https://orcid.org/0009-0000-6256-7588
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Figures
Abstract
Background
Cardiotoxin (CTX) from Naja atra venom is a principal virulence factor responsible for progressive local tissue necrosis and systemic inflammation following snakebite. Despite its clinical importance, the molecular mechanisms underlying CTX-induced skin injury remain poorly defined.
Methods
We employed a two-arm strategy combining transcriptome-guided discovery with mechanistic functional validation. In the transcriptomic arm, C57BL/6 mice received intradermal CTX injection (120 μg/50 μL), and skin tissues were harvested at 6, 12, and 24 h post-injection for RNA-seq analysis. Differentially expressed genes (DEGs) were screened and subjected to KEGG/GO enrichment and ssGSEA-based cell death mode scoring. In the functional validation arm, a separate cohort of mice was assessed at 72 h post-injection, with gross necrosis area quantified, followed by H&E staining, immunohistochemistry (IHC), and Western blot. In vitro validation was performed in human HaCaT keratinocytes using CCK-8 cytotoxicity assay, optical microscopy, transmission electron microscopy (TEM), propidium iodide/DAPI (PI/DAPI) dual staining, ROS detection, ELISA for IL-1β, LDH release assay, and pharmacological inhibition with MCC950 (NLRP3 inhibitor), VX-765 (caspase-1 inhibitor), and N-acetylcysteine (NAC, ROS scavenger).
Results
RNA-seq identified 2,490 DEGs (|log2FC| > 1, FDR < 0.01; 1,047 upregulated, 1,443 downregulated). KEGG enrichment revealed that the NOD-like receptor signaling pathway was the most significantly enriched pathway (enrichment fold = 6.8, p_adj < 0.001, with 42 differentially expressed genes annotated to this pathway). Among eight assessed cell death modalities, ssGSEA demonstrated that pyroptosis had the highest activation score (p < 0.05). Six canonical NLRP3/caspase-1/GSDMD pathway genes—Nlrp3, Pycard, Gsdmd, Il18, Nfkb, and Tlr4—were continuously upregulated from 6 to 24 h. Western blot confirmed both full-length GSDMD and its cleaved N-terminal fragment (GSDMD-N), along with NLRP3 inflammasome activation, in CTX-treated skin tissues and HaCaT cells. In vitro, CTX induced characteristic pyroptotic morphology and pyroptotic bodies (1–5 μm by TEM). Western blot confirmed NLRP3/GSDMD-N upregulation in HaCaT cells. CTX also induced dose-dependent intracellular ROS accumulation (DCFH-DA fluorescence). Importantly, all three inhibitors—NAC (ROS scavenger), MCC950 (NLRP3 inhibitor), and VX-765 (caspase-1 inhibitor)—significantly attenuated CTX-induced LDH release, IL-1β secretion, and GSDMD cleavage (all p < 0.0001), confirming the mechanistic hierarchy: ROS → NLRP3 → caspase-1 → GSDMD.
Conclusion
CTX induces intracellular ROS accumulation that activates the NLRP3/caspase-1/GSDMD pyroptotic cascade as an important mechanism contributing to skin tissue necrosis through membrane pore formation and inflammatory amplification. Pharmacological inhibition (NAC, MCC950, VX-765) confirmed a hierarchical ROS → NLRP3 → caspase-1 → GSDMD cascade. The ROS–NLRP3–caspase-1–GSDMD axis constitutes a tractable therapeutic target for Naja atra envenomation.
Author summary
Chinese cobra (Naja atra) snakebite causes severe local tissue necrosis, often leading to permanent disability or amputation. The primary toxin responsible is cardiotoxin (CTX), but its mechanism of action has been unclear. Using transcriptomics and functional experiments in mice and human skin cells, we found that CTX triggers pyroptosis—an explosive form of programmed cell death that releases inflammatory signals and amplifies tissue injury. Mechanistically, CTX induces reactive oxygen species (ROS) accumulation, activating a cascade of NLRP3 → caspase-1 → GSDMD, leading to membrane pores and cell death. Importantly, three inhibitors—NAC (ROS scavenger), MCC950 (NLRP3 inhibitor), and VX-765 (caspase-1 inhibitor)—significantly reduced pyroptosis in human skin cells. These findings identify the ROS–NLRP3–GSDMD pathway as a promising therapeutic target for reducing tissue necrosis after cobra snakebite, potentially decreasing the need for amputation.
Citation: Qin N, Zhai Y, Cao H, Dong C, Liao J, Li G, et al. (2026) Cardiotoxin from Naja atra Activates the NLRP3/Caspase-1/GSDMD Pyroptosis Pathway to Induce Skin Tissue Injury. PLoS Negl Trop Dis 20(7): e0014495. https://doi.org/10.1371/journal.pntd.0014495
Editor: Wuelton Monteiro, Fundação de Medicina Tropical Doutor Heitor Vieira Dourado: Fundacao de Medicina Tropical Doutor Heitor Vieira Dourado, BRAZIL
Received: December 12, 2025; Accepted: June 23, 2026; Published: July 13, 2026
Copyright: © 2026 Qin 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 raw RNA-sequencing data generated and analyzed in this study have been deposited in the NCBI Gene Expression Omnibus (GEO) and are publicly accessible under accession number GSE334138 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE334138).
Funding: The National Natural Science Foundation of China (Grant nos. 81560318 to WW and 81860346 to WW), Training Project of “139” Program for High-Level Medical Talents in Guangxi (No. G201903034 to WW), and Research and Demonstration Application of Snakebite Bigdata Platform Based on Virtual Standardized Snakebite Treatment Model (No. 22QYCX0046 to WW). The funders had no role in 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.
1. Introduction
Snakebite envenomation remains a critical neglected tropical disease, causing an estimated 81,000–138,000 deaths and 400,000 permanent disabilities annually worldwide [1,2]. Naja atra (Chinese cobra) is among the most medically significant venomous snakes in Southeast and East Asia, where its bites account for a substantial proportion of reported envenomation cases [3]. The clinical hallmark of N. atra envenomation is severe local tissue necrosis—a debilitating complication that frequently progresses to deep tissue destruction, secondary infection, and in refractory cases, amputation [4,5].
Cardiotoxin (CTX), also known as cytotoxin, constitutes 40–60% of the total protein in N. atra venom and is the primary effector of local tissue injury [6]. CTX belongs to the three-finger toxin superfamily, characterized by a compact β-sheet structure stabilized by four conserved disulfide bonds that confers remarkable thermal and proteolytic stability [7,8]. At the cellular level, CTX inserts directly into phospholipid bilayers, disrupting membrane integrity via loop-region penetration of the hydrophobic bilayer core, triggering downstream cytotoxic cascades [9]. However, the precise intracellular signaling events linking CTX-mediated membrane perturbation to the observed pattern of progressive necrotic tissue injury remain incompletely characterized.
Pyroptosis is a lytic, inflammatory form of programmed cell death mediated by the gasdermin (GSDM) protein family, which executes the final effector step by forming membrane pores that release pro-inflammatory cytokines IL-1β and IL-18 [10–12]. The canonical pyroptosis pathway involves pattern recognition receptor (PRR)-mediated inflammasome assembly—most commonly the NLRP3 inflammasome—leading to autocatalytic cleavage of procaspase-1 into active caspase-1, which in turn cleaves pro-IL-1β, pro-IL-18, and gasdermin D (GSDMD) [13]. The N-terminal fragment of GSDMD (GSDMD-N) then oligomerizes and inserts into the plasma membrane, forming large transmembrane pores (10–15 nm inner diameter) that cause osmotic lysis and secondary release of cytosolic DAMPs [14, 15]. This NLRP3/caspase-1/GSDMD axis has been implicated in a growing range of sterile injury conditions, including burns, ischemia-reperfusion injury, chemical toxicant-induced tissue damage, and crystalline substance-induced sterile inflammation [16,17].
Recent transcriptomic studies have begun to reveal the complexity of gene expression reprogramming following venom exposure in elapid and viperid species [18,19]. Nonetheless, systematic functional validation of specific cell death pathways triggered by snake venom cytotoxins—and in particular, whether pyroptosis constitutes a causally required mechanism of tissue damage—has not been rigorously established.
In this study, we employed an integrated transcriptomics-to-mechanism strategy in a validated N. atra CTX mouse skin injury model. We first performed unbiased genome-wide transcriptomic profiling across three time points to identify the dominant cell death signaling programs activated by CTX. We then conducted comprehensive functional validation, including pharmacological inhibition of the NLRP3/caspase-1/GSDMD pathway both in vivo and in vitro, to establish causality. Our findings demonstrate that CTX activates pyroptosis as its dominant cytotoxic mechanism and identify key nodes within this pathway as potentially tractable therapeutic targets.
2. Materials and methods
2.1. Ethics statement
Adult C57BL/6 mice (8–10 weeks; 22–25 g, Equal numbers of males and females) were obtained from Laboratory Animal Center of Guangxi Medical University and maintained under specific-pathogen-free (SPF) conditions (12 h light/dark cycle, ad libitum food and water; 22 ± 2°C). All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Guangxi Medical University Laboratory Animal Ethics Committee (Approval No: 202504006) and conducted in accordance with the ARRIVE guidelines.
2.2. Reagents and antibodies
CTX from N. atra venom (≥98% purity by RP-HPLC; Lot No: #335) was purchased from ZhongxinDongtai (Laiyang) Nanogene Biotechnology Co, Ltd. (Laiyang, China). The following primary antibodies were used: anti-NLRP3 (Cat. No. AG-20B-0014; AdipoGen Life Sciences), anti-caspase-1 (Cat. No. 22915–1-AP; Proteintech), anti-GSDMD (Cat. No. 20770–1-AP; Proteintech), anti-IL-1β (Cat. No. 16806–1-AP; Proteintech), anti-IL-18 (Cat. No. 10663–1-AP; Proteintech). MCC950 (Selleckchem, S6598), VX-765 (Selleckchem, S2228) and N-acetylcysteine(Selleckchem, S3656) were dissolved in DMSO and stored at −80°C. CCK-8 kit (Beyotime), DCFH-DA probe (Beyotime), LDH cytotoxicity assay kit (Beyotime), and IL-1β ELISA kit (Beyotime) were used according to manufacturers’ instructions.
2.3. Experimental models and animal grouping
Transcriptomic arm: C57BL/6 mice (n = 40) were randomly assigned to four groups (n = 10/group): healthy control (Control, PBS vehicle, 50 μL intradermal injection), CTX-6 h, CTX-12 h, and CTX-24 h. was selected based on pilot experiments showing consistent necrotic lesions of approximately 0.5 cm in diameter at 72 h post-injection, which was used as the criterion for successful model establishment [20–22], ensuring good reproducibility and reliability of the model. CTX was administered by intradermal injection into the dorsal skin. Tissues at the injection site were harvested at the indicated time points, snap-frozen in liquid nitrogen, and stored at −80°C for RNA-seq.
Functional validation arm: A separate cohort of C57BL/6 mice was assigned to two groups (n = 6/group): Control (PBS vehicle, 50 μL intradermal injection) and CTX (120 μg/50 μL). All mice were assessed at 72 h post-injection for gross necrosis area, histopathological analysis (H&E staining), IHC and Western blot analysis.
A sample size justification was performed based on prior studies reporting GSDMD pathway activation under similar inflammatory injury models; power analysis (α = 0.05, power = 0.80, effect size estimated from pilot experiments) indicated a minimum of 6 animals per group; 10 animals per group were used to account for potential attrition.
2.4. Gross observation and necrosis area measurement
At 72 h post-injection, animals were photographed under standardized conditions (fixed camera height, white background). Necrosis area (mm²) was measured from calibrated digital photographs using ImageJ software (NIH, v1.53) by two independent observers blinded to group allocation. Inter-observer agreement was assessed by intraclass correlation coefficient (ICC).
2.5. Histopathological assessment
Harvested skin tissue specimens were fixed in 10% neutral-buffered formalin for 24 h, paraffin-embedded, and sectioned at 5 μm. Sections were stained with hematoxylin and eosin (H&E) and evaluated by a board-certified pathologist (blinded to treatment groups) for epidermal disruption, dermal edema, inflammatory cell infiltration, and tissue necrosis.
2.6 RNA extraction, library preparation, and transcriptomic analysis
Total RNA was extracted from frozen skin tissue using TRIzol reagent (Invitrogen) per the manufacturer’s protocol. RNA quality was assessed by Agilent 2100 Bioanalyzer (RNA Integrity Number ≥ 7.0 required). Strand-specific mRNA libraries were prepared from polyA-selected RNA using the NEBNext Ultra RNA Library Prep Kit. Paired-end sequencing (150 bp) was performed on an Illumina NovaSeq 6000 platform to a depth of ≥20 million reads per sample. Raw reads were quality-controlled with FastQC and trimmed with Trimmomatic. Clean reads were aligned to the Mus musculus reference genome (GRCm38/mm10) using HISAT2 (v2.2.1). Gene expression was quantified with featureCounts (Subread v2.0.0). DEGs between CTX-treated and Control groups were identified using DESeq2 (v1.34) with thresholds of |log2FC| > 1 and adjusted p-value (Benjamini–Hochberg FDR) < 0.01. GO and KEGG functional enrichment analyses were performed using clusterProfiler (R package, v4.2). The ssGSEA algorithm (GSVA R package) was applied to score eight cell death mode gene signatures.
2.7. Cell culture and CTX treatment
Human immortalized HaCaT keratinocytes Cell Bank, Chinese Academy of Sciences (Shanghai, China) were maintained in DMEM supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin at 37°C under 5% CO2. For dose-response and time-course experiments, cells were seeded at 1 × 105 cells/well in 96-well plates, allowed to adhere overnight, and then treated with CTX at concentrations of 0, 5, 10, 15, and 20 μg/mL for varying durations (2, 4, and 8 h). For pathway inhibition experiments, cells were pre-treated with MCC950 (10 μM, 1 h), VX-765 (20 μM, 1 h), or NAC (N-acetylcysteine, 5 mM, 1 h) prior to CTX stimulation.
The selection of HaCaT keratinocytes as the in vitro model is justified on the basis that skin keratinocytes are the primary cell type exposed to CTX following intradermal injection, and HaCaT cells are the most widely used human keratinocyte line for studying inflammatory and pyroptotic signaling in skin injury contexts. We acknowledge that as a transformed line, HaCaT cells lack some innate immune components present in primary keratinocytes; accordingly, results from this model are interpreted as demonstrating the cell-autonomous capacity for CTX to activate GSDMD-dependent pyroptosis in epithelial cells, complementing the in vivo mouse findings.
2.8 CCK-8 cell viability assay
Cell viability was determined by CCK-8 assay per the manufacturer’s protocol. Absorbance was measured at 450 nm using a microplate reader. Results are expressed as percentage of viable cells relative to untreated controls.
2.9 Morphological analysis
Optical microscopy: Cells were cultured in 6-well plates, treated with CTX (10 μg/mL, 8 h), and observed under an inverted optical microscope (Olympus CKX53) at ×100 and ×200 magnification. LPS (1 μg/mL, 6 h) + ATP (5 mM, 30 min) was included as a positive pyroptosis control. Morphological features of pyroptosis (cell swelling, membrane blebbing, cytoplasmic vacuolation) were documented.
Transmission electron microscopy (TEM): Cells were fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4), post-fixed in 1% osmium tetroxide, dehydrated through a graded ethanol series, and embedded in Spurr’s resin. Ultrathin sections (70 nm) were stained with uranyl acetate and lead citrate and examined on a JEOL JEM-1400 at 80 kV. Pyroptotic bodies were identified as membrane-bound vesicles 1–5 μm in diameter with condensed nuclear material.
2.10. Propidium Iodide/DAPI dual staining
Membrane integrity was assessed by PI/DAPI staining. Cells were washed with PBS, incubated with PI (1 μg/mL) and DAPI (1 μg/mL) in PBS for 15 min at room temperature, and imaged by fluorescence microscopy (excitation/emission: PI, 535/617 nm; DAPI, 360/460 nm). PI-positive cells (membrane-compromised) were quantified as a percentage of total DAPI-positive cells.
2.11. Intracellular ROS Detection
Intracellular ROS levels were measured using the fluorescent probe DCFH-DA. Cells were incubated with 10 μM DCFH-DA at 37°C for 20 min, washed twice with serum-free DMEM, and fluorescence intensity was measured (excitation/emission: 485/530 nm).
2.12. LDH release assay
LDH release into culture supernatant was quantified using the LDH Cytotoxicity Assay Kit and expressed as a percentage relative to the maximum LDH release (total lysis control).
2.13. IL-1β ELISA
Secreted IL-1β in conditioned medium and tissue lysates was quantified by ELISA according to the manufacturer’s instructions. Lower limit of detection: 2.0 pg/mL.
2.14. Western Blot
Protein was extracted from cells or tissues using RIPA lysis buffer supplemented with protease and phosphatase inhibitors. Protein concentration was determined by BCA assay. Equal quantities of protein (30 μg/lane) were resolved by SDS-PAGE (10% or 12% gels) and transferred to PVDF membranes (0.45 μm). Membranes were blocked with 5% non-fat milk in TBST for 1 h at RT and incubated with primary antibodies overnight at 4°C (anti-NLRP3, 1:1000; anti-caspase-1, 1:1000; anti-GSDMD, 1:1000 [recognizes both full-length and N-terminal fragment]; anti-IL-1β, 1:1000; anti-IL-18, 1:1000; anti-β-actin, 1:5000). After washing, membranes were incubated with HRP-conjugated secondary antibody (1:10,000) for 1 h at RT. Bands were visualized by ECL and imaged on a ChemiDoc system. Densitometry was performed using ImageJ; all protein levels were normalized to β-actin. Western blot experiments were performed in triplicate biological replicates (n = 3 independent experiments), and representative blots are shown.
2.15. Statistical analysis
Statistical analyses were performed in GraphPad Prism 9.0 and R (v4.2.0). Data are expressed as mean ± standard deviation (SD). For comparisons between two groups, unpaired two-tailed Student’s t-test was used for normally distributed data and Mann–Whitney U test for non-normal data. For multiple group comparisons, one-way ANOVA followed by Tukey’s post hoc test was applied. Correlation analyses were performed using Pearson’s correlation coefficient. All statistical tests were two-tailed; p < 0.05 was considered statistically significant.
3. Results
3.1. CTX induces progressive skin necrosis in a time-dependent manner
To establish a robust and reproducible CTX-induced skin injury model, C57BL/6 mice received intradermal injection of CTX (120 μg/50 μL). Gross examination revealed a dynamic, progressive tissue response. During the early phase (0–12 h), the dominant finding was progressive erythema and edema, with a local reaction area of 72.5 ± 5.3 mm² at 6 h expanding to 116.2 ± 10.7 mm² at 12 h. In the intermediate phase (12–24 h), inflammatory infiltration intensified with emergence of early necrotic changes. Frank necrosis developed and consolidated during the late phase (24–72 h), reaching a plateau necrosis area of 50.8 ± 4.4 mm² at 72 h (p < 0.05 vs. control; Fig 1A-1B). The necrosis area at 96 h (55.3 ± 3.9 mm²) was not significantly different from the 72 h measurement (p > 0.05), confirming that tissue injury stabilizes by 72 h post-injection. Histopathological assessment (H&E staining) at 72 h confirmed full-thickness epidermal disruption, massive inflammatory cell infiltration, and dermal edema in the CTX group compared to PBS-injected controls (Fig 1C). These findings establish that CTX induces a progressive, time-dependent skin injury response—from early edematous/erythematous reaction to established necrosis—and define 72 h as the optimal primary endpoint for functional intervention studies, consistent with the clinical presentation of N. atra envenomation [4,5].
(A) Representative gross photographs of control and CTX-treated (72 h) skin (scale bar: 1 cm; n = 6/group). (B) Quantification of necrosis area (mm²) at each time point (n = 6/group; mean ± SD; one-way ANOVA, Tukey post hoc). (C) H&E staining of CTX-exposed and control skin sections at 72 h (×100; scale bar: 100 μm). Arrows indicate epidermal disruption, inflammatory infiltrate, and dermal edema.
3.2. Transcriptomic profiling identifies extensive gene expression reprogramming following CTX exposure
RNA-seq analysis of CTX-exposed skin tissues (6, 12, and 24 h post-injection) versus healthy controls identified 2,490 DEGs (|log2FC| > 1, FDR < 0.01; 1,047 upregulated, 1,443 downregulated) (Fig 2A). Among the top upregulated genes were inflammatory mediators and cytoskeletal remodeling factors (Cfl1, Pfn1, Ifitm3, Timp1, Serpina3n), while genes associated with muscle structural integrity and metabolic homeostasis (Txnip, Car3, Tpm2, Tcap) were predominantly downregulated.
(A) Volcano plot showing DEGs (CTX 24 h vs. Control). Top 10 upregulated and downregulated genes labeled. (B) GO Biological Process enrichment. (C) GO Molecular Function enrichment. (D) GO Cellular Component enrichment. (E) KEGG pathway enrichment bubble plot; bubble size: gene count; color: adjusted p-value. NOD-like receptor signaling pathway highlighted.
GO term enrichment analysis revealed significant enrichment in biological processes including mRNA processing, cellular component disassembly, muscle cell differentiation, and ubiquitin-dependent protein catabolism (Fig 2B–2D). KEGG pathway enrichment identified the NOD-like receptor (NLR) signaling pathway as the most significantly enriched pathway (enrichment fold = 6.8, P_adj < 0.001, 42 DEGs), followed by signaling pathways governing cell growth and death, and protein processing in the endoplasmic reticulum (Fig 2E). The predominant enrichment of NLR signaling directly implicates inflammasome activation as a central molecular response to CTX.
3.3. Pyroptosis is the predominant cell death mode activated by CTX
To systematically assess which cell death pathways are activated by CTX, ssGSEA was applied to score eight distinct cell death gene signatures across all samples. Compared to healthy controls, CTX treatment significantly elevated scores for seven of eight cell death modes (apoptosis, autophagy-related death, entotic cell death, ferroptosis, lysosome-dependent cell death, necroptosis, and pyroptosis; all p < 0.05); cuproptosis showed no significant change (Fig 3A–3H). Critically, pyroptosis exhibited the highest activation score among all eight modalities (Fig 3I), consistent with the predominant enrichment of the NLR/inflammasome pathway identified in KEGG analysis. Pairwise comparison confirmed that the pyroptosis score was significantly higher than each of the other seven cell death mode scores (Fig 3J, all comparisons p < 0.05).
(A–H) ssGSEA signature scores for eight cell death modes comparing CTX vs. Control (Wilcoxon test; ns: not significant; *p < 0.05; **p < 0.01). (I) Comparison of all eight cell death signature scores in the CTX group; pyroptosis highlighted in red. (J) Pairwise comparisons of pyroptosis score versus each other modality score.
3.4. Pyroptosis is rapidly activated and sustained across the observation period
To characterize the temporal dynamics of pyroptosis activation, ssGSEA scores were computed separately for each time point. The pyroptosis score was significantly elevated as early as 6 h post-CTX injection and remained significantly elevated at 12 h and 24 h (Fig 4A). Importantly, the magnitude of pyroptosis activation did not differ significantly between the 6 h, 12 h, and 24 h time points (Fig 4B), indicating that CTX triggers near-maximal pyroptosis pathway activation within the first 6 h, with sustained engagement thereafter. This rapid-onset, persistent activation pattern is consistent with CTX’s known mechanism of direct membrane perturbation creating an acute danger signal that continuously drives inflammasome assembly.
(A) Pyroptosis ssGSEA score at Control, 6 h, 12 h, and 24 h. (B) Inter-timepoint comparisons of pyroptosis scores (ns at all comparisons, indicating rapid onset with sustained plateau).
3.5. Core NLR signaling pathway genes are continuously upregulated following CTX exposure
Targeted analysis of canonical pyroptosis pathway components identified six genes—Nlrp3, Pycard (ASC), Gsdmd, Il18, Nfkb, and Tlr4—as the most significantly upregulated canonical NLRP3/caspase-1/GSDMD pathway components across all CTX time points versus healthy controls (Fig 5A– 5F). All six genes showed statistically significant upregulation at each individual time point (6 h, 12 h, and 24 h versus Control; Figs 6–8), confirming sustained transcriptional activation throughout the observation window. These six genes constitute the canonical NLRP3 inflammasome assembly-to-execution axis: NLRP3 (sensor), PYCARD/ASC (adaptor), CASP1 (effector protease), GSDMD (executor), and IL-1β/IL-18 (inflammatory outputs).
(A–F) Violin plots with individual data points showing expression of *Nlrp3*, *Pycard*, *Gsdmd*, *Il18*, *Nfkb*, and *Tlr4* in CTX (pooled 6/12/24 h) vs. Control groups.
(A–F) Violin plots with individual data points showing expression of *Nlrp3*, *Pycard*, *Gsdmd*, *Il18*, *Nfkb*, and *Tlr4* in the CTX 6 h group versus Control.
(A–F) Violin plots with individual data points showing expression of *Nlrp3*, *Pycard*, *Gsdmd*, *Il18*, *Nfkb*, and *Tlr4* in the CTX 12 h group versus Control.
(A–F) Violin plots with individual data points showing expression of *Nlrp3*, *Pycard*, *Gsdmd*, *Il18*, *Nfkb*, and *Tlr4* in the CTX 24 h group versus Control.
3.6. GSDMD cleavage is confirmed as direct evidence of pyroptosis execution
Western blot analysis of CTX-treated skin tissues demonstrated significant upregulation of NLRP3, phospho-P65 (P-P65; with stable total P65, confirming NF-κB pathway activation rather than overall protein induction), ASC, TLR4, and IL-18, as well as the appearance of the GSDMD N-terminal fragment (GSDMD-N, ~ 31 kDa) — direct evidence of caspase-1-mediated GSDMD cleavage (Fig 9A). Densitometric quantification normalized to β-actin revealed the following patterns (Fig 9B): NLRP3 showed modest upregulation at 6 h (1.11-fold) that was sustained through 24 h; P-P65 showed early peak activation at 6 h (1.27-fold, p < 0.05) with partial recovery at 12 h and renewed elevation at 24 h (1.14-fold, p < 0.05), while total P65 protein remained stable across all time points (n.s.), confirming that the observed increase in P-P65 reflects NF-κB phosphorylation rather than protein-level induction; ASC was significantly upregulated at 6 h (1.34-fold, p < 0.01) and 24 h (1.17-fold, p < 0.05). Among the downstream effectors, TLR4 showed progressive upregulation from 6 h (1.46-fold, p < 0.01) to 12 h (1.61-fold, p < 0.001) and 24 h (1.58-fold, p < 0.01); GSDMD-N was consistently elevated across all CTX timepoints (1.47–1.52-fold, p < 0.01), confirming sustained pyroptotic execution; and IL-18 exhibited the most robust upregulation (1.82–1.90-fold, p < 0.001 at all timepoints), consistent with ongoing inflammasome-driven cytokine processing and release. These findings provide protein-level evidence that CTX activates not only NLRP3 inflammasome assembly but also the downstream GSDMD cleavage step that commits cells to lytic pyroptotic death.
(A) Representative Western blot images. Left panel (Gel 1): NLRP3 (118 kDa), phospho-P65 (P-P65, 65 kDa), total P65 (65 kDa), ASC (24 kDa), and β-actin (42 kDa, loading control). Right panel (Gel 2): TLR4 (75 kDa), GSDMD-N (N-terminal fragment, ~31 kDa), IL-18 (18 kDa), and β-actin (42 kDa). Lanes: Control, CTX-6 h, CTX-12 h, CTX-24 h. (B) Densitometric quantification of all seven proteins normalized to β-actin and expressed relative to control (=1.0). Note: total P65 protein levels were stable across all timepoints (n.s.), confirming that the observed P-P65 increase reflects NF-κB pathway phosphorylation. Data represent mean ± SD of n = 3 independent biological replicates. *p < 0.05; **p < 0.01; ***p < 0.001 vs. Control.
3.7. In Vitro characterization of CTX-Induced Pyroptosis in HaCaT Keratinocytes
To dissect the molecular mechanism of CTX-induced pyroptosis in a controlled cellular system, we established a CTX dose-response model in HaCaT keratinocytes. CCK-8 assay demonstrated concentration- and time-dependent cytotoxicity; treatment at 10 μg/mL for 8 h resulted in approximately 69.5 ± 1.1% cell viability and was selected as the standard treatment condition for subsequent mechanistic experiments (Fig 10A).
(A) CCK-8 dose-response (0–20 μg/mL) and time-course (2, 4, and 8 h) cell viability curves. (B) Optical microscopy of Control, CTX-treated (10 μg/mL, 8 h), and LPS + ATP-stimulated cells (×100). Arrowheads indicate membrane blebbing and cell swelling. (C) Transmission electron microscopy (TEM) images of Control and CTX-treated cells; arrows indicate pyroptotic bodies (1–5 μm diameter; scale bar: 1 μm). (D) PI/DAPI dual staining showing membrane permeabilization. Each condition displayed as three fluorescence channels: Merge, PI (red; membrane-compromised cells), and DAPI (blue; all nuclei). Upper row: Control; lower row: CTX-treated (10 μg/mL, 8 h). Scale bar: 20 μm. All data: mean ± SD; n = 3 independent experiments; *p < 0.05, **p < 0.01, ***p < 0.001 vs. Control.
Morphological features of pyroptosis: Optical microscopy revealed that CTX-treated HaCaT cells (10 μg/mL, 8 h) displayed characteristic pyroptotic morphology including cell swelling, cytoplasmic vacuolation, and membrane blebbing—changes qualitatively similar to those observed in LPS + ATP-stimulated positive control cells (Fig 10B). TEM of CTX-treated cells revealed pyroptotic bodies (1–5 μm membrane-bound vesicles with condensed nuclear content) not present in control cells (Fig 10C), providing ultrastructural evidence of pyroptotic execution.
Membrane permeabilization: PI/DAPI dual staining demonstrated concentration- and time-dependent membrane permeabilization in CTX-treated cells (Fig 10D).
Pyroptosis pathway activation:Western blot analysis confirmed that CTX dose-dependently upregulated NLRP3 protein levels (1.64-fold at 10 μg/mL, p < 0.01) and full-length GSDMD (F-GSDMD), while simultaneously inducing accumulation of the GSDMD N-terminal cleavage fragment (N-GSDMD; 2.18-fold at 10 μg/mL, 3.82-fold at 15 μg/mL, all p < 0.001 vs. control), providing direct protein-level evidence of progressive pyroptotic execution in a dose-dependent manner (Fig 11A–11B). Cleaved caspase-1 was undetectable in untreated cells but emerged prominently at 10 and 15 μg/mL CTX (2.53- and 2.79-fold, p < 0.001), confirming caspase-1 activation as an upstream requirement for GSDMD cleavage. IL-1β secretion into conditioned medium was significantly elevated (Fig 11C), consistent with pore-dependent cytokine release.
(A) Representative Western blot images showing dose-dependent upregulation of NLRP3 inflammasome pathway components in HaCaT keratinocytes treated with increasing concentrations of CTX (0, 5, 10, and 15 μg/mL) for 8 h. Protein bands shown: NLRP3 (118 kDa), full-length GSDMD (F-GSDMD, 55 kDa), GSDMD N-terminal fragment (N-GSDMD, 33 kDa), cleaved caspase-1 (Casp-1, 17 kDa), and β-actin (42 kDa, loading control). (B) Densitometric quantification of NLRP3, F-GSDMD, N-GSDMD, and cleaved Casp-1 protein bands normalized to β-actin and expressed relative to untreated control (= 1.0). Note: cleaved Casp-1 was undetectable in untreated control cells, consistent with caspase-1 activation being an inflammasome-dependent event. N-GSDMD showed the most dramatic dose-dependent increase (3.82-fold at 15 μg/mL, p < 0.001 vs. control), directly evidencing progressive GSDMD cleavage at higher CTX doses. Data represent mean ± SD; n = 3 independent biological replicates. *p < 0.05; **p < 0.01; ***p < 0.001 vs. Control (one-way ANOVA with Tukey’s post hoc test). (C) IL-1β secretion (pg/mL) into conditioned medium measured by ELISA in Control vs. CTX-treated (10 μg/mL, 8 h) HaCaT cells. CTX induced a 5.5-fold increase in IL-1β secretion (p < 0.01), consistent with NLRP3 inflammasome-driven caspase-1-mediated pro-IL-1β cleavage and pore-dependent cytokine release. Data represent mean ± SD; n = 3 independent experiments. **p < 0.01 (Student’s t-test).
3.8. CTX induces intracellular ros accumulation in vitro, serving as the upstream activator of the NLRP3 inflammasome
CTX treatment induced dose-dependent intracellular ROS accumulation in HaCaT keratinocytes, as quantified by DCFH-DA fluorescence (Fig 12A-12B). ROS levels were significantly elevated at all tested CTX concentrations (5–20 μg/mL) compared to untreated controls (all p < 0.05). Furthermore, intracellular ROS accumulation showed a strong positive correlation with CTX-induced cellular cytotoxicity as measured by LDH release (Pearson r = 0.81, p < 0.01; Fig 12C), supporting a mechanistic link between oxidative stress and membrane damage. These findings indicate that CTX-mediated membrane perturbation generates a burst of reactive oxygen species that serves as the upstream danger signal initiating NLRP3 inflammasome assembly. Intracellular ROS elevation supports a mechanistic sequence in which CTX-induced oxidative stress serves as an upstream danger signal priming the NLRP3 inflammasome in exposed keratinocytes, consistent with the established TXNIP–NLRP3 interaction mechanism [29,30].
(A) Representative fluorescence microscopy images showing intracellular ROS levels detected by DCFH-DA staining in HaCaT keratinocytes. Each condition is displayed as three channels: Merge (DCFH-DA signal overlaid with DAPI), ROS (DCFH-DA channel; excitation/emission: 485/530 nm; pseudo-colored red), and DAPI (nuclear counterstain; blue). Upper row: untreated control cells, showing minimal background ROS fluorescence. Lower row: CTX-treated cells (10 μg/mL, 8 h), showing markedly elevated DCFH-DA signal distributed throughout the cytoplasm. Scale bar: 20 μm. (B) Quantification of intracellular ROS fluorescence intensity in HaCaT keratinocytes treated with increasing concentrations of CTX (0, 5, 10, 15, and 20 μg/mL) for 8 h. ROS levels were significantly elevated at all tested concentrations compared to untreated controls (all p < 0.05). Data represent mean ± SD; n = 3 independent experiments. *p < 0.05; **p < 0.01; ***p < 0.001 vs. Control (one-way ANOVA with Tukey’s post hoc test). (C) Positive correlation between intracellular ROS fluorescence intensity and CTX-induced cellular cytotoxicity (LDH release, %) across all treatment conditions. Each data point represents one treatment group (mean value). Pearson r = 0.81, p < 0.01. The linear regression line with 95% confidence interval is shown.
3.9. Pharmacological Validation of the ROS–NLRP3–Caspase-1–GSDMD Axis In Vitro
To confirm the mechanistic hierarchy of the ROS → NLRP3 → caspase-1 → GSDMD axis in mediating CTX-induced pyroptosis, we employed selective pharmacological inhibitors targeting distinct nodes: NAC (ROS scavenger, upstream signal), MCC950 (NLRP3 inhibitor), and VX-765 (caspase-1 inhibitor).
MCC950 (NLRP3 inhibitor): Pre-treatment with MCC950 (10 μM, 1 h before CTX) suppressed cleaved caspase-1, GSDMD-N, and also reduced NLRP3 protein levels (Fig 13A), consistent with MCC950 disrupting the upstream NLRP3 sensor itself. This contrasts with VX-765 (caspase-1 inhibitor), which showed compensatory NLRP3 upregulation, suggesting that active caspase-1 normally provides a negative feedback signal to suppress NLRP3 transcription. MCC950 significantly reduced LDH release (from 9.7% to 5.5%, p < 0.0001; Fig 13B) and IL-1β secretion (from 80.7 ± 5.4 to 53.0 ± 4.3 pg/mL, p < 0.0001; Fig 13C). Collectively, these results demonstrate that NLRP3 blockade by MCC950 significantly attenuates CTX-induced LDH release and IL-1β secretion, confirming the causal role of NLRP3 in mediating CTX-induced pyroptosis.
MCC950 (NLRP3 inhibitor): (A) Western blot of Control, CTX, and CTX + MCC950 groups showing unchanged NLRP3 (consistent with post-translational inhibition), suppressed cleaved caspase-1, and suppressed GSDMD-N. (B) LDH release (%) in CTX vs. CTX + MCC950 (****p < 0.0001). (C) IL-1β secretion (pg/mL) in CTX vs. CTX + MCC950 (****p < 0.0001). VX-765 (caspase-1 inhibitor): (D) Western blot of Control, CTX, and CTX + VX-765 groups; note compensatory NLRP3 upregulation, suppressed cleaved caspase-1 and GSDMD-N. (E) LDH release (%) in CTX vs. CTX + VX-765 (****p < 0.0001). (F) IL-1β secretion (pg/mL) in CTX vs. CTX + VX-765 (****p < 0.0001).
VX-765 (caspase-1 inhibitor): Pre-treatment with VX-765 (20 μM, 1 h before CTX) markedly suppressed cleaved caspase-1 and GSDMD-N (Fig 13D). Notably, NLRP3 protein expression was increased in VX-765-treated cells, consistent with a compensatory transcriptional upregulation of NLRP3 when downstream caspase-1-mediated cleavage products are absent; this likely reflects a caspase-1-to-NF-κB feedback loop that normally suppresses NLRP3 transcription under active inflammasome conditions. VX-765 similarly suppressed LDH release (Fig 13E) and IL-1β secretion (Fig 13F) to levels comparable to MCC950, demonstrating that downstream caspase-1 activity is required for CTX-induced membrane permeabilization, cytokine release, and cell death.
NAC (ROS scavenger, upstream inhibitor): Pre-treatment with N-acetylcysteine (NAC, 5 mM, 1 h) significantly attenuated CTX-induced intracellular ROS accumulation and partially restored cell viability in CTX-treated cells (Fig 14E, p < 0.001 vs. CTX alone). NAC also suppressed downstream LDH release and IL-1β secretion, confirming that ROS generation is required for full activation of the NLRP3 → caspase-1 → GSDMD cascade (Fig 14A-14D). These results establish ROS accumulation as the upstream initiating event in the CTX-induced pyroptotic pathway.
(A) Representative western blot images showing the expression of NLRP3, full-length GSDMD (F-GSDMD), N-terminal GSDMD (N-GSDMD), and cleaved caspase-1 in Control, NAC, CTX, and NAC + CTX groups, with β-actin used as the loading control. (B) Densitometric quantification of NLRP3, F-GSDMD, N-GSDMD, and cleaved caspase-1 protein levels normalized to β-actin. (C) LDH release assay showing cell membrane damage in each group. (D) ELISA analysis of IL-1β secretion in the culture supernatant. NAC pretreatment markedly attenuated CTX-induced activation of the pyroptosis-related signaling cascade, as evidenced by reduced N-GSDMD and cleaved caspase-1 expression, accompanied by decreased LDH release and IL-1β secretion. (E) CCK-8 cell viability (%) in Control, CTX, and CTX + NAC groups; NAC partially restored viability (***p < 0.001 vs. CTX). Data are presented as mean ± SD from three independent experiments. ns, not significant; *P < 0.05; ****P < 0.0001.
Taken together, these results establish that CTX activates intracellular ROS accumulation as the upstream initiating signal, which primes NLRP3 inflammasome assembly and drives the caspase-1 → GSDMD cascade as the dominant mechanism of pyroptotic cell death in CTX-exposed keratinocytes. The mechanistic hierarchy is: CTX → ROS↑ → NLRP3 activation → caspase-1 cleavage → GSDMD-N pore formation → pyroptotic death and IL-1β/IL-18 release.
4. Discussion
Our study provides comprehensive transcriptomic and mechanistic evidence that pyroptosis, specifically the ROS → NLRP3 → caspase-1 → GSDMD axis, is a key pathway driving CTX-induced skin tissue injury.
Pyroptosis as one of the mechanistically important cell death modalities. Previous studies have established that cobra CTX and related cytotoxins can trigger diverse forms of cell death—including mitochondria-mediated apoptosis, endoplasmic reticulum stress-dependent apoptosis, autophagy, necrosis of skeletal muscle, and necroptosis in leukemia cells [23,24]. Our genome-wide ssGSEA analysis confirmed that multiple cell death pathways are activated following CTX exposure; however, pyroptosis showed the highest activation score among all eight assessed modalities in the in vivo skin tissue transcriptomic context, corroborated by predominant KEGG enrichment of the NOD-like receptor signaling pathway. It should be noted that this ssGSEA ranking reflects the composite transcriptomic signature of intact tissue—which encompasses inflammasome-competent tissue-resident immune cells (macrophages, monocytes, dendritic cells) that amplify the NLR/pyroptosis gene expression program—and may differ from findings in isolated cell line systems. Prior in vitro studies have identified necroptosis and apoptosis as prominent CTX-induced death modes in pure cell line models [25,26], and these pathways are also significantly activated in our transcriptomic dataset. Importantly, in our in vitro functional validation experiments, pharmacological inhibition of the NLRP3/caspase-1 axis with selective inhibitors (MCC950 and VX-765) substantially reduced LDH release, IL-1β secretion, and GSDMD cleavage in CTX-exposed HaCaT keratinocytes, establishing that pyroptosis constitutes an important, functionally significant contributor to CTX-induced epithelial injury, coexisting with other activated cell death pathways. This finding is consistent with prior work in hepatic ischemia-reperfusion injury and cardiac pyroptosis models demonstrating that GSDMD-mediated membrane pore formation amplifies sterile tissue damage through cytokine release and DAMP dissemination, exceeding the primary cytotoxic insult [27,28].
Mechanism of CTX-mediated NLRP3 inflammasome activation. Our data indicate that CTX-induced intracellular ROS accumulation serves as the likely upstream activator of the NLRP3 inflammasome, given that intracellular ROS levels were markedly elevated in CTX-treated HaCaT cells in a dose-dependent manner. This is consistent with established mechanisms by which mitochondrial ROS, oxidized mitochondrial DNA, and disruption of mitochondrial membrane potential collectively drive NLRP3 inflammasome priming and assembly [29–31]. Specifically, mitochondria-derived reactive oxygen species trigger thioredoxin-interacting protein (TXNIP) dissociation, enabling TXNIP–NLRP3 interaction and inflammasome oligomerization; oxidized mitochondrial DNA released from damaged mitochondria provides an additional endogenous DAMP that sustains NLRP3 activation [32]. We propose that CTX-mediated membrane perturbation, by inserting into the bilayer and disrupting mitochondrial membrane potential, generates a burst of reactive oxygen species that provides both the danger signal (DAMPs) and the metabolic context necessary for NLRP3 oligomerization. The subsequent caspase-1-mediated cleavage of GSDMD creates transmembrane pores that amplify the initial injury by releasing mature IL-1β, IL-18, and cytosolic DAMPs, establishing a self-sustaining feed-forward inflammatory cascade that drives progressive tissue necrosis.
Distinction from classical infectious inflammasome activation. It is important to note that CTX is not an infectious pathogen; rather, it is a non-replicating chemical toxin with direct membrane-disrupting activity [23]. CTX’s capacity to activate NLRP3 in the absence of pathogen-associated molecular patterns (PAMPs) suggests it acts as a powerful damage-associated molecular pattern (DAMP)-generating stimulus, activating the so-called “sterile inflammation” arm of the NLRP3 pathway. This distinguishes our findings from canonical infectious inflammasome activation but places them within the growing literature on toxin-induced sterile pyroptosis. Indeed, NLRP3/GSDMD-mediated pyroptosis has now been documented in response to diverse non-microbial danger signals—including snake venom Lys49-phospholipase A2 homologues, arthropod venom components, mycotoxins such as T-2 toxin, and crystalline substances such as uric acid crystals and silica—highlighting a broadly conserved role for the NLRP3 inflammasome as a cellular sensor of membrane damage and cytotoxic stress that is not restricted to pathogen recognition [33–35].
Time course and biological interpretation. The rapid onset of pyroptosis (detectable at 6 h) with sustained activation through 24 h—without significant inter-timepoint differences in most genes—reflects the membrane-disrupting mechanism of CTX. Unlike pattern recognition receptor-mediated infections that require intracellular replication cycles to reach threshold PAMP concentrations, CTX can directly perturb cell membranes at the moment of exposure, rapidly releasing DAMPs and activating NLRP3 with near-immediate kinetics. The observation that Gsdmd shows a significant increase at 12 h relative to 6 h likely reflects progressive recruitment of inflammasome-competent cells (including infiltrating macrophages and monocytes) into the injury site during the secondary inflammatory wave, rather than a cell-intrinsic escalation of the pyroptotic program.
Cross-species and cell model considerations. The combination of mouse in vivo and human HaCaT keratinocyte in vitro models in this study is intentional. The in vivo mouse model provides the physiologically relevant tissue context and injury endpoint (necrosis area) for assessing the functional consequences of CTX-induced signaling. The HaCaT in vitro model enables controlled, mechanistic interrogation of the NLRP3/GSDMD cascade in the predominant cell type of the primary injury site (keratinocytes) without the confounding influence of infiltrating immune cells. We acknowledge that HaCaT cells, as a transformed cell line, may have altered inflammasome responsiveness relative to primary human keratinocytes; the pharmacological inhibitor results nonetheless clearly demonstrate the cell-autonomous capacity of keratinocytes to activate caspase-1-dependent GSDMD cleavage in response to CTX. Importantly, human keratinocytes—including HaCaT cells—have recently been confirmed to execute GSDMD-dependent pyroptosis in response to diverse exogenous stimuli including inflammatory cytokines and xenobiotic nanoparticles [36,37], providing independent evidence that the NLRP3/GSDMD axis is fully operative in this cell type and supporting the validity of our in vitro model system. We also note that NLR signaling pathway components are conserved between mouse and human; the mouse transcriptomic data therefore provide a valid basis for pathway-level interpretation.
Therapeutic implications. Our in vitro data identify three experimentally validated pharmacological nodes in the CTX-induced pyroptotic cascade: (1) upstream ROS scavenging (N-acetylcysteine, NAC), (2) NLRP3 inflammasome inhibition (MCC950 and related NLRP3 inhibitors), and (3) downstream caspase-1 blockade (VX-765 and related inhibitors). MCC950 achieves NLRP3 blockade by engaging the ATP-hydrolysis motif within the NACHT domain, preventing conformational changes required for oligomerization [38,39]. In our model, MCC950 additionally reduced NLRP3 protein expression, which may reflect disruption of a caspase-1/NF-κB-dependent feed-forward loop that sustains NLRP3 transcription under active inflammasome conditions; this is consistent with published evidence that NF-κB signaling provides the ‘Signal 1’ priming required for NLRP3 upregulation [39]. These findings suggest that the NLRP3-caspase-1-GSDMD axis represents a tractable therapeutic target in CTX-induced skin injury, pending in vivo validation. Consistent with its role as an upstream activator, pharmacological scavenging of ROS with N-acetylcysteine (NAC) significantly attenuated CTX-induced LDH release, IL-1β secretion, and GSDMD cleavage (Fig 14), confirming that ROS accumulation is required for full NLRP3 inflammasome activation in this model. Thus, in addition to direct NLRP3 and caspase-1 blockade, our data identify NAC as a functionally validated ROS-scavenging agent that interrupts the pyroptotic cascade at its earliest initiating step. While these in vitro findings are robust, further in vivo studies are needed to evaluate the therapeutic potential of NAC, particularly given its established clinical safety profile and low cost, which would facilitate translation for snakebite management. Future studies should evaluate these agents in clinically relevant post-bite treatment protocols using optimized dosing regimens and administration routes.
Limitations.This study has several limitations. First, pharmacological validation of the NLRP3/caspase-1/GSDMD axis was performed exclusively in vitro (MCC950, VX-765); dedicated in vivo pharmacological intervention experiments are required to establish the causal contribution of this pathway to tissue necrosis in intact animal models and to assess therapeutic feasibility in clinically relevant post-bite treatment scenarios. Second, HaCaT cells lack the full complement of innate immune signaling components present in primary skin keratinocytes and tissue-resident immune cells; primary keratinocyte or ex vivo skin explant models would strengthen the translational relevance. Third, although we have identified NLRP3/caspase-1/GSDMD as the dominant pyroptosis axis, the contribution of non-canonical pyroptosis pathways (caspase-4/5/11-mediated GSDMD cleavage [40]) was not directly assessed; future work should also examine whether secondary wound contamination provides LPS-derived triggers for non-canonical inflammasome engagement.
5. Conclusion
This study demonstrates that CTX from Naja atra venom activates the NLRP3/caspase-1/GSDMD pyroptotic cascade as a key mechanism driving skin tissue injury. Transcriptomic profiling identified pyroptosis as the dominant cell death modality activated by CTX in vivo, with sustained upregulation of core pathway genes (Nlrp3, Pycard, Gsdmd, Il18) across 6–24 h. Pharmacological inhibition in HaCaT keratinocytes established the hierarchical cascade: ROS accumulation → NLRP3 inflammasome assembly → caspase-1 activation → GSDMD cleavage → pyroptotic death and IL-1β release. Notably, each of the three experimentally validated inhibitors—NAC (ROS scavenger), MCC950 (NLRP3 inhibitor), and VX-765 (caspase-1 inhibitor)—significantly attenuated CTX-induced pyroptosis, identifying multiple actionable nodes in this pathway. These findings establish the ROS–NLRP3–caspase-1–GSDMD axis as a high-priority therapeutic target for mitigating local tissue necrosis following Naja atra envenomation.
Supporting information
S1 Raw Images. Original uncropped Western blot images corresponding to the cropped immunoblots presented in Fig 9, Fig 11, Fig 13, and Fig 14.
https://doi.org/10.1371/journal.pntd.0014495.s001
(PDF)
Acknowledgments
The authors thank all employees of The First Affiliated Hospital of Guangxi Medical University and Guilin Municipal Hospital of Traditional Chinese Medicine for their helpful discussions related to this project. The authors also appreciate the help of the reviewers in significantly improving the manuscript.
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