted at 10× magnification to determine plaque-forming units per milliliter (PFU/mL).

4.4. Western Blot Analysis

HIS cells from one confluent well of a six-well plate infected or mock-infected with HSV-1 KOS804 for 1, 3, or 6 h were lysed by adding RIPA lysis buffer supplemented with a protease inhibitor cocktail in an ice bath for 30 min. The lysates were centrifuged at 12,000× g for 10 min at 4 °C, and the supernatants containing total proteins were prepared. Then, 2× loading buffer was added, and 50 μg of total cell lysate was separated by 8% Tris-Glycine SDS-PAGE and transferred to 0.45 μm Nitrocellulose membranes (Bio-Rad Laboratories, Hercules, CA, USA) using established protocols. The membranes were blocked in 5% non-fat dry milk for 1 h and incubated with anti-VP16 (1-21) 1:200 (Santa Cruz, Dallas, TX, USA, Cat. No. sc-7545) overnight at 4 °C. The membranes were washed four times with TBS + 0.1% Tween-20. The membranes were incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (goat anti-mouse IgG Fc BP-HRP, Santa Cruz, Dallas, TX, Cat. No. sc-525416) and developed using enhanced chemiluminescence (ECL) substrate. The results were confirmed by at least three biological replicates.

4.5. Immunofluorescence

HIS cells were cultured on 12 mm coverslips and infected with 2 MOI of HSV-1 for 1, 3, 6, and 12 h. Following fixation, cell membranes were permeabilized using 1% Triton-X100 for five minutes and washed three times in PBS. Samples were then blocked using 1% bovine serum albumin (BSA, Sigma-Aldrich, St. Louis, MO, USA) in PBS with 0.2% Tween-20 (PBS-T). After blocking, coverslips were washed again and incubated with mouse anti-Heparan Sulfate (1:500 dilution, Amsbio, Cambridge, MA F58-10E4, catalog no: 370255-S), anti-CD138 (syndecan-1) monoclonal antibody (1:200 dilution, ThermoFisher, Waltham, MA, DL-101, catalog no: 14-1389-82), anti-syndecan-2 polyclonal antibody (1:125 dilution, ThermoFisher, Waltham, MA, USA, catalog no: 36-6200).

Coverslips were then washed with PBS-T three times and incubated with the appropriate Alexa Fluor 594 secondary antibodies (1:1000, Invitrogen, Carlsbad, CA, USA) for 1 h in 1% BSA in PBS-T. Samples were then washed and incubated with 300 nM DAPI (Sigma) for 15 min and mounted onto glass slides using Thermo Immu-Mount (New York, NY, USA). Samples were then imaged using an EVOS ^TM^ M5000 Imaging system (ThermoFisher Scientific, Waltham, MA, USA). Images were analyzed in FIJI. Images were pre-processed using a gaussian blur filter (0.2 μm) and rolling pin filter (5 μm). Mean fluorescent intensity measurements were taken of 300 × 300 μ regions of interest. Data was then normalized by nuclei identified using an Otsu threshold. Samples stained normally, but without primary antibody, were used as controls. Datasets were then normalized by 1 h mock control for visualization.

4.6. RNA Extraction

Total RNA was extracted from cells in Trizol using the Trizol method following manufacturer’s instructions (ThermoFisher Scientific, Waltham, MA, USA). Total RNA samples were quantified using Qubit 2.0 Fluorometer (Life Technologies, Carlsbad, CA, USA) and RNA integrity was checked using Agilent TapeStation 4200 (Agilent Technologies, Palo Alto, CA, USA).

4.7. Library Preparation with PolyA Selection and Illumina Sequencing

RNA sequencing libraries were prepared using the NEBNext Ultra II RNA Library Prep Kit for Illumina using manufacturer’s instructions (NEB, Ipswich, MA, USA). RNA fragmentation was performed in a mildly basic buffer (10 mM Tris-HCl, pH 8.0) according to the NEBNext Ultra II RNA Library Prep protocol. Briefly, mRNAs were initially enriched with Oligod(T) beads. Enriched mRNAs were fragmented for 15 min at 94 °C. The first strand and second strand cDNA were subsequently synthesized. cDNA fragments were end repaired and adenylated at 3’ends, and universal adapters were ligated to cDNA fragments, followed by index addition and library enrichment by PCR with limited cycles. The sequencing library was validated on the Agilent TapeStation (Agilent Technologies, Palo Alto, CA, USA), and quantified by using Qubit 2.0 Fluorometer (Invitrogen, Carlsbad, CA, USA) as well as by quantitative PCR (KAPA Biosystems, Wilmington, MA, USA). The sequencing libraries were multiplexed and clustered onto a flow cell on the Illumina NovaSeq instrument or equivalent according to manufacturer’s instructions. The samples were sequenced using a 2 × 150 bp Paired End (PE) configuration. Image analysis and base calling were performed by using NovaSeq Control Software (NCS v1.8.1; Azenta Life Sciences, Waltham, MA, USA). Raw sequence data (BCL files) generated from the Illumina NovaSeq were converted to FASTQ files and demultiplexed using Illumina bcl2fastq v2.20, allowing one mismatch for index sequence identification. Following demultiplexing, sequencing reads were subjected to quality control and adapter trimming using standard parameters. High-quality reads were aligned to the reference genome using a splice-aware aligner, and aligned reads were filtered to remove low-quality and ambiguously mapped reads. Gene-level read counts were generated from the aligned files using annotated gene models, and the resulting count matrices were used for downstream normalization and differential expression analysis.

4.8. Differential Expression, Gene Set Enrichment Analysis (GSEA), and Interaction Network Analysis

Count normalization and differential expression analysis was performed using DESeq2, version 3.22 [97] in R. Transcripts were considered differentially expressed when FDR was <0.05 and expression changed ≥2-fold in either direction in HSV-1 treated cells compared to vehicle-treated cells at the same time point. GSEA was performed using the R package fgsea [98] using Hallmark and Reactome databases downloaded from Molecular Signatures Database (MSigDB v2025.1Hs) [99]. Maps of interaction networks were constructed with high confidence (0.7) evidence of both functional and physical interactions between mapped proteins encoded by differentially expressed transcripts using STRING v12.0 (http://string-db.org). Functional KEGG pathway enrichments were determined for all time points, as well as for overlapping mapped proteins across all time points. Note that for the 3 h timepoint, only 606 out of 790 differentially expressed transcripts could be mapped to proteins. For the 6 h timepoint, only 2050 out of 2708 differentially expressed transcripts could be mapped to proteins.

5. Conclusions

This study was designed to define early host transcriptomic signatures during HSV-1 entry, focusing specifically on window when viral attachment and fusion occur. An MOI of 2 was determined to be optimal, achieving 100% infection by 6 h post-infection; transcriptomic profiling was therefore conducted at 1, 3, and 6 h. The transcriptomic analyses presented herein establish a comprehensive framework for characterizing host cellular responses that govern HSV-1 infection dynamics, immune activation, and structural remodeling (Figure 9). The relatively low levels of heparan sulfate (HS) expression detected at early time points (Figure 7) likely reflect the fact that HSV-1 entry primarily depends on pre-existing cell surface HS rather than immediate de novo synthesis. During the early stages of infection, host transcriptional responses are predominantly driven by innate immune activation and cellular stress signaling. In contrast, modulation of HS biosynthetic and modifying enzymes, including members of the HS3ST family, is more likely to occur at later stages as part of virus-induced cellular remodeling. Accordingly, future studies examining both dose- and time-dependent effects on HS and HS-related gene expression would help determine whether these changes represent secondary transcriptional responses rather than immediate entry-associated events. By delineating the mRNA signatures and molecular circuits associated with viral persistence and pathogenesis, these datasets will provide a foundation for rational target prioritization and therapeutic development. Integrating transcriptomic insights with functional and temporal validation will be essential to resolve the sequence of host–virus interactions and to identify regulatory nodes most susceptible to pharmacologic modulation. In this direction, targeting HSV-1 across multiple phases of its life cycle, including viral entry, post-entry processing, and early transcriptional activation represents a rational and potentially transformative therapeutic strategy [100]. Interventions at these upstream stages could prevent the downstream amplification of immune-mediated inflammation and tissue injury that culminate in irreversible damage of the iris with or without corneal involvement during ocular HSV infection. In parallel, therapeutic agents designed to inhibit viral attachment, modulate cytokine networks, preserve epithelial architecture, and enhance cellular repair mechanisms may confer both antiviral and cytoprotective benefits. Such dual-acting or multi-targeted approaches hold strong translational promises to bridge the existing gap in effective interventions for HSV-associated ocular disease. In this study, we utilized the laboratory-adapted HSV-1 KOS (804) strain rather than clinical isolates, as KOS is a gold-standard model extensively employed to dissect mechanisms of viral entry and cell-to-cell spread. Importantly, even with this laboratory strain, our analyses identified multiple transcriptional signatures that closely recapitulate those reported for clinical HSV-1 isolates, highlighting the translational relevance and robustness of our findings. Nevertheless, future studies incorporating genetically diverse clinical HSV-1 isolates, both in the presence and absence of the antiviral drug acyclovir, will be essential to further validate and extend these observations. Moreover, ongoing and future efforts will focus on defining key molecular effectors and their regulatory networks that drive HSV-1 pathogenesis, with a particular emphasis on screening high-affinity inhibitors targeting HSV-1 glycoprotein D (gD) and critical pro-inflammatory cytokines [100]. Collectively, these precision-based therapeutic strategies have the potential to redefine the treatment paradigm by concurrently limiting viral propagation, attenuating immunopathology, and preserving visual integrity in affected patients.