https://orcid.org/0009-0001-1698-9592
Figures
Abstract
Lumpy skin disease virus (LSDV) is a critical transboundary pathogen that causes devastating infections in cattle, buffalo, and other ruminants. The virus induces characteristic clinical manifestations, including cutaneous nodules, marked reduction in milk yield, and impaired production performance, leading to severe economic losses in the global livestock sector. Although LSDV exhibits remarkable multi-tissue tropism and persistent viral shedding in various organs, posing significant challenges for disease control, the molecular mechanisms underlying its tissue-specific adaptation remain poorly understood. Here, we established both bovine cell models and golden hamster models to elucidate the tissue-specific pathogenic mechanisms of LSDV, and further validated these findings in bovine kidney and mammary tissue samples to demonstrate their relevance in natural hosts. Our findings revealed that LSDV employs distinct cell death pathways in different tissues to facilitate host adaptation. In kidney tissue, the viral envelope protein ORF117 specifically interacts with host GAPDH, triggering its nuclear translocation and subsequent activation of the GAPDH-Siah1/p53 signaling cascade, culminating in Caspase-3-mediated apoptosis. Conversely, in mammary tissue, LSDV induces Caspase-8-dependent cleavage of Gasdermin C, promoting pyroptosis in mammary epithelial cells and substantial release of inflammatory cytokines IL-1β and IL-18. This study provides the first mechanistic insight into the molecular basis of LSDV’s tissue-specific activation of distinct cell death pathways, establishing a theoretical framework for developing targeted therapeutic interventions against lumpy skin disease.
Author summary
Lumpy skin disease virus (LSDV), a significant pathogen of cattle, causes severe economic losses in livestock industries worldwide. Although LSDV exhibits broad tissue tropism, the molecular mechanisms underlying its tissue-specific adaptation remain elusive. Our study revealed that LSDV orchestrates distinct cell death pathways in different tissues. Using both in vitro cell culture systems and in vivo hamster models, we demonstrated that LSDV triggers tissue-specific cell death mechanisms, and further validated these findings in bovine kidney and mammary tissue samples, confirming their relevance in natural hosts. In kidney cells, the viral protein ORF117 induces apoptosis through interaction with host GAPDH, promoting its nuclear translocation and subsequent activation of death signaling pathways. Conversely, in mammary tissue, LSDV activates pyroptosis via Gasdermin C-mediated membrane permeabilization, leading to inflammatory cytokine release. These findings uncovered a sophisticated viral strategy where LSDV employs immunologically silent cell death in kidney tissue while promoting inflammatory responses in mammary tissue. This newly discovered tissue-specific adaptation mechanism provides potential therapeutic targets for controlling this economically important livestock disease.
Citation: Wen Y, Wang T, Zhang S, Wang J, Li C, Liu C, et al. (2026) Cell death modulation dictates tissue-specific tropism of lumpy skin disease virus. PLoS Pathog 22(2): e1013982. https://doi.org/10.1371/journal.ppat.1013982
Editor: Pinghui Feng, University of Southern California, UNITED STATES OF AMERICA
Received: July 19, 2025; Accepted: February 6, 2026; Published: February 26, 2026
Copyright: © 2026 Wen et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The RNA‑seq data generated in this study have been deposited in the NCBI BioProject database under accession number PRJNA1423217. All other relevant data are within the manuscript and its Supporting Information files.
Funding: This work was supported by the National Key Research and Development Program of China under grant number 2023YFF0611504. Funding was provided to F.X. The funder’s website is https://www.most.gov.cn. The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Poxviruses represent significant zoonotic pathogens that draw global attention in both public health and livestock sectors due to their extensive host range and distinctive tissue tropism. These viruses are characterized by large double-stranded DNA genomes, sophisticated replication mechanisms, and remarkable capacity for cross-species transmission. These viruses can infect diverse animal hosts, including humans, typically causing systemic infections with multi-organ pathology [1–3]. Although smallpox virus was eradicated in the twentieth century [4], other poxvirus family members, notably monkeypox virus and goatpox virus, have been emerging and spreading rapidly in recent years. These viruses frequently cross species barriers, presenting escalating challenges for disease control and posing serious threats to global public health and livestock industries [5–7]. Understanding their pathogenic mechanisms and developing targeted intervention strategies have become increasingly urgent.
A distinctive feature of poxvirus infections is their pronounced tissue tropism: the ability to adapt to diverse host tissue microenvironments and efficiently replicate in multiple tissues, including skin, mucous membranes, and visceral organs [8–12]. For instance, in African squirrels, monkeypox virus induces pathological changes in multiple organs, including skin, lungs, and kidneys [13], whereas goatpox virus causes severe respiratory and visceral damage in goats [12]. This multi-tissue infectivity enhances viral pathogenicity and significantly complicates disease control strategies. Traditionally, tissue tropism has been attributed to the synergistic effects of multiple factors, including virus-host receptor interactions, viral immunomodulatory genes, and host physical and immune barriers [11,14–16]. However, emerging evidence suggests that the dynamic regulation of programmed cell death (PCD) pathways in host cells may be a critical determinant of tissue-specific viral colonization and dissemination [17–19].
PCD is a crucial host defense mechanism against viral infections, encompassing various forms including apoptosis, pyroptosis, and programmed necrosis [18,20]. Apoptosis, a low-inflammatory process, effectively constrains viral spread by controlling the elimination of infected cells, although it may inadvertently facilitate viral latency and persistent replication. In contrast, pyroptosis and programmed necrosis trigger the release of inflammatory mediators, which enhance immune responses to restrict viral dissemination. However, these inflammatory forms of cell death may paradoxically promote viral spread through tissue damage [21]. Poxviruses have evolved to encode multiple regulatory proteins that modulate host PCD pathways, enabling efficient viral infection and transmission [15,22,23]. Nevertheless, the molecular mechanisms underlying tissue-specific PCD regulation by poxviruses and their strategic selection of distinct PCD modes in different tissues remain incompletely understood.
Lumpy skin disease virus (LSDV), a significant transboundary pathogen within the Poxviridae family, has been emerging globally in recent years, inflicting substantial economic losses on the cattle industry [24,25]. LSDV infection manifests primarily as widespread cutaneous nodules, accompanied by reduced lactation, weight loss, and decreased productivity [26,27]. Like other poxviruses, LSDV exhibits remarkable tissue tropism, establishing infection in multiple organs including skin, lungs, liver, spleen, and kidneys, leading to significant organ damage [28]. However, the molecular mechanisms underlying LSDV’s tissue-specific modulation of programmed cell death patterns remain to be elucidated.
To address these knowledge gaps, this study investigated the molecular mechanisms underlying LSDV-induced tissue-specific cell death using in vivo and in vitro models. Systematic analysis revealed distinct regulatory mechanisms governing virus-host interactions across different tissues. The findings not only advance our understanding of poxvirus pathogenesis but also identify potential therapeutic targets for developing targeted interventions and control strategies against poxvirus infections.
Results
Significant differences in infection characteristics of LSDV in kidney and mammary tissue and cells
An experimental golden hamster infection model was established to investigate LSDV’s tissue-specific infection characteristics (Fig 1A). LSDV infection significantly suppressed body weight gain in infected animals (Fig 1B) and induced characteristic pox-like cutaneous lesions (red arrows; Fig 1C). Viral loads were analyzed using a newly developed quantitative PCR assay (S1 Fig) in various tissues at 14 days post-infection. Although LSDV was detected in all examined tissues, viral loads varied significantly among different organs. The highest viral titers were observed in dorsal skin, followed by kidney, mammary, and liver tissue, indicating distinct tissue tropism (Fig 1D).
(A) Schematic diagram of the golden hamster infection model. Created in BioRender. wen, y. (2025) https://BioRender.com/5sa93q9 (B) Body weight changes in LSDV-infected and mock-infected hamsters over 14 days post-infection (dpi). (C) Representative image demonstrating characteristic pox-like skin lesions (red arrows) in LSDV-infected hamsters. (D) Viral loads in different tissues at 14 dpi quantified by qPCR (n = 5). (E) Growth kinetics of LSDV in MDBK and MAC-T cells infected at MOI = 1. (F) Phase-contrast microscopy images of morphological changes in LSDV-infected MDBK and MAC-T cells at 24 hpi compared to mock-infected controls. Scale bars = 20 μm. (G) TEM images illustrating ultrastructural changes in LSDV-infected cells at 24 hpi. Scale bars = 500 nm. Statistical significance was determined using one-way ANOVA with multiple comparisons (*p < 0.05, **p < 0.01, ***p < 0.001). Error bars represent mean ± SD.
To elucidate the molecular mechanisms underlying LSDV’s tissue-specific infection patterns, viral infection characteristics were analyzed in two bovine cell lines: kidney-derived Madin-Darby bovine kidney (MDBK) cells and mammary-derived bovine mammary alveolar cells (MAC-T). After infection at a multiplicity of infection (MOI) of 1, viral growth kinetics revealed that LSDV initiated logarithmic growth at 4–8 hours post-infection (hpi) and reached plateau at 24–36 hpi. Notably, LSDV demonstrated significantly higher replication efficiency in MDBK cells compared to MAC-T cells, consistent with the tissue tropism patterns observed in the hamster model (Fig 1D-1E).
LSDV infection induced distinct morphological changes in these cell lines. MDBK cells exhibited marked shrinkage, aggregation, and volume reduction, whereas MAC-T cells showed characteristic swelling and membrane blebbing (Fig 1F). Transmission electron microscopy (TEM) revealed that whereas control cells maintained intact cytoplasmic structure and normal nuclear morphology, infected cells in both cell types displayed varying degrees of disrupted cytoplasmic architecture with multiple vacuoles in MAC-T and evident cellular disintegration in MDBK. Characteristic brick-shaped to oval viral particles with distinct envelope structures were visible in infected cells (Fig 1G). The morphological and ultrastructural observations demonstrate significant differences in LSDV infection patterns between kidney and mammary cells.
LSDV infection induces apoptosis in MDBK cells
To elucidate the molecular mechanisms underlying LSDV’s tissue-specific infection patterns, transcriptome analysis was first performed on LSDV-infected MDBK cells. RNA sequencing revealed significant activation of multiple cell death-related pathways, including apoptosis, p53 signaling, and TNF signaling pathways (Fig 2A). Considering that viruses often modulate host cell death pathways to optimize their replication and spread, it was hypothesized that LSDV potentially exploits apoptotic mechanisms to facilitate its infection in kidney cells.
(A) KEGG pathway enrichment analysis of differentially expressed genes in LSDV-infected MDBK cells. Pathways associated with apoptosis, including the p53 signaling pathway and TNF signaling pathway, are significantly enriched. Circle size corresponds to the number of genes, and color intensity indicates statistical significance (-log10(q-value)). (B) TUNEL assay demonstrated DNA fragmentation in MDBK cells infected with LSDV, staurosporine (positive control), or mock treatment. Green TUNEL-positive signals indicated LSDV-induced apoptotic DNA fragmentation. Scale bars = 20 µm. (C) TEM images of MDBK cells infected with LSDV or mock-treated. Scale bars = 500 nm. (D) Quantification of early apoptotic, late apoptotic, and necrotic cells from flow cytometry analysis of MDBK and MAC-T cells. (E) Flow cytometry analysis of Annexin V-FITC and PI staining in MDBK and MAC-T cells infected with LSDV (12, 24, 48, 72, and 96 hpi) or mock-treated. (F) Western blot analysis of Caspase-3 expression and activation (Cleaved caspase-3) in MDBK and MAC-T cells following LSDV infection (12, 24, 48, 72 and 96 hpi) or mock treatment (96 h). α-tubulin was used as loading control.
TUNEL assays revealed extensive DNA fragmentation in LSDV-infected MDBK cells, a hallmark of apoptotic cell death (Fig 2B). TEM further confirmed the apoptotic phenotype, with characteristic ultrastructural changes including nuclear envelope disruption and chromatin condensation and margination; mitochondrial degeneration with cristae loss and vacuolization; and membrane blebbing with formation of apoptotic bodies (Fig 2C). In contrast, mock-infected cells exhibited normal nuclear architecture with homogeneous chromatin distribution and intact mitochondrial morphology.
To determine whether this apoptotic response is cell-type specific, apoptosis levels were compared between MDBK and MAC-T cells using Annexin V-FITC/PI double staining and flow cytometry. LSDV infection significantly increased both early apoptotic (Annexin V + PI–) and late apoptotic (Annexin V + PI+) cell populations in MDBK cells, with peak effects observed between 72 and 96 hpi (Fig 2D-2E). In contrast, MAC-T cells exhibited notably lower apoptotic rates following LSDV infection, indicating that LSDV induces different apoptotic responses in kidney and mammary cells.
To further characterize the molecular basis of this cell-type specific apoptotic response, examination was conducted on the activation of Caspase-3, a key executioner protease in apoptotic signaling. Western blot analysis revealed time-dependent increases in cleaved Caspase-3, an established marker of apoptosis, in MDBK cells, while MAC-T cells showed no detectable Caspase-3 activation under the same experimental conditions (Fig 2F). These findings establish that LSDV specifically activates Caspase-3-dependent apoptotic signaling in kidney cells.
To verify that the observed apoptotic response in MDBK cells was independent of viral burden, viral RNA levels were quantified at 16 hpi for MDBK cells and 20 hpi for MAC-T cells, confirming comparable viral loads between the two cell types. Immunofluorescence analysis further demonstrated Cleaved Caspase-3 activation in LSDV-infected MDBK cells, while no such activation was detected in MAC-T cells under the same conditions (S2A Fig).
LSDV ORF117 interacts with GAPDH and induces apoptosis in MDBK cells
Viral envelope proteins not only mediate viral entry and replication but also modulate host cell death pathways and immune responses, thereby affecting viral spread and host defense [29–32]. To investigate the molecular mechanism of LSDV-induced apoptosis, all LSDV envelope protein genes (ORF024, ORF028, ORF046, ORF060, ORF064, among others) were cloned and expressed as GFP fusion proteins (Fig 3A). Using a Caspase-3/7 activity assay, ORF117 was identified as a key effector that significantly activated Caspase-3/7, showing markedly higher activity compared to other viral protein candidates (Fig 3B). Notably, several proteins including ORF024, ORF046, and ORF070 exhibited anti-apoptotic effects, highlighting the functional diversity among LSDV envelope proteins.
(A) Fluorescence microscopy images of Vero cells transfected with plasmids expressing GFP-tagged LSDV ORFs. Scale bars = 100 µm. (B) Caspase-3/7 activity assay in Vero cells transfected with plasmids expressing the indicated LSDV ORFs. (n = 3) (C) Fluorescence microscopy images of MDBK cells stably expressing ORF117, demonstrating expression of ORF117-associated GFP fluorescence. Scale bars = 100 µm. (D) Western blot analysis confirming LSDV ORF117 expression in MDBK cells stably expressing ORF117. α-tubulin was used as the loading control. (E) Caspase-3/7 activity assay in MDBK cells stably expressing ORF117. (F) SDS-PAGE and silver staining of GST pull-down assay using recombinant GST-tagged ORF117 protein. (G) Mass spectrometry analysis of proteins interacting with ORF117. (H) Co-IP assay in 293T cells co-transfected with FLAG-tagged GAPDH and Myc-tagged ORF117. (I) Co-IP assay in MDBK cells infected with LSDV or mock-treated. (J) Confocal microscopy images of 293T cells co-transfected with Flag-GAPDH and Myc-ORF117 or MDBK cells infected with LSDV. Line-scan plot profiles of fluorescence intensity demonstrate overlapping signals of GAPDH and ORF117. Scale bars = 10 µm. Statistical significance was determined using one-way ANOVA with multiple comparisons (*p < 0.05, **p < 0.01, ***p < 0.001). Error bars represent mean ± SD.
To confirm the role of ORF117 in apoptosis, a stable MDBK cell line expressing ORF117 (MDBK-ORF117) was established (Fig 3C). Western blot analysis revealed a specific protein band corresponding to ORF117 (~17 kDa) in MDBK-ORF117 cells but not in control cells (Fig 3D). Caspase-3/7 assays further showed that Caspase-3/7 activity in MDBK-ORF117 cells was significantly higher than in control cells after 48 hours of culture (Fig 3E). These results establish ORF117 as a potent inducer of apoptosis in MDBK cells.
To elucidate the molecular mechanism underlying ORF117-induced apoptosis, protein interaction studies were performed to identify potential host targets of ORF117. ORF117 was cloned into the PGEX vector, and the PGEX-ORF117 fusion protein was purified using an E. coli expression system. GST pull-down assays followed by SDS-PAGE analysis revealed host proteins interacting with ORF117 (Figs 3F, S3A and S3B). Liquid chromatography-tandem mass spectrometry analysis identified 74 differentially interacting proteins (S1 Table). Subsequent analysis using the STRING database and Cytoscape identified GAPDH as the host protein with the highest interaction score with ORF117 (Figs 3G, S3C and S3D).
To validate this interaction, co-immunoprecipitation (Co-IP) assays revealed the interaction in 293T cells co-transfected with Flag-GAPDH and Myc-ORF117. Bidirectional Co-IP experiments using both anti-Flag and anti-Myc antibodies confirmed the interaction between ORF117 and GAPDH (Fig 3H). Furthermore, in LSDV-infected MDBK cells, immunoprecipitation with an anti-ORF117 monoclonal antibody (9E3) confirmed the association between viral ORF117 and endogenous GAPDH under physiological infection conditions (Fig 3I).
To further investigate the intracellular localization of ORF117 and GAPDH, indirect immunofluorescence assays revealed the subcellular distribution patterns. In 293T cells co-transfected with pcDNA3.1-flag-GAPDH and pcDNA3.1-myc-ORF117, both proteins exhibited cytoplasmic distribution with significant colocalization, as confirmed by fluorescence intensity profile analysis (Fig 3J, upper panels). Similarly, in LSDV-infected MDBK cells, viral ORF117 (detected using 9E3 antibody) colocalized with endogenous GAPDH in the cytoplasm, evidenced by overlapping fluorescence signals and intensity profiles (Fig 3J, lower panels). These observations indicate direct interaction between ORF117 and GAPDH within the host cell cytoplasm.
LSDV induces apoptosis in MDBK cells via ORF117-mediated activation of the GAPDH-Siah1/p53 pathway
Previous studies have shown that nuclear translocation of GAPDH plays a key role in oxidative stress-mediated apoptosis [33]. To investigate whether LSDV infection affects GAPDH localization, its subcellular distribution was examined using IFA. Results revealed that while GAPDH was predominantly cytoplasmic in uninfected cells, LSDV infection induced significant nuclear accumulation of GAPDH after 48 hours (Fig 4A). Similarly, Western blot assay after isolation of nuclear/plasma proteins demonstrated increased GAPDH levels in the nuclear fraction of LSDV-infected cells compared to control cells (Fig 4B). These data indicate that LSDV infection triggers the nuclear translocation of GAPDH.
(A) Immunofluorescence analysis of GAPDH nuclear translocation in LSDV-infected MDBK cells compared to mock controls. DAPI (blue) stains nuclei, GAPDH (red) detected using specific antibodies, and merged images highlight nuclear localization. Enlarged views indicate nuclear GAPDH (arrowhead). Scale bars = 10 µm. (B) Western blot analysis of GAPDH localization in nuclear and cytoplasmic fractions of LSDV-infected and mock-infected MDBK cells. Histone H3 and α-tubulin were used as nuclear and cytoplasmic markers, respectively. Quantification of nuclear GAPDH relative to Histone H3 is shown (n = 3). (C) Quantitative PCR analysis of pro-apoptotic genes (Bim, Puma, and Noxa) in MDBK cells infected with LSDV or mock-treated at 12, 24, and 48 hpi. LSDV infection significantly upregulates the expression of these genes in a time-dependent manner (n = 3). (D) Efficiency of GAPDH knockdown in MDBK cells using siRNA targeting GAPDH, confirmed by Western blot. (E) Western blot analysis of Cleaved caspase-3 and Siah1 levels in LSDV-infected MDBK cells following GAPDH knockdown. (F) Caspase-3 activity assay in LSDV-infected MDBK cells with GAPDH knockdown compared to siRNA-NC and mock controls. (G) qRT-PCR analysis of Bim, Puma, and Noxa mRNA levels in LSDV-infected MDBK cells following GAPDH knockdown compared to controls. (H) Western blot analysis of cleaved caspase-3 and Siah1 levels in MDBK cells overexpressing ORF117 compared to control cells. (I) Western blot analysis of GAPDH localization in nuclear and cytoplasmic fractions of MDBK-ORF117 and MDBK cells. Histone H3 and α-tubulin were used as nuclear and cytoplasmic markers, respectively. (J) qRT-PCR analysis of Bim, Puma, and Noxa mRNA levels in MDBK cells overexpressing ORF117 compared to control cells (n = 3). (K) Western blot analysis of Cleaved Caspase-3 in MDBK-ORF117 cells with or without GAPDH knockdown using siRNA. (L) qRT-PCR analysis of Bim, Puma, and Noxa mRNA levels in MDBK-ORF117 cells with or without GAPDH knockdown (n = 3). Statistical significance was determined using one-way ANOVA with multiple comparisons (*p < 0.05, **p < 0.01, ***p < 0.001). Error bars represent mean ± SD.
The nuclear translocation of GAPDH in apoptosis requires its interaction with the E3 ubiquitin ligase Siah1. This interaction is critical as Siah1 provides the nuclear localization signal (NLS) for nuclear transport [34]. Once in the nucleus, GAPDH mediates apoptosis through two main mechanisms: (1) S-nitrosylated GAPDH binds to p300/CBP, enhancing its acetyltransferase activity and activating pro-apoptotic genes including p53 [35], and (2) GAPDH stabilizes Siah1, promoting ubiquitin-mediated degradation of nuclear proteins to amplify apoptotic signals [36]. To determine if LSDV infection affects the GAPDH-Siah1 interaction, co-immunoprecipitation was performed using anti-Siah1 antibodies. LSDV infection significantly enhanced GAPDH-Siah1 complex formation in MDBK cells (S4 Fig). Furthermore, qRT-PCR analysis revealed significant upregulation of p53 pathway target genes, including Bim, Puma, and Noxa (Fig 4C), demonstrating activation of p53-dependent apoptotic signaling during LSDV infection.
To further confirm the role of GAPDH in this process, RNA interference (RNAi) reduced GAPDH expression in MDBK cells (Figs 4D and S5). GAPDH knockdown led to a significant reduction in the protein expression levels of Cleaved Caspase-3 and Siah1 (Fig 4E). Furthermore, Caspase-3/7 activity and the transcription levels of Bim, Puma, and Noxa were markedly reduced in GAPDH-deficient cells (Fig 4F and 4G). These results demonstrate that the GAPDH-Siah1/p53 pathway mediates LSDV-induced apoptosis in MDBK cells.
To determine whether ORF117, as a GAPDH-interacting protein, activates the GAPDH-Siah1/p53 pathway, the effects on apoptotic signaling were analyzed in this study using the MDBK-ORF117 cell line. Western blot analysis showed that ORF117 overexpression significantly increased protein levels of both Siah1 and Cleaved Caspase-3 (Figs 4H and S6), while simultaneously promoting the nuclear translocation of GAPDH (Fig 4I). Moreover, qRT-PCR analysis showed that ORF117 overexpression markedly upregulated transcription of p53-dependent pro-apoptotic genes (Bim, Puma, and Noxa) (Fig 4J). These data indicate that ORF117 promotes activation of both the Siah1-dependent pathway and p53-mediated apoptotic signaling.
To further validate the causal relationship between ORF117 and GAPDH in mediating apoptosis, GAPDH expression was knocked down in MDBK-ORF117 cells, and the effects on Cleaved Caspase-3 expression and p53 target gene transcription were analyzed. Western blot results revealed that GAPDH knockdown significantly reduced the expression of Cleaved Caspase-3 in MDBK-ORF117 cells (Fig 4K). Additionally, qRT-PCR analysis demonstrated that GAPDH knockdown markedly suppressed the transcription levels of p53-dependent pro-apoptotic genes (Bim, Puma, and Noxa) in MDBK-ORF117 cells (Fig 4L). In summary, these findings confirm that LSDV activates the GAPDH-Siah1/p53 pathway via ORF117 to induce apoptosis in MDBK cells.
To further investigate the tissue-specific cell death effects of LSDV infection in vivo, analysis was performed on apoptosis markers in kidney tissue from LSDV-infected golden hamsters. Immunofluorescence microscopy revealed significantly increased Cleaved Caspase-3 signals in kidney sections from LSDV-infected animals compared to mock-infected controls (Fig 5A). Western blot analysis of kidney tissue lysates from three independent groups confirmed the substantial elevation of Cleaved Caspase-3 protein levels in LSDV-infected samples relative to mock controls (Fig 5B). Quantitative analysis demonstrated significantly higher Caspase-3 enzymatic activity in infected kidney tissues compared to PBS-treated controls (Fig 5C). These data demonstrate that LSDV infection activates Caspase-3-dependent apoptotic signaling in kidney tissues in vivo.
(A) Immunofluorescence analysis of Cleaved-Caspase-3 (green) in kidney tissues from LSDV-infected and mock-treated hamsters. LSDV (red) detected using specific antibodies, and DAPI (blue) indicates nuclear staining. Merged and enlarged images highlight apoptotic regions. Scale bars = 50 µm. (B) Western blot analysis of Cleaved-Caspase-3 expression in kidney tissues from LSDV-infected and mock-treated hamsters. (C) Caspase-3 enzymatic activity in kidney tissues from LSDV-infected and mock-treated hamsters. (D) Immunofluorescence analysis of GSDMC-N (green) in mammary tissues from LSDV-infected and mock-treated hamsters. LSDV (red) detected using specific antibodies, and DAPI (blue) indicates nuclear staining. Merged and enlarged images highlight pyroptotic regions. Scale bars = 50 µm. (E) Western blot analysis of GSDMC-F and GSDMC-N expression in mammary tissues from LSDV-infected and mock-treated hamsters. (F) Concentrations of pro-inflammatory cytokines IL-18 and IL-1β in plasma and mammary gland tissues from LSDV-infected and mock-treated hamsters. (G) Western blot analysis of Cleaved-Caspase-3 expression in kidney tissues from LSDV-infected and uninfected controls samples (left). Caspase-3 enzymatic activity in bovine kidney tissues was quantified using a specific assay kit (right). (H) Western blot analysis of GSDMC-N expression and concentrations of IL-18 and IL-1β in mammary gland tissues from LSDV-infected and mock-treated bovine samples. Statistical significance was determined using one-way ANOVA with multiple comparisons (*p < 0.05, ***p < 0.001). Error bars represent mean ± SD.
To validate this finding in natural hosts, we conducted similar analyses on bovine clinical tissue samples (S7A Fig) provided by the China Animal Health and Epidemiology Center. Quantitative PCR revealed significantly higher viral copy numbers in all tissues compared to controls (S7B Fig). Western blot analysis showed a marked increase in Cleaved Caspase-3 protein levels in kidney tissues from LSDV-infected cattle compared to uninfected controls (Fig 5G, left). Furthermore, enzymatic activity assays revealed significantly elevated Caspase-3 activity in LSDV-infected bovine kidney tissues, consistent with the observations in the hamster model (Fig 5G, right). These results further support the activation of Caspase-3-mediated apoptosis as a key mechanism of kidney tissue damage during LSDV infection in both experimental and natural hosts.
LSDV infection induces pyroptosis in MAC-T cells
To investigate the mechanism of cell death in MAC-T cells, microscopic analysis revealed morphological changes after LSDV infection. Infected MAC-T cells displayed characteristic pyroptotic features, including cytoplasmic swelling and membrane bubbling, with a distinct fried-egg-like central protrusion (Fig 1F). Meanwhile, to enable a direct comparison between the two cell types, we also performed RNA-seq on MAC-T cells post-infection using the same library preparation and sequencing conditions as for MDBK. The results showed that both cell types exhibited robust innate immune and stress responses; however, MAC-T demonstrated stronger enrichment of inflammasome/pyroptosis-related modules (including NOD-like receptor, IL-17, JAK-STAT, and TLR/RIG-I-like/C-type lectin pathways), consistent with the phenotypic data (S8 Fig).
To confirm the role of pyroptosis in LSDV-induced cell death, MAC‑T cells were pretreated with Z‑VAD‑FMK, a pan‑caspase inhibitor commonly used to block caspase‑dependent cell death including pyroptosis, prior to LSDV infection. Cell viability assays demonstrated that Z-VAD-FMK pretreatment significantly enhanced cell survival in a concentration-dependent manner after LSDV-induced cell death (Fig 6A). TEM analysis revealed that Z-VAD-FMK-treated cells maintained normal cellular architecture, while untreated LSDV-infected cells exhibited characteristic pyroptotic morphological changes (Fig 6B).
(A) Cell viability analysis demonstrating dose-dependent protective effects of Z-VAD-FMK (0-50 μM) against LSDV-induced cell death in MAC-T cells. (B) Phase-contrast microscopy images of mock-infected cells (left), LSDV-infected cells exhibiting cellular swelling and rounding (middle), and Z-VAD-FMK-pretreated cells maintaining normal morphology after LSDV infection (right). Scale bars = 20 μm. (C) LDH release assay in MAC-T and MDBK cells at 24, 48, and 72 hpi. (D) ELISA quantification of IL-1β and IL-18 secretion in culture supernatants from LSDV-infected MAC-T cells at 24, 48, and 72 hpi. (E) TEM images of mock-infected (upper panel) and LSDV-infected (lower panel) MAC-T cells. Higher magnification insets (right) demonstrate normal cellular ultrastructure in mock-infected cells, while LSDV-infected cells exhibit characteristic features of pyroptosis including plasma membrane rupture (green arrows) and cytoplasmic vacuolization (red arrows). Scale bars = 20 µm. Statistical significance was determined using one-way ANOVA with multiple comparisons (**p < 0.01, ***p < 0.001, ****p < 0.0001). Error bars represent mean ± SD.
To quantitatively assess cell membrane integrity during LSDV infection, analysis was performed to measure the release of lactate dehydrogenase (LDH), a stable cytoplasmic enzyme that leaks into the extracellular space during cell lysis. LSDV-infected MAC-T cells showed significantly elevated LDH release compared to mock-infected controls, with levels progressively increasing from 24 to 72 hpi. In contrast, MDBK cells infected with LSDV showed no significant increase in LDH release (Fig 6C). Since pyroptosis is an inflammatory form of cell death, examination of inflammasome-related genes expression was conducted in LSDV-infected cells. RT-qPCR analysis revealed that, compared to mock-infected controls, LSDV infection significantly upregulated the mRNA levels of inflammasome components NLRP1 and NLRP3, as well as pro-inflammatory cytokines IL-1β, IL-18, and IL-1α in MAC-T cells (S9 Fig). ELISA analysis of culture supernatants confirmed that the secretion of IL-1β and IL-18 in infected MAC-T cells increased in a time-dependent manner (Fig 6D).
TEM analysis revealed ultrastructural features of pyroptosis in LSDV-infected MAC-T cells (Fig 6E). Mock-infected cells exhibited intact structures, including clear nuclear membranes, visible nucleoli, and uniformly distributed organelles. In contrast, infected cells displayed hallmark pyroptotic features, including plasma membrane rupture (green arrows) and pronounced cytoplasmic vacuolization (red arrows), indicative of increased membrane permeability and leakage of cellular contents. Compared with Fig 2C, LSDV-infected MDBK cells exhibited typical apoptotic ultrastructural features, including nuclear condensation, chromatin margination, and apoptotic body formation, highlighting the structural differences between the two cell types following LSDV infection.
LSDV induces MAC-T cell pyroptosis through the caspase-8-GSDMC pathway
Previous studies have shown that poxvirus-induced pyroptosis is mediated through GSDMB [37]. To investigate the pyroptotic pathway in LSDV infection, analysis was performed on gasdermin family members in MAC-T cells at 48 hpi. Western blot analysis revealed no substantial changes in GSDME-N or GSDMD-N levels, but showed a significant increase in the N-terminal active fragment of GSDMC (GSDMC-N) (Fig 7A). In contrast, MDBK cells infected under identical conditions showed no detectable GSDMC-N, indicating the absence of appreciable GSDMC cleavage (S10 Fig). These data indicate GSDMC as the key mediator of pyroptosis in LSDV-infected MAC-T cells, indicating a distinct mechanism from previously described poxvirus-induced cell death pathways. To explore the upstream regulation of GSDMC activation, analysis of caspase-8, a known activator of GSDMC [38,39], revealed that both cleaved caspase-8 and GSDMC-N levels increased in a dose- and time-dependent manner following LSDV infection (Fig 7B).
(A) Western blot analysis of GSDMD, GSDME and GSDMC active fragments after LSDV infection of MAC-T cells. (B) Dose- and time-dependent expression of GSDMC-F, GSDMC-N and Cleaved-Caspase 8 proteins in MAC-T cells after LSDV infection at different multiplicities of infection (MOIs: 0.1, 0.5, 1, 2, and 5) and time points (24, 48, and 72 hpi). (C-D) Immunofluorescence microscopy of MAC-T cells infected with LSDV or mock-infected, immunostained for key pyroptosis markers. Scale bars = 10 µm.
To further investigate the molecular mechanism of LSDV-induced pyroptosis, analysis of the subcellular localization of GSDMC-N and cleaved caspase-8 was performed using confocal microscopy. In mock-infected cells, GSDMC-N exhibited diffuse cytoplasmic distribution without apparent membrane accumulation. However, LSDV infection induced significant GSDMC-N aggregation at the plasma membrane, accompanied by nuclear loss in some cells (indicated by arrows) (Fig 7C). Meanwhile, cleaved caspase-8 expression was low and primarily cytoplasmic in mock-infected cells but exhibited marked nuclear accumulation following LSDV infection (Fig 7D). These data indicate that GSDMC-N oligomerization at the plasma membrane leads to pore formation and subsequent pyroptotic cell death.
To confirm that the distinct pyroptotic response in MAC-T cells was not due to differences in viral burden, viral RNA levels were quantified at 16 hpi (MDBK) and 20 hpi (MAC-T) after infection with the same MOI (MOI = 2). The results showed comparable viral loads between the two cell types. Immunofluorescence analysis further demonstrated that LSDV infection induced plasma membrane localization of GSDMC-N in MAC-T cells, consistent with pyroptotic pore formation, whereas no such redistribution was observed in MDBK cells (S2B Fig).
To elucidate the role of caspase-8 in LSDV-induced pyroptosis, LSDV-infected MAC-T cells were treated with the pan-caspase inhibitor Z-VAD-FMK. Administration of Z-VAD-FMK significantly suppressed caspase-8 cleavage and GSDMC-N production, while increasing LSDV replication (Fig 8A). Consistently, Z-VAD-FMK markedly decreased LSDV-induced cell death, as demonstrated by reduced LDH release (Fig 8B).
(A) Western blot analysis showing expression of Cleaved-Caspase 8, GSDMC-F, GSDMC-N, and LSDV proteins in MAC-T cells mock-infected or infected with LSDV, with or without Z-VAD-FMK treatment. (B) LDH release measuring pyroptotic cell death in response to LSDV infection, with or without Z-VAD-FMK treatment. (C) Knockdown efficiency of GSDMC using siRNA (siRNA-GSDMC1, siRNA-GSDMC2, siRNA-GSDMC3, and siRNA-NC [negative control]) evaluated using Western blot. (D) Cell viability of MAC-T cells after GSDMC knockdown and LSDV infection. (E) LDH release measuring pyroptosis in MAC-T cells following GSDMC knockdown and LSDV infection. (F) Viral load (log10 copies/mL) in MAC-T cells after GSDMC knockdown and LSDV infection. (G) Dose-dependent inhibition of Cleaved-Caspase 8 by Z-IETD-FMK (10–60 µM) in LSDV-infected MAC-T cells, determined by Western blot. (H-I) Western blot analysis showing expression of Cleaved-Caspase 8, GSDMC-F, GSDMC-N, and LSDV proteins in MAC-T cells mock-infected or infected with LSDV, with or without Z-IETD-FMK treatment. (J) LDH release measuring pyroptotic cell death in MAC-T cells treated with Z-IETD-FMK, with or without LSDV infection. Statistical significance was determined using one-way ANOVA with multiple comparisons (*p < 0.05, **p < 0.01, ***p < 0.001). Error bars represent mean ± SD.
To further investigate the specific involvement of GSDMC in this process, siRNA-mediated knockdown of GSDMC was performed. Western blot analysis demonstrated efficient GSDMC knockdown across multiple siRNAs (Fig 8C). Functionally, GSDMC knockdown significantly enhanced cell survival (Fig 8D), reduced LDH release (Fig 8E), and notably increased viral replication (Fig 8F). These data indicate GSDMC as a central effector of LSDV-induced pyroptosis.
Additionally, analysis was performed on the effects of Z-IETD-FMK, a specific caspase-8 inhibitor. Optimal Z-IETD-FMK concentrations were determined (Fig 8G). Western blot analysis showed that Z-IETD-FMK treatment effectively reduced Cleaved Caspase-8 and GSDMC-N levels while significantly enhancing viral replication (Fig 8H and 8I). Additionally, LDH release assays demonstrated that Z-IETD-FMK significantly suppressed LSDV infection-induced pyroptosis (Fig 8J). These data indicate that LSDV-induced pyroptosis is mediated through a caspase-8-GSDMC axis.
Finally, analysis was performed on pyroptosis markers in the mammary tissue of LSDV-infected golden hamsters. Immunofluorescence analysis revealed enhanced expression of GSDMC-N in infected mammary tissues (Fig 5D), corroborated by Western blot analysis showing significantly increased GSDMC-N levels compared to controls (Fig 5E). ELISA analysis demonstrated elevated levels of pro-inflammatory cytokines IL-18 and IL-1β in both mammary tissues and plasma of infected animals (Fig 5F), demonstrating that LSDV infection triggers both local and systemic inflammatory responses. These data show that LSDV infection not only induces localized tissue damage through GSDMC-mediated pyroptosis but also promotes systemic inflammation through viremia, potentially facilitating viral dissemination and modulating host immune responses.
To validate these findings in the natural host, bovine mammary tissues were analyzed for pyroptosis markers. Western blot analysis confirmed significantly increased GSDMC-N levels in mammary tissues from LSDV-infected cattle compared to controls (Fig 5H, left). Similarly, ELISA results showed elevated concentrations of IL-18 and IL-1β in infected bovine mammary tissues, consistent with the hamster model (Fig 5H, right). These findings further demonstrate that LSDV infection induces GSDMC-mediated pyroptosis and inflammatory cytokine production in mammary tissues of natural hosts.
Discussion
Poxviruses exhibit complex replication strategies and exceptional adaptability, facilitating immune evasion, cross-species transmission, and systemic infections [14,40,41]. Host cell death, a crucial aspect of viral pathogenesis, functions as both a defense mechanism to restrict viral replication and a tool exploited by viruses to optimize their spread [42,43]. During viral infection, the form of host cell death not only determines the efficiency of infected cell clearance but also significantly influences viral replication and disease progression through regulation of inflammatory responses and immune cell recruitment [44–46].
The present study demonstrates that LSDV exhibits tissue-specific regulation of cell death, utilizing distinct mechanisms to induce apoptosis in kidney-derived MDBK cells and pyroptosis in mammary-derived MAC-T cells. Specifically, LSDV-induced apoptosis in MDBK cells is mediated through the ORF117 protein, which activates the GAPDH-Siah1/p53 pathway. Conversely, pyroptosis in MAC-T cells is triggered via Caspase-8-mediated cleavage of GSDMC. This dual regulatory strategy indicates the virus’s remarkable adaptability to different host cellular environments and provides new insights into its pathogenic mechanisms.
Apoptosis and pyroptosis, two major types of programmed cell death, fulfill distinct and sometimes opposing roles in viral infections [19,47]. Studies demonstrate that certain poxviruses actively inhibit apoptosis to prolong host cell survival. For instance, the Vaccinia virus encodes a Bcl-2-like protein, F1L, which prevents mitochondrial outer membrane permeability and Caspase-9 activation [15,22]; Similarly, the Cowpox virus produces the CrmA protein to block Caspase-1 and Caspase-8 activity, thereby suppressing TNF-induced apoptosis and associated inflammatory responses [23]. The current data demonstrate that LSDV employs a contrasting strategy by specifically promoting apoptosis in MDBK cells.
In MDBK cells, LSDV activates the Caspase-3-dependent apoptotic pathway, leading to significant upregulation of Cleaved Caspase-3 and hallmark apoptotic features such as DNA fragmentation. By inducing this immunologically silent form of cell death, LSDV creates a favorable replication environment while minimizing the risk of triggering inflammatory responses. The current study identified the viral protein ORF117 as a key mediator of this process. ORF117 interacts with the host metabolic enzyme GAPDH, promoting its nuclear translocation and activating the GAPDH-Siah1/p53 signaling pathway, which ultimately drives apoptosis. This discovery demonstrates LSDV’s ability to hijack host metabolism-related factors and repurpose their non-glycolytic functions to regulate cell death. Previous studies have established that GAPDH nuclear translocation can initiate apoptosis under stress conditions [33,36]. The present data link this process to LSDV infection, providing a novel perspective on the virus’s pathogenic strategies and its ability to manipulate host cell machinery for its benefit.
Unlike apoptosis, pyroptosis is a pro-inflammatory form of programmed cell death mediated by the Gasdermin family of proteins, characterized by the formation of membrane pores, cell lysis, and the release of inflammatory cytokines such as IL-1β and IL-18 [47]. Inflammasomes are the core regulators of pyroptosis, and many viruses induce host cell pyroptosis by activating inflammasomes. For example, Influenza Virus promotes host cell pyroptosis by activating the NLRP3 inflammasome and Caspase-1, thereby enhancing antiviral immunity [48], while SARS-CoV-2 induces pyroptosis in lung epithelial cells via the Caspase-8-GSDME pathway, significantly exacerbating inflammatory responses [49]. Although the role of pyroptosis in various viral infections has been extensively studied, the mechanisms of pyroptosis associated with poxviruses require further elucidation. Available evidence indicates that Monkeypox Virus triggers astrocyte pyroptosis by inducing the cleavage of Gasdermin B [37]. These data indicate that poxviruses may regulate pyroptosis through specific molecular mechanisms, but the exact processes require further investigation.
LSDV induces Caspase-8-mediated pyroptosis in MAC-T cells. By cleaving GSDMC and releasing its N-terminal active fragment, LSDV facilitates the formation of membrane pores, leading to the leakage of cellular contents and the release of pro-inflammatory cytokines such as IL-1β and IL-18. As a typical form of inflammatory cell death, pyroptosis plays dual roles in antiviral immunity. On one hand, it activates the host immune system by clearing infected cells and releasing immune signaling molecules, thereby limiting viral spread; on the other hand, excessive inflammatory responses may cause damage to host tissues and could even be exploited by the virus to enhance its transmission [50,51]. This mechanism was further validated using the pyroptosis inhibitor Z-VAD-FMK and the Caspase-8-specific inhibitor Z-IETD-FMK. Analysis demonstrated that inhibiting pyroptosis significantly reduced LDH release, but simultaneously led to a marked increase in viral replication levels. This phenomenon indicates that LSDV-induced pyroptosis is not only a critical immune mechanism to restrict viral spread but also a potential cause of host cell damage due to exacerbated inflammatory responses. These data demonstrate the complex immunoregulatory role of pyroptosis in antiviral processes and emphasize the strategy employed by LSDV to adapt to the host environment by modulating the balance of inflammation. Additionally, the role of caspase activity in regulating LSDV replication was further validated by treating MDBK and MAC-T cells with the pan-caspase inhibitor Z-VAD-FMK (S11 Fig). In MDBK cells, caspase inhibition significantly reduced viral replication, highlighting the pro-replicative role of apoptosis in this cell type. Conversely, in MAC-T cells, caspase inhibition led to a marked increase in viral replication, indicating that pyroptosis plays a critical antiviral role by restricting viral propagation. These findings further support the tissue-specific strategies employed by LSDV to modulate host cell death pathways for optimal replication and dissemination.
The differential induction of apoptosis and pyroptosis by LSDV in different host cells may represent a strategy to adapt to distinct tissue environments and functional demands. Kidneys, as key organs for maintaining systemic homeostasis, exhibit relatively mild immune responses. By inducing apoptosis, LSDV achieves immune silencing, a strategy that avoids triggering excessive inflammatory responses, thereby minimizing damage to renal function [52]. In contrast, mammary tissue, located at the frontline of host-pathogen interactions, features a ductal and glandular environment conducive to pathogen dissemination [53]. LSDV induces pyroptosis to rapidly activate local inflammatory responses and enhance immune cell recruitment, thereby more effectively limiting viral spread within mammary tissues. Similar regulatory patterns are observed in other viruses, such as SARS-CoV-2, which induces pyroptosis via GSDME in lung epithelial cells while evading immune clearance by triggering apoptosis in human monocytes [49]. This tissue-specific regulation of cell death likely reflects an evolutionary advantage of LSDV in adapting to the functional and immune microenvironments of different tissues, offering a novel perspective for understanding its pathogenic strategies.
However, a limitation of this study is the lack of experiments using skin- or lung-derived cells, despite these tissues showing high viral loads in hamsters (Fig 1D). This limitation arises due to the unavailability of widely used bovine skin or lung cell lines, which makes mechanistic dissection in these tissues challenging. Nevertheless, kidney and mammary tissues were chosen because of their clinical relevance to lumpy skin disease. The kidney is involved in systemic viral dissemination and excretion, while the mammary gland is critical due to its association with reduced milk production, a significant cause of economic loss. The findings in kidney-derived MDBK cells and mammary-derived MAC-T cells provide valuable insights into LSDV-induced cell death mechanisms and offer a foundation for future studies once suitable skin and lung models are available.
In summary, the present study reveals novel tissue-specific mechanisms by which LSDV regulates programmed cell death in host cells. The data demonstrate that LSDV induces apoptosis in kidney epithelial cells through ORF117-mediated activation of the GAPDH-Siah1/p53 pathway, while triggering pyroptosis in mammary epithelial cells via the Caspase-8-GSDMC pathway. The differential regulation of cell death was validated in both golden hamster and bovine models, where LSDV infection led to significant apoptosis in renal tissues and prominent pyroptosis in mammary tissues. Notably, inhibition of pyroptosis reduced inflammatory responses but enhanced viral replication, highlighting its dual role in host defense and tissue damage. These data indicate that LSDV has evolved distinct strategies to optimize its replication and spread in different tissue environments: employing immunologically silent apoptosis in kidney cells while activating inflammatory pyroptosis in mammary tissues to regulate local immune responses. This study provides critical insights into the molecular mechanisms underlying LSDV’s tissue tropism and pathogenesis, establishing a theoretical foundation for the development of targeted therapeutics and improved vaccines against this emerging transboundary disease.
Materials and methods
Ethics statement
All experimental procedures were conducted according to animal welfare and ethical guidelines and in accordance with the norms established by the Laboratory Animal Ethics Committee of the China Animal Health and Epidemiology Center. Animal ethics review number: DWFL-2024-04 and DWFL-2024-01.
Male golden hamsters (3 weeks old) were obtained from Beijing Vitalriver Laboratory Animal Technology Co. Bovine kidney and mammary gland tissue samples were collected from clinical specimens obtained through field sampling by the China Animal Health and Epidemiology Center. All experiments involving live LSDV and animals were performed in the BSL-3 facility of the China Animal Health and Epidemiology Center.
Cells and viruses
Human embryonic kidney cells 293 (HEK293T) (ATCC CRL-3216), Vero cells (ATCC CCL-81), MDBK (ATCC CCL-22) and MAC-T (ATCC CRL-10274) were obtained from American Type Culture Collection (ATCC). HEK293T, Vero, and MAC-T cells were maintained in DMEM (Gibco) supplemented with 10% FBS and 1% penicillin/streptomycin (Thermo Scientific), while MDBK cells were maintained in RPMI 1640 (Gibco) supplemented with 10% FBS and 1% penicillin/streptomycin. All cells were maintained at 37°C in 5% CO2. The LSDV/Hongkong/2021 strain was provided by the National Research Center for Exotic Animal Diseases, China Animal Health and Epidemiology Center, Qingdao, China (GenBank: MW732649.1).
Antibodies and reagents
The monoclonal antibody against LSDV ORF117 (9E3) was generated as previously described [54]. The mouse anti-Myc monoclonal antibody and rabbit anti-Flag monoclonal antibody were obtained from Santa Cruz Biotechnology (Dallas, TX, USA). Alexa Fluor 488-conjugated goat anti-mouse IgG secondary antibody, Alexa Fluor 647-conjugated goat anti-rabbit IgG secondary antibody, rabbit anti-DYKDDDDK tag polyclonal antibody, Histone H3 polyclonal antibody, GAPDH polyclonal antibody, Alpha Tubulin polyclonal antibody, Caspase 3/p17/p19 rabbit pAb and DFNA5/GSDME polyclonal antibody were obtained from Proteintech Biotechnology (Wuhan, China). Caspase 3 (active) rabbit monoclonal antibody was obtained from Beyotime (Shanghai, China). GSDMC rabbit polyclonal antibody and Siah1 polyclonal antibody were obtained from Abclonal (Wuhan, China). GSDMD recombinant rabbit monoclonal antibody was obtained from Bioworlde (Nanjing, China), and cleaved-Caspase 8 (Asp384) antibody was obtained from Affinity Biosciences (Cincinnati, OH, USA).
The following reagents were obtained: Annexin V-FITC Apoptosis Detection Kit (Beyotime, Shanghai, China); Cell Counting Kit-8 (Yeasen, Shanghai, China); One-step TUNEL In Situ Apoptosis Kit (Elabscience, Wuhan, China); Caspase 3/7 Activity Assay Kit (Elabscience, Wuhan, China); Glutathione Agarose Beads (Smart Lifesciences, Changzhou, China); LDH Cytotoxicity Assay Kit (Yeasen, Shanghai, China); Staurosporine (MCE, Monmouth Junction, NJ, USA); Z-VAD-FMK and Z-IETD-FMK (Felix, Beijing, China); and Bovine IL-1β and Bovine IL-18 ELISA Kits (Meikebio, Wuhan, China).
Plasmid construction and lentiviral transfection
Based on the LSDV/Hongkong/2021 genome sequence, specific primer sequences were generated using Primer Premier 5 for LSDV envelope proteins (ORF024, ORF028, ORF046, ORF060, ORF064, ORF070, ORF073, ORF100, ORF104, ORF105, ORF108, ORF109, ORF113, ORF117, ORF118, ORF120, ORF122, ORF123 and ORF141) (S2 Table). The amplified target fragments were cloned into pEGFP-N1 vector. The full-length sequences of LSDV ORF117 and GAPDH were cloned into pcDNA3.1-Myc (+) and pcDNA3.1-FLAG (+), respectively. The ORF117 gene was cloned into pGEX-4T-1. The pcDNA3.1-Myc-ORF117 recombinant plasmid was synthesized by Sangyo Bioengineering (Shanghai, China) (S3 Table).
Referring to the genome sequences of pSLenti-EF1-EGFP vector and LSDV/HONGKONG/2021, specific primers for the CDS sequence of ORF117 protein were designed using Primer Premier 5 (S4 Table). The overexpression plasmid pSLenti-EF1-EGFP-ORF117 was constructed and co-transfected with PSPAX and PMD2G plasmids into 293T cells to generate packaged lentivirus. MDBK cells were then infected with the packaged lentivirus and selected using puromycin (2 μg/mL final concentration). The expression of the target gene and protein was verified using fluorescence microscopy and Western blot analysis, respectively.
Plasmids PSPAX and PMD2G for packaging lentivirus were stored in our laboratory. Plasmid transfection was performed using Hieff Trans Liposomal Transfection Reagent (Yeasen, Shanghai, China) and Lipofectamine 3000 (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s instructions.
SiRNA transfection
The following siRNA sequences were designed for GAPDH and GSDMC knockdown: GAPDH1 (GACCUUCACUACAUGGUCUTT), GAPDH2 (UCGGAGUGAACGGAUUCGGTT), GAPDH3 (CCUCAAGAUUGUCAGCAAUTT), GAPDH4 (GGGAAGCUGUGGCGUGACGTT), GSDMC1 (GAACUUGGCGACAAAGACUTT), GSDMC2 (AGGACCCUCUCUCAGUUCUTT), GSDMC3 (AGAGGGAUCGGGUCUCUCAAATT), and GSDMC4 (GCGGGAAGACUCAAGAAAUTT). All siRNAs were synthesized by Gemma Genetics (Suzhou, China). For transfection, siRNA (50 pmol per well) was diluted in transfection buffer containing transfection reagent, mixed with siRNA-Mate plus transfection reagent, and incubated for 10 min at room temperature. The siRNA-transfection reagent mixture was then added to cells at 70–80% confluence and incubated for 48 hours prior to viral infection.
Flow cytometry for apoptosis detection
MDBK and MAC-T cells were plated in six-well plates and infected with LSDV. At designated time points, supernatants were collected, and cells were washed three times with PBS. Cells were detached using trypsin, combined with the collected supernatants, and centrifuged at 1000 × g for 5 minutes. The cell pellet was resuspended in 1 mL PBS for cell counting. Subsequently, 50,000–100,000 cells were centrifuged, resuspended in 195 μL Annexin V-FITC binding buffer, and stained with 5 μL Annexin V-FITC and 10 μL PI. After incubation in the dark at room temperature for 10–20 minutes, samples were kept on ice until analysis. Apoptosis was analyzed by flow cytometry, with Annexin V-FITC-positive cells detected in the FITC channel and PI-positive cells in the PE channel.
Establishment of viral growth curve
MDBK cells were infected with LSDV at an MOI of 1 and incubated for 48 hours. RNA was extracted using the SteadyPure Universal RNA Extraction Kit (Accurate Biotechnology, China) and reverse-transcribed into cDNA with the Evo M-MLV Reverse Transcription Kit. Specific primers targeting the ORF043 gene were designed (amplicon length: 208 bp) using Primer Premier 5 and synthesized by Tsingke Biotechnology Co., Ltd (S5 Table). A standard plasmid containing the full-length ORF043 gene (945 bp) was constructed by amplifying the target sequence, ligating it into the pClone007 vector, and transforming it into DH5α cells. Plasmid copy numbers were calculated, and a 10-fold dilution series was prepared to establish a standard curve via SYBR Green I-based qPCR. Viral growth kinetics were analyzed by infecting MDBK cells with LSDV and collecting supernatants at multiple time points. Viral DNA was extracted from the supernatants and quantified using the established qPCR method to generate a viral growth curve.
Viral growth kinetics in MAC-T cells were determined in parallel using the same infection conditions and qPCR-based quantification described above.
Transmission electron microscopy for cytopathic effects
MDBK and MAC-T cells were plated in 6 cm dishes and infected with LSDV at an MOI of 1. After 24 hours, cells were collected using a cell scraper, centrifuged at 1000 × g for 5 mins, and the supernatant was removed. The cell pellet was fixed in 1 mL of 2.5% glutaraldehyde at 4°C overnight. Fixed samples were sent to Wuhan Servicebio Technology Co., Ltd. for ultrastructural analysis of cellular alterations and viral particle distribution.
Western blot for apoptosis detection
MDBK and MAC-T cells were infected with LSDV and total protein was collected at specific time points. Cells were lysed with RIPA buffer containing 1 mM PMSF on ice for 30 min, followed by ultrasonic fragmentation (4 s on/4 s off cycle) and centrifugation at 12,000 × g for 15 min at 4°C to collect supernatants. Protein concentration was determined by BCA method using bovine serum albumin as standard. Proteins were mixed with 5 × Sampling Buffer, boiled for 10 minutes, and stored at -80°C.
Protein samples (30–40 μg per lane) were separated by SDS-PAGE at a constant voltage of 130 V and then wet transferred to PVDF membranes at 300 mA, with the transfer time adjusted to the molecular weight of the target protein. Membranes were blocked with 10% skim milk for 2 hours, incubated overnight at 4°C with primary antibody, rinsed with PBST, and then incubated with secondary antibody for 40 minutes at room temperature. Protein expression was visualized using the ECL detection system and imaged in a dark room.
TUNEL assay for apoptosis detection
MDBK cells were seeded on coverslips in 12-well plates and cultured to 80% confluence. Cells were infected with LSDV at an MOI of 1, with staurosporine-treated cells as a positive control and PBS-treated cells as a negative control. After 48 hours, apoptosis was detected using a one-step TUNEL in situ apoptosis kit.
Cells were fixed at room temperature for 20 minutes, permeabilized at 37°C for 10 minutes, and incubated with 100 μL of TdT equilibration buffer at 37°C for 10–30 minutes. After removing the buffer, cells were incubated with 50 μL of labeling solution at 37°C for 1 hour in the dark, followed by washing with PBS. Subsequently, cells were incubated overnight at 4°C with a monoclonal antibody targeting LSDV ORF117 protein. After washing, cells were incubated with a red fluorescent secondary antibody at room temperature for 30 minutes, counterstained with DAPI for 5 minutes, and mounted with anti-fade mounting medium. Apoptotic cells (green fluorescence) and LSDV-infected cells (red fluorescence) were visualized using an inverted fluorescence microscope.
Transcriptome sequencing to analyze the effects of LSDV on host cell transcription
MDBK cells were seeded in 10 cm dishes and divided into 8 groups (4 infection groups and 4 control groups), with three replicates per group. When cells reached 80% confluence, the infection groups were infected with LSDV (MOI = 1), and the control groups were treated with an equal volume of UV-inactivated LSDV. Total RNA was extracted at 2, 8, 16, and 24 hpi.
RNA was extracted using the Trizol method and assessed for quality and concentration using a Nanodrop spectrophotometer and Agilent 2100 bioanalyzer. RNA integrity was verified by agarose gel electrophoresis. The 24 RNA samples that met quality standards were sent to Guangzhou Gene Denovo Biotechnology Co., Ltd. for transcriptome sequencing. The transcriptome sequencing data generated in this study have been deposited in the NCBI BioProject database under the accession number [PRJNA1423217] (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1423217).
For transcriptome analysis of MAC-T cells, cells were infected with LSDV at an MOI of 1 or treated with UV-inactivated virus as control, and total RNA was collected at 24 hpi (n = 3 per group). Library preparation, sequencing and bioinformatic analyses were performed using the same protocols as described above for MDBK cells.
GST pull-down assay
The pGEX-ORF117 recombinant plasmid was transformed into BL21 cells, and recombinant protein expression was induced with 1 mM IPTG at 16°C for 16 hours. The protein was purified using AKTA chromatography and analyzed by SDS-PAGE and Coomassie blue staining. Purified pGEX-ORF117 protein was incubated with MDBK cell lysates, and glutathione agarose beads were used for pulldown. After washing with NP40 lysis buffer, the bound proteins were analyzed by SDS-PAGE and sent for mass spectrometry at Genecreate Biotech.
Co-immunoprecipitation (Co-IP)
Both pcDNA3.1-myc-ORF117 and pcDNA3.1-flag-GAPDH recombinant plasmids were co-transfected into 293T cells using liposome reagents. After 48 hours, cells were lysed with NP40 lysis buffer, and lysates were incubated with anti-flag or anti-myc antibodies for immunoprecipitation. Protein A/G agarose beads were used to capture immune complexes, which were washed and analyzed by Western blot. Additionally, MDBK cells infected with LSDV (MOI = 1) were used for Co-IP with ORF117 monoclonal antibody 9E3 to confirm interactions.
Indirect immunofluorescence for colocalization
293T cells were seeded in confocal dishes and co-transfected with pcDNA3.1-flag-GAPDH and pcDNA3.1-myc-ORF117 plasmids, with pcDNA3.1-flag-GAPDH and pcDNA3.1-myc as controls. After 48 hours, cells were fixed with 4% paraformaldehyde at 4°C for 15 minutes, permeabilized with 0.5% Triton X-100, and blocked with 5% goat serum at 37°C for 30 minutes. Cells were incubated overnight at 4°C with mouse anti-Myc and rabbit anti-Flag primary antibodies. After washing, Alexa Fluor 488-conjugated anti-mouse and Alexa Fluor 647-conjugated anti-rabbit secondary antibodies were added and incubated at 37°C for 1 hour. DAPI was used for nuclear staining, and samples were mounted with antifade reagent for observation under a laser confocal microscope.
MDBK cells were infected with LSDV (MOI = 1) for 48 hours, with PBS-treated cells as a control. Cells were fixed and permeabilized as described above. ORF117 monoclonal antibody 9E3 and rabbit anti-GAPDH antibody were used as primary antibodies, and the same secondary antibodies were applied. Colocalization of ORF117 and GAPDH was observed under a laser confocal microscope.
Lactate dehydrogenase release assay
MDBK and MAC-T cells were inoculated in 96-well plates and cultured to approximately 70% confluence for LSDV infection. According to the LDH Cytotoxicity Assay Kit manual, the experimental groups included: wells without cell culture medium (background control), PBS-treated cells (negative control), uninfected cells for maximal enzyme activity determination, and LSDV-infected cells (time points: 24, 48, 72 hpi; MOI: 0.5, 1, 2, 3, 5).
One hour before the assay time point, 10% of the original culture volume of LDH-releasing reagent was added to the maximal enzyme activity control wells and incubated until the assay time point. The plates were centrifuged at 400 × g for 5 min, and 120 μL of supernatant from each well was transferred to a new 96-well plate. The 1 × INT solution and LDH assay working solution were prepared according to instructions. Working solution (60 μL) was added to each well and incubated for 30 min at room temperature in the dark. The absorbance was measured at 490 nm using a plate reader.
The cytotoxicity rate was calculated using the following formula: cytotoxicity rate (%) = (absorbance of treated samples - absorbance of sample control wells)/(absorbance of maximal enzyme activity - absorbance of sample control wells) × 100.
Animal experiment
Three-week-old male golden hamsters were selected and randomly divided into two groups (n = 5). The experimental group was inoculated with 1 mL of 106 TCID₅₀/mL of the LSDV strain by subcutaneous injection, and the control group was inoculated with the same volume of PBS. After inoculation, body weight changes were recorded at 2-day intervals, and samples of serum, heart, liver, spleen, lungs, kidneys, dorsal skin, and mammary gland tissues were collected on day 14.
Genomic DNA was extracted from bovine kidney and mammary gland tissues, as well as from hamster tissues. The viral load in each tissue was subsequently quantified by quantitative real-time PCR. Kidney and mammary tissues were sent to Servicebio Company for indirect immunofluorescence with antibodies against Caspase-3 and GSDMC, respectively.
Supporting information
S1 Fig. A fluorescence quantitative PCR method for detection of LSDV was designed based on ORF043 gene.
(A) ORF043 gene was amplified by PCR; (B-C) Construction of the standard curve for the fluorescence quantitative PCR method.
https://doi.org/10.1371/journal.ppat.1013982.s001
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S2 Fig. Detection of apoptotic and pyroptotic markers in LSDV-infected MDBK and MAC-T cells with comparable viral loads.
(A) Under comparable viral loads, Cleaved Caspase-3 (green) was detected in MDBK cells, indicating apoptosis, but not in MAC-T cells. LSDV (red) and nuclei (DAPI, blue) are also shown. (B) With comparable viral loads, LSDV infection induced plasma membrane localization of GSDMC-N (green) in MAC-T cells, indicative of pyroptosis, while no such redistribution was observed in MDBK cells. Scale bars = 10 μm.
https://doi.org/10.1371/journal.ppat.1013982.s002
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S3 Fig. The GST-pull down assay was used to screen the interacting proteins between ORF117 and MDBK cells.
(A) Double-digestion identification of PGEX-ORF117 plasmid, M: DL5000 DNA marker, 1: PGEX-ORF117 post digestion product, 2: PGEX-ORF117 plasmid (B) SDS-PAGE analysis of the expression and purification of PGEX-ORF117 fusion protein, M: protein molecular weight standard, 1: PGEX protein purification product, 2: PGEX-ORF117 protein purification product.
https://doi.org/10.1371/journal.ppat.1013982.s003
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S4 Fig. Effect of LSDV infection on GAPDH-Siah1 interaction.
https://doi.org/10.1371/journal.ppat.1013982.s004
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S5 Fig. Identification of GAPDH knockdown efficiency.
(A) Quantitative analysis of GAPDH knockdown efficiency detected by Western blot; (B) Relative quantitative analysis of GAPDH knockdown efficiency detected by fluorescence quantitative PCR.
https://doi.org/10.1371/journal.ppat.1013982.s005
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S6 Fig. ORF117 overexpression enhances key markers of apoptotic signaling in MDBK cells.
https://doi.org/10.1371/journal.ppat.1013982.s006
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S7 Fig. Clinical manifestations of LSDV‑infected cattle and the viral loads in tissues.
(A) Representative photographs of LSDV‑infected cattle showing characteristic multifocal cutaneous nodules and plaques on the trunk and udder region. (B) Quantification of viral loads in bovine kidney and mammary gland tissues by qPCR (n = 3).
https://doi.org/10.1371/journal.ppat.1013982.s007
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S8 Fig. KEGG pathway enrichment analysis of differentially expressed genes in LSDV-infected MAC-T cells.
Circle size corresponds to the number of genes, and color intensity indicates statistical significance (-log10(q-value)).
https://doi.org/10.1371/journal.ppat.1013982.s008
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S9 Fig. LSDV infection upregulates the expression of inflammasome-related genes in MAC-T cells.
https://doi.org/10.1371/journal.ppat.1013982.s009
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S10 Fig. Western blot analysis of GSDMC active fragments after LSDV infection of MDBK and MAC-T cells.
https://doi.org/10.1371/journal.ppat.1013982.s010
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S11 Fig. Effects of caspase inhibition on LSDV replication in MDBK and MAC-T cells.
MDBK and MAC-T cells were infected with LSDV at an MOI of 1, with or without treatment with the pan-caspase inhibitor Z-VAD-FMK. Viral genome copies were quantified at different time points post-infection using qPCR.
https://doi.org/10.1371/journal.ppat.1013982.s011
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S1 Table. Mass spectrometric identification of LSDV ORF117-interacting proteins.
https://doi.org/10.1371/journal.ppat.1013982.s012
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S5 Table. Primer design for fluorescent quantitative PCR of LSDV ORF043 gene.
https://doi.org/10.1371/journal.ppat.1013982.s016
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