Rhaponticin alleviates pancreatic acinar cell necrosis by attenuating oxidative stress via modulation of the HIF-1α signaling pathway.

Acute pancreatitis (AP) is a clinical inflammatory disease of the pancreas with diverse pathogenic factors, primarily induced by cholelithiasis, chronic alcohol abuse, hyperlipidemia, pharmacological agents, or infectious pathogens. Pathologically, the disease is initiated by premature intracellular activation of trypsin within the pancreas, leading to autodigestion of pancreatic tissue, followed by edema, hemorrhage, and necrosis. These pathological changes further trigger a cascade of local and systemic inflammatory responses [ 1–3 ]. The clinical progression of AP may be rapid, and severe AP poses a substantial risk to survival, as evidenced by a mortality rate of 15–30% [ 4 , 5 ]. Studies have indicated that pancreatic acinar cell injury or death constitutes a pivotal initiating event in AP pathogenesis [ 6 ]. The degree of acinar cell damage is closely related to disease severity and clinical prognosis. Preclinical studies demonstrate that improved clinical outcomes are achieved by suppressing pancreatic acinar cell death in initial phases, supporting the pivotal role of this process in AP progression [ 7 ]. Consequently, therapeutic strategies focused on protecting acinar cells are considered to be crucial for blocking the AP development, and have consequently emerged as a major focus of research on the pathological mechanisms of AP and targeted drug discovery. Pancreatic acinar cells serve as the fundamental functional units responsible for exocrine secretion in the pancreas. The specific form of acinar cell demise and its subsequent inflammatory injury are critical determinants of AP progression and patient prognosis from the outset [ 8 ]. Without timely and effective treatment, these pathological processes may escalate into severe complications, including extensive pancreatic destruction and systemic inflammatory response syndrome [ 9 , 10 ]. Acinar cell death is manifested through various pathways, including NLRP3 inflammasome-mediated pyroptosis [ 11 ], ferroptosis driven by iron-dependent lipid peroxide accumulation [ 12–14 ], and necroptosis—a recently highlighted mechanism in inflammation and immune-related disease researches [ 15–17 ]. In light of these complex mechanisms, this study aims to identify and validate therapeutic strategies capable of effectively inhibiting acinar cell death and promoting cellular repair in AP. Rhaponticin (Rha), a natural stilbene glycoside derived from rhubarb rhizomes, has diverse activities such as anti-inflammation, anti-microbe, free-radical scavenging, and antitumor [ 18 , 19 ]. Growing evidence indicates that Rha can significantly mitigates tissue damage by suppressing inflammatory pathways in multiple pathological conditions such as colitis [ 20 ], arthritis [ 21 ], and human endothelial cell inflammation [ 22 ], highlighting its potential therapeutic versatility. These collective findings suggest that Rha may possesses pleiotropy and multi-target mechanisms, particularly in inflammation-associated tissue injury. However, a comprehensive assessment of Rha's potential for treating AP is lacking, and the molecular basis for its protective effects is still unclear. In this study, the therapeutic effects of Rha were comprehensively assessed by AP models. And its potential mechanism was further investigated by integrating network pharmacology, molecular docking, and in vitro and in vivo experiments, which provided the scientific basis for the clinical treatment of AP.

In this study, to establish an in vitro injury cell model, primary acinar cells from mice were subjected to CCK stimulation. Relative to controls, the CCK-modeled group showed markedly elevated LDH release. While different doses of Rha could dose-dependently suppress the CCK-triggered LDH release, with the 5 μM concentration exhibiting the most substantial cytoprotective effect ( Figure 1A and B ). Consequently, this optimal dosage was selected for subsequent experiments. Additionally, calcein-AM/PI staining further validated that 5 μM Rha effectively preserved the viability of acinar cells and ameliorated the CCK-induced cellular damage ( Figure 1C and E ). As is well established, acinar cell injury is accompanied by pronounced oxidative stress, with mitochondrial dysfunction recognized as a central event contributing to cellular damage. The mitochondrial membrane potential (MMP) and superoxide generation, which reflect oxidative phosphorylation and electron transport processes, serve as key indicators of mitochondrial function. In healthy cells, JC-1 aggregates and emits red fluorescence, whereas in damaged cells with collapsed mitochondrial membranes, JC-1 fails to accumulate and instead emits green fluorescence. The JC-1 red/green ratio and MitoSOX were employed to determine the MMP and mitochondrial ROS production, respectively. As expected, exposure to CCK significantly increased mitochondrial ROS levels and decreased JC-1 accumulation in acinar cells, while Rha treatment markedly abrogated these changes ( Figure 1D , F–G ). Rha also effectively restored the NAD+/NADH ratio and intracellular ATP levels ( Figure 1H–I ). Furthermore, western blot analysis demonstrated that Rha rescued the CCK-induced suppression of key OXPHOS complex proteins (Complex I-V) ( Figure 1J–K ). Collectively, these findings demonstrated that Rha could significantly alleviate the CCK-induced acinar cell injury under in vitro conditions. Rha treatment attenuated caerulein-induced pancreatic injury in a mouse model of AP. (A) Schematic representation of the in vivo experimental protocol. (B) Representative H&E-stained sections of pancreatic tissue at 100 ×  and 400 ×  magnification. Scale bars: 200 μm (100×), 50 μm (400×). Yellow arrows indicate areas of pancreatic injury, including necrosis, edema, and inflammatory infiltration. (C-D) Serum concentrations of amylase and lipase. Figure 2D was analyzed using non-parametric tests (Kruskal-Wallis test). (E–H) Pathological score of pancreatic tissue. (I-J) Pancreatic tissue amylase (red) and DAPI (dark blue) immunofluorescence. Yellow arrows indicate the abnormal distribution or loss of amylase (red fluorescence) in pancreatic acinar cells, reflecting cellular damage. One-way ANOVA followed by Tukey's post hoc test was used for comparisons among multiple groups, unless otherwise indicated. ns, no significance, * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001. A Cae-induced AP model was employed to further assess the therapeutic potential of Rha ( Figure 2A ). Compared with the control group, pancreatic samples from Cae-treated mice displayed substantial edema, inflammatory infiltration, and focal acinar necrosis. Rha treatment could substantially ameliorate these pathological alterations, with the medium dose (200 mg/kg) demonstrating the most robust protective effect, surpassing the high-dose (400 mg/kg) and low-dose (100 mg/kg) regimens ( Figure 2B and E–H ). Additionally, serum amylase and lipase concentrations were markedly increased in Cae-treated mice relative to control animals. However, Rha administration markedly reduced these enzymatic indicators, with the 200 mg/kg dose exhibiting the most pronounced normalization effect ( Figure 2C and D ). Based on these comprehensive results, the medium dose of Rha was identified as optimal and selected for subsequent mechanistic studies. Immunofluorescence staining using an anti-amylase antibody further confirmed the protective effect of Rha. While amylase expression was notably reduced in pancreatic tissues of the Cae group compared to the NC group, this reduction was effectively reversed following Rha intervention ( Figure 2I–J ). The above results indicated that Rha could effectively improve AP, highlighting its considerable therapeutic potential. To evaluate the therapeutic generalizability of Rha across AP induced by different etiologies, we further examined its protective effects in an alcohol-related AP model (Fig. S1A) and NaT-AP models (Fig. S2A). Histopathological analysis showed that both models exhibited typical features of pancreatic injury, including acinar cell necrosis, inflammatory cell infiltration, and interstitial edema. Notably, Rha treatment markedly alleviated these pathological changes (Fig. S1B,E–H and Fig. S2 B,E–H). Furthermore, serum biochemical assays indicated that Rha significantly reduced the levels of serum amylase and lipase compared to the respective model groups (Fig. S1C-D and Fig. S2C-D). Taken together, the consistent protective effects of Rha across Cae-induced, alcohol-related, and NaT-induced AP models suggest that Rha may exert beneficial effects across distinct etiologies of AP and holds potential as a therapeutic candidate. To assess pancreatic inflammatory cell infiltration, serum levels of key pro-inflammatory cytokines (IL-1β, IL-6, TNF- α ) were determined. Rha administration markedly lowered these cytokine levels versus the Cae group, showing the most pronounced effect at the 200 mg/kg dose ( Figure 3A–C ). The suppression of systemic inflammation correlated well with the amelioration of pancreatic acinar cell injury. Immunostaining for F4/80 and MPO showed that Rha markedly reduced pancreatic infiltration of macrophages and neutrophils compared to the Cae group ( Figure 3D-E ). Thus, Rha alleviated Cae-induced AP through effective suppression of inflammatory cell recruitment and cytokine production. Rha inhibited inflammatory cell infiltration in pancreatic tissue. (A–C) Serum levels of IL-1β, TNF- α and IL-6 were measured by ELISA ( n  = 6). Figure 3B was analyzed using non-parametric tests (Kruskal-Wallis test). (D) F4/80 immunohistochemistry of pancreatic tissue. Red arrows indicate F4/80-positive macrophage infiltration in the pancreatic interstitial spaces. (E) MPO immunohistochemistry of pancreatic tissue ( n  = 6); scar bar: 100×/200 μm, 400/50 μm. Red arrows indicate MPO-positive neutrophil infiltration in the pancreatic tissue. One-way ANOVA followed by Tukey's post hoc test was used for comparisons among multiple groups. * p  < 0.05, *** p  < 0.001, **** p  < 0.0001. Target screening identified 674 Rha-related targets, 10,283 AP-associated targets, and 2185 cell death-related targets. Intersection analysis revealed 222 overlapping core targets among these datasets ( Figure 4A ). Subsequent KEGG pathway assessment showed notable enrichment for the HIF-1α signaling axis within these overlapping targets when ranked by statistical significance ( Figure 4B ). The PPI network built from the STRING database contained 217 nodes and 4,238 interactions, averaging 39.1 connections per node. Network topology analysis identified HIF-1α as a central hub within the network architecture ( Figure 4C ). Based on existing guidelines [ 24 , 25 ], molecular binding strength was classified into strong (<−6.0 kcal/mol), moderate (−6.0 to −5.0 kcal/mol), and weak (−5.0 to −4.0 kcal/mol) categories. Molecular docking revealed a binding energy of −7.7 kcal/mol between Rha and HIF-1α, demonstrating robust binding ( Figure 4D ). These results suggested that Rha might function as an antagonist of the HIF-1α signaling pathway through direct binding to HIF-1α. Network pharmacology analysis identified HIF-1α as a key target of Rha in alleviating AP. (A) Overlapping Venn diagram of AP-related targets, Rha-related targets and cell death-related targets. (B) KEGG pathway enrichment map. (C) AP-Rha-cell death' network diagram. (D) Molecular docking results of Rha and HIF-1α. To examine the effect of Rha on HIF-1α during pancreatitis, HIF-1α protein levels in pancreatic tissue were analyzed by western blot. The results demonstrated significant upregulation of HIF-1α and necroptosis markers (RIP3 and p -MLKL) in the Cae-induced AP model, which was markedly suppressed by Rha treatment ( Figure 5A–D ). Immunohistochemical staining further confirmed enhanced expression levels of HIF-1α and p -MLKL in pancreatic tissues of the Cae group, with these increases being substantially attenuated following Rha intervention ( Figure 5E and G-H ). In addition, the model group exhibited a significant increase in 4-HNE expression and a marked decrease in GPX4 expression while treatment with Rha effectively reversed these alterations, demonstrating its potent anti-oxidative stress activity ( Figure 5F and I-J ). Collectively, these results showed that Rha could mitigate the inflammatory pathogenesis of AP mice by inhibiting the HIF-1α signaling pathway. Rha inhibited HIF-1α–mediated necroptosis and oxidative stress in AP. (A) The expression levels of HIF-1α, RIP3 and phosphorylated MLKL ( p -MLKL) in acinar tissues were examined by western blot analysis. (B–D) Relative expression levels of HIF-1α, RIP3 and phospho-MLKL. GAPDH served as the loading control for HIF-1α, RIP3, while total MLKL was used for p -MLKL normalization. (E, G, and H) Immunohistochemistry of HIF1α and p -MLKL in pancreatic tissue. Red arrows indicate the increased expression of HIF-1α and p -MLKL (positive brown staining) in pancreatic acinar cells. (F, I, and J) Immunohistochemistry of 4HNE and GPX4 in pancreatic tissue ( n  = 6) scar bar: 100×/200 μm, 400/50 μm. Red arrows indicate 4HNE-positive areas and regions of GPX4 loss. One-way ANOVA followed by Tukey's post hoc test was used for comparisons among multiple groups. * p  < 0.05, *** p  < 0.001, **** p  < 0.0001. Based on the cytoprotective profile of Rha, rescue experiments employing the HIF-1α inhibitor PX-478 were conducted to delineate its molecular mechanism. To identify the most effective concentration, we conducted a dose-response screening using an LDH release assay in primary pancreatic acinar cells. As shown in Figure 6A , we evaluated a concentration gradient of PX-478 (1.25, 2.5, and 5 μM). Among these, 1.25 μM of PX-478 exhibited the most significant protective effect in reducing CCK-induced cell injury. Higher concentrations did not yield further significant protective benefits. Consequently, 1.25 μM was selected as the optimal concentration for all subsequent cellular mechanistic experiments. Subsequent rescue assessments through LDH quantification and calcein-AM/PI dual staining revealed that Rha provided no additive therapeutic benefits beyond PX-478 monotherapy ( Figure 6B–D ). Furthermore, we successfully established a HIF-1α knockdown model in 266-6 cells using siRNA ( Figure 6E–F ). Consistent with our previous findings, silencing HIF-1α alleviated acinar cell injury. Notably, treatment with Rha in HIF-1α–knockdown cells did not result in additional protective effects ( Figure 6G–H ). These findings suggested that Rha could ameliorate AP through the modulation of the HIF-1α signaling axis, effectively attenuating the damage of pancreatic acinar cells. The HIF-1α inhibitor PX-478 abolished the protective effect of Rha on acinar cells in AP. (A-B) LDH release in acinar cells ( n  = 6). (C-D) The dead cell rate of acinar cells co-incubated with CCK, PX-478 and Rha in live/dead cell staining ( n  = 6). (E) The expression levels of HIF-1α were examined by western blot analysis. (F) Relative expression levels of HIF-1α, GAPDH served as the loading control for HIF-1α. (G-H) The dead cell rate of HIF-1α-knockdown 266-6 cells as determined by live/dead staining ( n  = 6). Yellow arrows indicate PI-positive (dead) cells. Student's t-test was used for comparisons between two groups. One-way ANOVA followed by Tukey's post hoc test was used for comparisons among multiple groups. ns, no significance, * p  < 0.05, ** p  < 0.01, **** p  < 0.0001. Additionally, the effect of PX-478 on the in vivo protection of Rha was further investigated. Shen et al reported that administration of PX-478 at 100 mg/kg provided significant protective effects, effectively reducing acinar cell necrosis in a mouse model of AP [ 26 ]. Based on these findings, we adopted this dosage as an appropriate and evidence-based regimen for our study. In line with previous in vivo findings, pancreatic tissues from the Cae group exhibited obvious edema, inflammatory cell infiltration, and acinar cell necrosis compared with the NC group. Notably, co-administration of Rha with PX-478 did not produce additional therapeutic effects, as evidenced by comparable levels of pancreatic tissue damage and serum amylase/lipase concentrations to PX-478 monotherapy ( Figure 7A–G ). Thus, these results further validated that HIF-1α mediated Rha's pharmacological effects. Furthermore, Rha significantly attenuated TLCS-induced cell injury in human acinar cells, as evidenced by reduced LDH release (Fig. S3A) and decreased ROS accumulation (Fig. S3B-C). These findings are highly consistent with our observations in murine pancreatic acinar cells. The protective effect of Rha was comparable to that of PX478. Notably, co-administration of Rha and PX478 did not produce additional protective effects, suggesting that Rha exerts its therapeutic effects primarily through the HIF-1α pathway in the human cellular context. In summary, these findings in human primary acinar cells confirm that the protective effects of Rha against injury and oxidative stress are largely mediated through the inhibition of the HIF-1α pathway, a mechanism that is highly conserved between murine and human models. HIF-1α inhibitor PX-478 counteracted the protective effect of Rha on Cae-induced AP. (A) Representative HE staining of pancreatic tissue with magnifications of 100 times and 400 times. Scar bar: 100×/200 μm, 400×/50 μm. Yellow arrows indicate areas of pancreatic injury, including necrosis, edema, and inflammatory infiltration. (B-C) Serum levels of amylase and lipase. (D–G) Pathological score of pancreatic tissue. One-way ANOVA followed by Tukey's post hoc test was used for comparisons among multiple groups. ns, no significance, * p  < 0.05, ** p  < 0.01 and **** p  < 0.0001.

Rhaponticin (CAS No. 155-58-8) and Palmitoleic acid (P816247) were obtained from Macklin (Shanghai, China, No. R861350); PX-478 (HY-10231) was purchased from MCE (New Jersey, U.S.A.); Caerulein (Cae) (C9026), dihydroethidium (DHE) (D7008) and Taurocholic acid sodium salt Hydrate (T4009) were bought from Sigma-Aldrich (St. Louis, MO, U.S.A.); Taurolithocholic Acid 3-sulfate (sodium salt)(TLCS,64936-83-0) was purchased from CAYMAN(Ann Arbor, MI, U.S.A.); Cholecystokinin (CCK) (S9690) was provided by Echelon Biosciences (Utah, U.S.A.); Anti-RIP 3 antibody (sc-374639) was purchased from Santa Cruz Biotechnology (CA, U.S.A.); Anti-amylase (ab21156), anti-Hif1α (ab179483), anti-MLKL (ab243142), anti- p -MLKL (ab196436), anti-myeloperoxidase (MPO) (ab208670) and mouse total OXPHOS rodent WB antibody cocktail (ab110413) were obtained from Abcam (Cambridge, UK); Rabbit (7074) and mouse (7076) secondary antibodies were purchased from Cell Signaling Technologies (MA, U.S.A.); LDH Cytotoxicity Assay Kit (C0017), NAD + /NADH Assay Kit (S0175); Enhanced adenosine 5′-triphosphate (ATP) Assay kit (S0027) and BCA Protein Concentration Assay Kit (P0009) were bought from Beyotime Biotechnology (Beijing, China); Calcein-AM/PI double staining kit (40747ES) was purchased from Yeasen Biotechnology (Shanghai, China); Amylase Kit (D08-09) was bought from BioSino BioTechnology & Science Inc. (Beijing, China); Lipase Kit (A054-1-1) was purchased from Nanjing Jiancheng (Nanjing, China); F4/80 rabbit monoclonal antibody ( GB113373 ) was provided by Servicebio (Wuhan, China); DAPI (BL105A) was bought from Biosharp (Beijing, China). Male ICR wild-type mice (6–8 weeks old, 20–25 g) were obtained from the Institute of Comparative Medicine, Yangzhou University; all animals were healthy, immunocompetent, non-genetically modified, and naive to prior experimental procedures, with the study approved by the Institutional Animal Care and Use Committee of Yangzhou University (Approval No. 202509007) and conducted in strict compliance with the NIH Guide for the Care and Use of Laboratory Animals. Mice were housed under specific pathogen-free (SPF) conditions in a controlled environment (20–25 °C, 50–60% relative humidity, 12-h light–dark cycle) and had ad libitum access to sterilized water and standard laboratory chow. Using individual mice as the experimental unit, animals were randomly allocated to groups using a random-number table ( n  = 6 per group). A total of 90 mice were utilized across four independent experiments (Experiment 1: n  = 30; Experiment 2: n  = 24; Experiment 3: n  = 18; Experiment 4: n  = 18). In this study, the sample size of animal experiments is a scheme formulated in combination with the 3 R principle. Randomization was systematically applied to treatment assignment, outcome assessment, and cage placement to minimize potential bias. In the Cae-AP model, healthy ICR mice were randomly divided into a control (NC), a Cae-induced AP model (Cae) (20 μg/mL dissolved in PBS), and different treatment groups receiving Cae and Rha (dissolved in DMSO) (Cae + Rha), Cae and PX-478 (dissolved in PBS) (Cae + PX-478), or the combination of Cae, PX-478, and Rha (Cae + PX-478 + Rha). The NC group received standard handling without intervention. AP in the Cae group was induced by repeated intraperitoneal dosing of Cae (200 μg/kg), administered hourly for a total of eight times as shown in Figure 2A. After the first Cae injection, different treatment groups were administered with Rha at doses of 100, 200, or 400 mg/kg, respectively, while PX-478 (100 mg/kg) was co-administered with Rha as indicated. The NC and Cae groups received an equal volume of saline. At 12 hours post-induction, all mice were anesthetized with isoflurane to achieve deep anesthesia, followed by euthanasia via carbon dioxide inhalation; after confirmation of complete death by a qualified veterinarian, serum samples and pancreatic tissues were collected for subsequent analyzes, and the carcasses were transferred to a designated harmless treatment facility. To establish the alcoholic AP model, mice received two intraperitoneal injections of ethanol (1.35 g/kg) and palmitoleic acid (POA, 150 mg/kg) at a 1-hour interval. Prior to the ethanol+POA injections, 200 μL of normal saline was administered to prevent localized peritoneal organ damage. Control mice were treated with normal saline. In the treatment groups, mice received intraperitoneal injections of Rha simultaneously with the first ethanol and POA challenge. Animals were euthanized 24 hours after the initial injection, following the same sampling and sacrifice procedures described above. In Sodium taurocholate (NaT) in vivomodel, the model was also induced via retrograde infusion of 1% (w/v) NaT into the biliopancreatic duct. The control group (NC) received 0.9% (w/v) NaCl instead. Two hours post-surgery, Rha was administered via intraperitoneal injection. Mice were euthanized 24 hours after the completion of the surgical procedure, and samples were collected as previously specified. Primary outcome measures included serum biochemical markers and histopathological examination of pancreatic tissue, which were used to assess the severity of AP and the therapeutic efficacy of Rha. Rha treatment attenuated mitochondrial damage and enhanced the viability of pancreatic acinar cells in vitro. (A) The physical map and molecular structure of Rha. (B) LDH release in CCK-treated acinar cells. (C) Live/dead cell staining images of acinar cells after co-incubation with CCK and Rha (green: live cells; red: dead cells). Yellow arrows indicate PI-positive (dead) cells. (E) The dead cell rate of acinar cells co-incubated with CCK, Rha in live/dead cell staining ( n  = 6). (D, F-G) Potential alterations in the mitochondrial membrane and mitochondrial ROS levels were detected using JC-1-specific fluorescence and the MitoSOX probe. Yellow arrows indicate cells with decreased mitochondrial membrane potential and elevated mitochondrial ROS levels. (H-I) Intracellular levels of the NAD⁺/NADH ratio and ATP. (J-K) Representative western blot images and corresponding quantitative analysis of mitochondrial OXPHOS system components. One-way ANOVA followed by Tukey's post hoc test was used for comparisons among multiple groups. ns, no significance, * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001. Following 48-hour fixation in 4% paraformaldehyde (4 mL paraformaldehyde + 96 mL ddH₂O), pancreatic tissues underwent routine processing for dehydration and paraffin embedding. For histological evaluation, sections underwent routine H&E staining and microscopic examination. A blinded scoring system was implemented to evaluate pancreatic injury based on four key parameters: edema, inflammatory response, leukocyte infiltration, and acinar cell necrosis. Each parameter was graded on a 0–4 point scale, with the cumulative score representing the overall severity of pancreatic tissue damage. We analyzed the intermediate grades with 0.5 increments to standardize the statistical process. Pancreatic tissue specimens were fixed, paraffin-embedded, and sectioned at 5 μm thickness. After mounting on glass slides, the sections underwent dewaxing, rehydration, antigen retrieval, and blocking of endogenous peroxidase activity using normal goat serum. For immunohistochemical analysis, sections were incubated overnight at 4 °C with primary antibodies—anti-HIF-1α (1:100), anti- p -MLKL (1:100), anti-F4/80 (1:500), and anti-MPO (1:500). Then, the sections were incubated with a biotin-labeled secondary antibody (1:500) for 15 min at ambient temperature. Detection was performed using DAB chromogenic substrate with hematoxylin counterstaining. For immunofluorescence, sections were exposed to anti-amylase (1:75) overnight at 4 °C, rinsed with PBS, and then treated with Cy3-tagged goat anti-mouse IgG for 2 hours at room temperature under light-safe conditions. Following washing, nuclei were counterstained with DAPI according to the manufacturer's protocol. Fluorescent signals were acquired by a microscope with suitable filter sets. Following serum collection from experimental mice, the enzymatic activities of amylase and lipase in the pancreas were measured by corresponding test kits. Subsequently, the concentrations of pivotal inflammatory mediators (IL-1β, IL-6, and TNF- α ) were quantified with standard ELISA kits. Every measurement was carried out in strict adherence to the supplied instructions. Pancreatic acinar cells were extracted from mice with a collagenase-based technique, as previously established [ 23 ]. A cellular injury model was established by treating acinar cells with CCK for 6 hours. Following treatment completion, cellular extracts and culture supernatants were collected for subsequent biochemical and molecular analyzes. Human pancreatic acinar cells (hPACs) were isolated from normal pancreatic tissues distal to benign tumors obtained during partial pancreatectomy. The study was conducted in accordance with the principles of the Declaration of Helsinki and was approved by the Ethics Committee of the Affiliated Hospital of Yangzhou University (Approval No. 2018-YKL11-27-(K3)-ET). Written informed consent was obtained from all donors. Briefly, the tissues were minced and subjected to an optimized Collagenase V digestion at 37 °C, coupled with mechanical dissociation. The resulting cell suspension was then filtered through a 70 μm nylon mesh and collected by centrifugation. Following treatment completion, both murine and human cellular extracts and culture supernatants were collected for subsequent biochemical and molecular analyses. The sequence targeting HIF-1α was designed and synthesized by the company. The sequences were as follows: si-HIF-α (sense: 5’-GAUGGAAGCACUAGACAAATT-3’; antisense: 5’-UUUGUCUAGUGCUUCCAUCTT-3’), si-Negative control (si-NC) (sense:5’-UUCUCCGAACGUGUCACGUTT-3’; antisense: 5’ACGUGACACGUUC GAGAATT-3). siRNAs were transfected into 60-70% confluent 266-6 cells with Lipofectamine 3000 regent. After 6 h of incubation, medium containing transfection reagents was refreshed with complete 266-6 cell medium and the cells were cultured for another 24 h before other experiments. Intracellular reactive oxygen species (ROS) levels in primary acinar cells were determined using a dihydroethidium (DHE) fluorescent probe. Fluorescence images were captured using a Nikon confocal microscope and subsequently quantified using ImageJ software. ATP (Beyotime S0027, Shanghai, China) and NAD + / NADH levels (Beyotime S0175, Shanghai, China) were analyzed according to the manufacturer's protocol. The ratio of NAD + / NADH was calculated by standard curve. Cellular toxicity in acinar cells was determined by quantifying LDH leakage using a commercially available test system. Cell culture supernatants were collected and analyzed for the LDH activity following the manufacturer's guidelines. A total of 5 μL Calcein-AM solution (2 mM) and 15 μL propidium iodide (PI) solution (1.5 mM) were mixed with 5 mL 1 × Assay Buffer. Freshly isolated pancreatic acinar cells were washed three times and subsequently stained with calcein-AM and PI at 4 °C for 15 minutes. After sample preparation, cellular fluorescence was visualized under a fluorescence microscope using an excitation wavelength of 490 nm. Viable cells exhibited green cytoplasmic fluorescence, while non-viable or apoptotic cells displayed distinct red nuclear staining. Protein concentration was determined via a BCA assay following the manufacturer's protocol.​ Equal amounts of protein samples were separated by 10% SDS-polyacrylamide gel electrophoresis, then transferred onto PVDF membranes.​ Membranes were blocked with 5% (w/v) skimmed milk for 2 hours at room temperature, and afterward incubated overnight at 4 °C with specific primary antibodies: anti-HIF-1α (1:1000), anti-GAPDH (1:2000), anti-RIP3 (1:1000), anti-MLKL (1:1000), and anti- p -MLKL (1:1000) and mouse total OXPHOS rodent WB antibody cocktail (1:250).​ The next day, membranes were washed three times with TBST (15 minutes each) and incubated with horseradish peroxidase (HRP)-linked secondary antibody (1:5000) for 2 hours at room temperature.​ After three more TBST washes (10 minutes each), protein bands were visualized by the Tanon ECL Plus chemiluminescence system. Band intensity quantification was conducted via grayscale analysis with ImageJ software. Potential therapeutic targets associated with AP were systematically identified through comprehensive mining of the GeneCards, DisGeNET, and OMIM databases. Putative targets of Rha were retrieved from multiple pharmacological and traditional medicine-oriented databases, including SymMap, Swiss Target Prediction, SuperPred, and TargetNet. Furthermore, targets associated with cell death were screened via the NIH, OMIM, and CTD databases. Common targets shared among these three datasets were identified and visualized via a Venn diagram. Common targets were uploaded to the STRING database (version 11.0; http://string-db.org ) for PPI network construction.​ Biological species were restricted to Homo sapiens, with a minimum interaction confidence cutoff set at 0.9.​ Unconnected nodes were excluded from visualization, and all other parameters remained at default settings.​ AutoDock Vina v1.1.2 (Scripps Research, CA, U.S.A.) was used for molecular docking to investigate the binding interactions between Rha and key targets in the HIF-1α signaling pathway. The two-dimensional structures of Rha-derived ligands were acquired and converted into three-dimensional mol2 format via ChemOffice software, followed by energy minimization and structural characterization. Both ligand and receptor structures were prepared using PyMOL and the three-dimensional crystal structures of human HIF-1α were retrieved from the RCSB Protein Data Bank and exported in PDBQT format. Docking simulations and binding affinity assessments of Rha with individual target proteins were performed with AutoDock Vina. The obtained docking poses, demonstrating specific molecular interactions of Rha with HIF-1α, were examined and visualized through PyMOL. All statistical analyzes were conducted using GraphPad Prism version 10 (GraphPad Software Inc., U.S.A.). Prior to analysis, all datasets were tested for normality. Data that did not meet the assumption of normal distribution were analyzed using appropriate non-parametric tests. Intergroup comparisons were performed using Student's t-test for two groups and one-way analysis of variance (ANOVA) followed by Tukey's post hoc test for multiple groups. Statistical significance was defined as * p  < 0.05, ** p  < 0.01, *** p  < 0.001, and **** p  < 0.0001, with ‘ns’ indicating no significant difference.

AP is a common clinical disease characterized by severe upper abdominal pain, nausea, and vomiting. In recent years, both the incidence and mortality rates of AP have shown a progressive upward trend [ 27 , 28 ]. Current clinical management primarily relies on supportive strategies such as fluid resuscitation, antibiotic administration, analgesic therapy, and endoscopic retrograde cholangiopancreatography [ 29–31 ]. Nevertheless, the therapeutic effectiveness of these conventional approaches remains limited, with a notable absence of targeted interventions addressing acinar cell injury [ 32 ]. As the major secretory units of the exocrine pancreas, acinar cells are particularly vulnerable to oxidative and inflammatory injury during AP. Early acinar cell death leading to pancreatic necrosis constitutes a crucial determinant of disease severity and patient mortality [ 33 ]. Moreover, the degree of acinar injury correlates directly with clinical severity and prognostic outcomes in AP [ 26 ]. Therefore, interventions that protect acinar cells and restore redox balance are essential for improving AP outcomes [ 26 , 34 ]. In this study, Rha was shown to significantly mitigate acinar cell injury, attenuate inflammatory responses, and alleviate oxidative stress in three classic murine models of AP. Notably, Rha also exhibited a distinct protective effect in hPACs. Rha decreased mitochondrial ROS generation and preserved mitochondrial membrane potential, thereby improving cellular energy metabolism and viability. These findings reveal, for the first time, the protective effect of Rha against AP, supporting its potential as a novel antioxidant and anti-inflammatory agent. Rha is known for its strong redox-regulatory and anti-inflammatory properties. Traditional Chinese medicine (TCM) formulations containing Rha, such as Qingyi Granules [ 35 ] and Pancogrit [ 36 ], have shown broad efficacy across inflammatory conditions. Yet, its therapeutic value in AP had not been defined. Given that excessive ROS and necrosis amplify pancreatic injury [ 12 , 37 ], the antioxidant action of Rha may represent a crucial mechanism underlying its cytoprotective effects. Network pharmacology analysis identified HIF-1α as a key target of Rha in alleviating AP. HIF-1α is a redox-sensitive transcription factor that orchestrates cellular adaptation to hypoxia and inflammation [ 38 , 39 ]. Within the AP microenvironment, tissue ischemia and edema trigger HIF-1α activation, leading to mitochondrial dysfunction, necroptosis, and ROS accumulation. Consistent with prior reports [ 40 ], our in vitro and in vivo data revealed that Rha inhibited HIF-1α expression and downregulated RIP3 and p -MLKL, and mitigated oxidative damage, thereby preserving acinar cell function and tissue integrity. In conclusion, Rha ameliorated AP by reducing oxidative stress and acinar cell necrosis through inhibition of the HIF-1α signaling pathway. As a chemically defined botanical monomer with favorable biosafety and bioavailability, Rha demonstrates clear mechanistic advantages and translational potential. These findings highlight Rha as a promising redox-regulating therapeutic candidate for the treatment of AP.

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