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Abstract

Background

Cystitis glandularis is a metaplastic lesion of the bladder mucosa. The underlying pathogenesis of cystitis glandularis and the potential association between the therapeutic effects of pachymic acid and ferroptosis remain unclear.

Methods

Human bladder epithelial cells (SV40-immortalized human urothelial cells-1 (SV-HUC-1)) and Sprague–Dawley rat bladders were treated with lipopolysaccharide, with pachymic acid and ferrostatin-1 used as interventions. Cell viability was determined using the Cell Counting Kit-8 (CCK-8) assay. Levels of reactive oxygen species, glutathione, and malondialdehyde were quantified in both cells and tissues. Levels of tumor necrosis factor-α, interleukin-1β, and interleukin-6 in rat bladder tissues were quantified using enzyme-linked immunosorbent assay kits. The expression levels of nuclear factor erythroid 2-related factor 2, glutathione peroxidase 4, and solute carrier family 7 member 11 were analyzed, and bladder inflammation was evaluated using hematoxylin and eosin staining. The mitochondrial ultrastructure of bladder tissue was examined using transmission electron microscopy.

Results

Lipopolysaccharide treatment reduced cell viability, increased malondialdehyde and reactive oxygen species levels, and decreased glutathione levels in both cells and tissues. The levels of nuclear factor erythroid 2 related factor 2, glutathione peroxidase 4, and solute carrier family 7 member 11 were reduced, while those of inflammation markers were elevated. Lipopolysaccharide stimulation induced mitochondrial structural damage in rat bladder tissues. Treatment with pachymic acid and ferrostatin-1 partially reversed these effects.

Conclusions

Ferroptosis plays a critical role in lipopolysaccharide-induced cystitis glandularis. Pachymic acid may alleviate cystitis glandularis by inhibiting ferroptosis, offering a novel therapeutic strategy for the clinical management of this disease.

Keywords

Pachymic acid ferroptosis cystitis glandularis lipopolysaccharides inflammation

Introduction

Cystitis glandularis (CG) is a metaplastic condition of the bladder mucosa that typically develops as a response to chronic bladder inflammation. During prolonged inflammation, urothelial cells proliferate and form nests within the lamina propria, which eventually differentiate into cystic or glandular structures. Previous studies have reported that CG accounts for only 0.1%–1.9% of all bladder diseases.¹ In recent years, the reported incidence of CG has been steadily increasing, largely due to the increased use of cystoscopy.² The clinical symptoms primarily include recurrent urinary frequency, urgency, pain, difficulty urinating, and either microscopic or gross hematuria.³ The diagnosis and management of CG remain controversial. Given that CG frequently coexists with glandular carcinoma, a potential link may exist between the two diseases.^4,5 Most scholars believe that CG is a precancerous lesion that requires early clinical intervention.⁶ However, other experts contend that CG only requires regular follow-up.⁷ Understanding the mechanisms of CG and identifying novel treatment approaches are of great clinical significance.

Ferroptosis is a unique form of cell death, distinct from apoptosis, necrosis, or autophagy, characterized by iron-dependent lipid peroxidation. The biochemical features of ferroptosis include the accumulation of reactive oxygen species (ROS), mitochondrial structural changes, glutathione (GSH) depletion, dysfunction of GSH peroxidase 4 (GPX4), and lipid peroxide accumulation.⁸ Solute carrier family 7 member 11 (SLC7A11), also known as xCT, is an essential component of the Xc⁻ system. SLC7A11 facilitates the transport of extracellular oxidized cystine into the cell, where it is converted into cysteine, a crucial precursor for the synthesis of GSH.⁹ Inactivation of SLC7A11 reduces cystine transport into the cell, which in turn results in decreased GSH levels, impairing GPX4 activity. GPX4 is a key regulator of ferroptosis, and its inactivation promotes the accumulation of lipid peroxides, which in turn causes an increase in intracellular ROS levels.¹⁰ Nuclear factor erythroid 2-related factor 2 (Nrf2) is an intracellular transcription factor responsible for regulating the cell’s defense response to oxidative stress.¹¹ Numerous studies have shown that Nrf2 also plays a crucial role in ferroptosis by regulating iron homeostasis, GSH metabolism, and GPX4 expression, thereby inhibiting iron-dependent lipid peroxidation.¹² Many diseases have been associated with ferroptosis, including neurodegenerative disorders such as Alzheimer and Parkinson diseases¹³ and cardiovascular conditions such as ischemic heart disease.¹⁴ Additionally, several inflammation-related disorders, such as periodontitis and mastitis, have been shown to be associated with ferroptosis.^15,16 However, the association between ferroptosis and the pathogenesis and progression of CG remains unclear.

Numerous studies have shown that chronic infection is one of the most important causes of CG, primarily due to ascending urinary tract infections caused by Escherichia coli.¹⁷ Lipopolysaccharide (LPS), an endotoxin extracted from the outer leaflet of the outer membrane of gram-negative bacteria, is widely used in the construction of various inflammation models, such as acute kidney injury¹⁸ and rheumatoid arthritis.¹⁹ LPS has been proven to be an effective stimulant for activating the innate immune response through the Toll-like receptor (TLR)-4 pathway.²⁰ Numerous studies have utilized the LPS intravesical instillation method to establish animal models of cystitis. However, conventional protocols typically involve no more than 10 instillations over a short timeframe to induce inflammatory models such as those of acute cystitis or interstitial cystitis.^20,21 In a previous study, researchers successfully constructed a CG rat model by extending the LPS instillation regimen, administering LPS every other day for 40 days (a total of 20 instillations) in Sprague–Dawley (SD) rats.²²

Pachymic acid (PA) is a lanostane-type triterpenoid compound extracted from Poria cocos, exhibiting multiple pharmacological effects, including antitumor, anti-inflammatory, hypoglycemic, and hypnotic activities.²³ PA alleviates ferroptosis by upregulating key ferroptosis-related proteins, including GPX4, SLC7A11, heme oxygenase-1 (HO-1), and Nrf2. This mechanism confers protection against ischemia–reperfusion-induced acute kidney injury in mice, effectively reversing the resultant damage.²⁴ Reports indicate that PA suppresses inflammatory responses in chondrocytes by upregulating SIRT6 expression and inhibiting the nuclear factor kappa B (NF-κB) signaling pathway, thereby slowing the progression of osteoarthritis.²⁵ Previous studies have shown that PA significantly reduces the levels of interleukin-1 (IL-1), IL-6, and lactate dehydrogenase in the bladder tissues of CG mouse models.²⁶ However, it remains unclear whether its anti-inflammatory effects on CG are mediated through the inhibition of ferroptosis.

In the present study, we employed LPS stimulation of normal human bladder epithelial cells and intravesical LPS perfusion in rats to establish both in vitro and in vivo CG models. Treatment with PA in these models demonstrated that PA alleviates LPS-induced CG by inhibiting ferroptosis. This research highlights the therapeutic potential of PA for CG, providing new insights and directions for future treatment approaches.

Methods

Cell culture

The immortalized human bladder epithelial cell line SV40-immortalized human urothelial cells-1 (SV-HUC-1) was obtained from Procell (China, CL-0222). Cells were maintained in Ham’s F-12K (Kaighn’s) medium (Procell) supplemented with 10% fetal bovine serum (Sigma). The culture was incubated at 37°C in a humidified environment containing 5% CO₂. The medium was replaced every 2 days, and the cells were passaged every 3 days. When the culture reached 70%–80% confluency, the cells were harvested for subsequent experiments.

Cell viability assay

To evaluate cell viability, the Cell Counting Kit-8 (CCK-8; Biosharp) assay was employed. SV-HUC-1 cells were seeded in a 96-well plate at a density of 1 × 10⁴ cells per well. Following incubation in a CO₂ incubator until the confluency reached 80%, various concentrations of LPS (MCE, HY-D1056), ferrostatin-1 (Fer-1) (MCE, HY-100579), and PA (MCE, HY-N0371) were added according to the experimental groups. After 24 h of treatment, the culture medium was discarded, and 100 μL of 10% CCK-8 solution was added to each well. The plate was subsequently incubated at 37°C in 5% CO₂ for 4 h. Finally, absorbance at 450 nm was measured using a microplate reader (Bio-Rad). Cell viability was calculated using the following formula: (Experimental group − Blank group)/(Control group − Blank group) × 100%.

Measurement of GSH

The cells were harvested and rinsed three times with phosphate-buffered saline (PBS) and then resuspended in the same buffer for counting. The cells were then lysed ultrasonically and centrifuged at 1300 ×g for 10 min to obtain the supernatant. For the rat bladder tissue, the weight was recorded, and Reagent 1 was added in a 1:9 ratio (tissue weight in grams to reagent volume in milliliters). The tissue was then homogenized using a glass homogenizer and centrifuged at 1300 ×g for 10 min to collect the supernatant. The total GSH content in this supernatant was quantified according to the manufacturer’s instructions using a GSH assay kit (Nanjing Jiancheng, A006-2-1).

Measurement of malondialdehyde (MDA)

The cells were further lysed using western and immunoprecipitation (IP) cell lysis buffer (Beyotime, P0013) at a ratio of 0.1 mL per 1 million cells. For rat bladder tissue, the sample was homogenized in a glass homogenizer at a tissue mass-to-lysis buffer volume ratio of 1:9 (w/v). After homogenization or lysis, the samples were centrifuged for 10 min at 10,000–12,000 ×g and 4°C, and the supernatant was collected. The MDA assay kit (Beyotime, S0131) was used for subsequent measurements, according to the manufacturer’s instructions.

Measurement of ROS

The intracellular ROS levels were measured using a ROS assay kit (Beyotime, S0033) according to the manufacturer’s instructions. Briefly, after collecting the cells, the probe was loaded by diluting 2′7′-dichlorodihydrofluorescein diacetate (DCFH-DA) with Ham’s F-12K (Kaighn’s) basal medium at a ratio of 1:1000 and incubating at 37°C for 20 min. The mixture was gently inverted every 3–5 min to ensure thorough probe–cell interaction. Subsequently, the cells were resuspended in PBS. ROS detection was performed using a flow cytometer (CF-700, URIT). The excitation wavelength was set at 488 nm, and fluorescence from 10,000 cells was recorded through the fluorescein isothiocyanate (FITC) channel. Cells were gated by forward scatter (FSC) and side scatter, with a threshold of FSC <5 × 10³ to exclude debris and dead cells. Unstained cells served as a reference to eliminate background fluorescence and nonspecific staining while simultaneously setting the fluorescence threshold. Intracellular ROS oxidize nonfluorescent DCFH to generate fluorescent DCF. Intracellular ROS levels were determined based on DCF fluorescence. The mean fluorescence intensity for each group was calculated using FlowJo software.

For rat bladder tissue, 5-μm thick frozen sections were mounted on slides and washed three times with cold (4°C) PBS for 5 min each. In the dark, tissue sections were incubated with 5 μM superoxide anion with dihydroethidium (DHE; Beyotime, S0064S) probe working solution at 37°C for 30 min. The sections were then washed three times with PBS for 5 min each. Finally, the sections were visualized using a fluorescence microscope.

Enzyme-linked immunosorbent assay (ELISA)

After weighing the rat bladder tissue, precooled PBS was added at a ratio of 1:9 (w/v). The tissue was homogenized using a glass homogenizer, followed by sonication. The samples were centrifuged at 5000 ×g for 5 min at 4°C, and the supernatant was collected as the test sample. Before measurement, the ELISA kit was equilibrated to room temperature for approximately 30 min. The procedure was performed according to the instructions for the IL-1β, IL-6, and tumor necrosis factor-α (TNF-α) kits. First, 150 μL of diluent was added to the standard and left to stand for 10 min, which was then diluted to generate a concentration gradient. Next, 10 μL of diluent was added to the blank well, 40 μL of the test sample to the sample well, and 50 μL of the gradient concentration of the diluted standard to each standard well. Then, 10 μL of enzyme-conjugated antibody was added to each well, mixed thoroughly, and incubated at 37°C in a water bath for 30 min. Afterward, a chromogenic substrate, diaminobenzidine, was added to each well and mixed thoroughly. The absorbance (A) value of each well was measured at 535 nm and 450 nm using a microplate reader. A standard curve was generated using the standard concentrations, and the sample concentrations were calculated accordingly.

Western blot analysis

Cells and rat bladder tissues were collected and lysed using radioimmunoprecipitation assay (RIPA, Beyotime) lysis buffer supplemented with 1% phenylmethanesulfonyl fluoride (Beyotime). The protein concentration of each sample was quantified using the bicinchoninic acid (BCA) Protein Assay Kit (Beyotime). The lysates were separated on a 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) gel and then transferred to 0.45-μm polyvinylidene fluoride (PVDF) membranes (Millipore). The membranes were blocked with 5% nonfat milk diluted in Tris-buffered saline with Tween-20 (TBST) for 2 h and incubated with primary antibodies overnight at 4°C. The primary antibodies included BCL-2 (1:2000, Proteintech, 12789-1-AP), BAX (1: 20000, Proteintech, 50599-2-Ig), cleaved caspase 3 (1:500, Proteintech, 25128-1-AP), GPX4 (1:1000, Abcam, ab125066), SLC7A11/XCT (1:1000, Abcam, ab307601), Nrf2 (1:500, Affinity, AF0639), and GAPDH (1:50000, Proteintech, 60004-1-Ig). The secondary antibodies used were horseradish peroxidase (HRP)–labeled goat anti-rabbit IgG (H + L) (1:1000, Beyotime, A0208) and HRP-labeled goat anti-mouse IgG (H + L) (1:1000, Beyotime, A0216). The PVDF membranes were incubated at room temperature for 1 h, and protein bands were visualized using an enhanced chemiluminescence (ECL) kit (Epizyme), followed by imaging using a Bio-Rad imaging system (Bio-Rad, USA). Protein expression levels were quantified using ImageJ software.

Hematoxylin and eosin (H&E) staining

Bladder tissue specimens from each group of rats were fixed in 4% formaldehyde for 24 h. Using a microtome, 4-μm thick sections were obtained from the paraffin blocks. These sections were then mounted on glass slides and dewaxed in xylene for 30 min, followed by hydration through a series of ethanol solutions (100%, 95%, and 70%). Next, the slides were subsequently rinsed in distilled water, stained with hematoxylin (Beyotime, C0105S) for 5–10 min, and then rinsed under running tap water for 5 min to promote bluing. In addition, the sections were counterstained with eosin for 2–5 min. Following the staining procedure, the slides were dehydrated using increasing concentrations of ethanol (70%, 95%, and 100%), cleared in xylene, and finally mounted using an appropriate medium. Finally, the stained sections were examined under a light microscope, and the images were captured for further analysis. The pathological sections of bladder tissues from each group of rats were scored according to the grading criteria established by Zhou et al.²⁰ The scoring system evaluated three parameters—edema severity, inflammatory cell infiltration, and Brunn’s nest quantity—each parameter graded on a 0–3 scale (0, normal; 1, mild; 2, moderate; and 3, severe). The maximum total score was 9 points. Examples of Brunn’s nest scoring criteria were as follows: 0 points, no Brunn’s nests observed under high-power field (HPF); 1 point, ≤2 Brunn’s nests per HPF; 2 points, ≤5 Brunn’s nests per HPF; and 3 points, extensive distribution of Brunn’s nests per HPF.

Animal experiments

A total of 36 female SD rats (aged 6–8 weeks and weighing 190–210 g) were obtained from Hunan Sileike Jingda Laboratory Animal Co., Ltd., and housed in the Central Laboratory of the Second Affiliated Hospital of Guilin Medical University. All animal experimental procedures complied with the “Guide for the Care and Use of Laboratory Animals” issued by the National Institutes of Health in 1996. All rats were maintained under standard laboratory conditions (temperature: 22°C ± 2°C; humidity: 40%–70%), with a 12-h light/dark cycle and free access to food and water. This study was performed in accordance with the ethical principles outlined in the Declaration of Helsinki (1975, as revised in 2024).

The 36 rats were divided randomly into 4 groups (n = 9 per group): control group, LPS group, LPS+Fer-1 group, and LPS+PA group. The specific procedure is as follows: SD rats were placed in an induction chamber and anesthetized using inhaled isoflurane gas. After 40 s, anesthesia depth was assessed. Once anesthesia was successfully induced, the rats were fitted with a face mask to maintain anesthesia and positioned supine on the surgical table. The urethral opening and perineal skin were disinfected with povidone–iodine. A 7-cm disposable epidural catheter was inserted into the urethral opening until urine flowed out, maintaining an insertion depth of approximately 4 cm. The abdomen was gently pressed to ensure complete bladder emptying. The other end of the catheter was connected to a 0.5-mL disposable syringe, and 0.2 mL of LPS solution (1 mg/mL) was slowly injected into the bladder. The catheter was slowly withdrawn, and the urethral opening was immediately clamped for 30 min before being released. The control group underwent the same procedure, with the exception that physiological saline was instilled instead of LPS. Bladder instillation was performed every 2 days for a total of 20 sessions. After 20 days of LPS instillation, the LPS+Fer-1 group received intraperitoneal injections of Fer-1 (5 mg/kg), while the LPS+PA group received intraperitoneal injections of PA (40 mg/kg) every 2 days for a total of eight injections. Equal volumes of dimethyl sulfoxide solution were injected into the LPS and the control groups. After a total of eight intraperitoneal injections, the rats were euthanized via cervical dislocation.

Transmission electron microscopy (TEM)

Following collection, rat bladder tissues were rinsed with precooled PBS and cut into 1-mm cubes and fixed in 2.5% glutaraldehyde at 4°C for 24 h. Postfixation, tissues were incubated in 1% osmium tetroxide at room temperature for 1–2 h. The samples were then dehydrated through a graded ethanol series (30%, 50%, 70%, 90%, and 100%). After dehydration, the specimens were embedded in epoxy resin, sectioned into ultrathin slices (70–90 nm) using an ultramicrotome, and mounted on copper grids. The sections were subsequently stained with uranyl acetate for 10–15 min, followed by lead citrate for 5–10 min, and examined using a Hitachi-HT7700 electron microscope. Mitochondrial length was measured using ImageJ, and damaged mitochondria were manually quantified. The assessment of mitochondrial damage was based on the following three aspects: (a) cristae structure (presence or disappearance); (b) membrane integrity (normal (smooth outer membrane), shrunken (invaginated outer membrane), or ruptured (discontinuous outer membrane)); (c) degree of swelling (determined by the ratio of mitochondrial long-axis to short-axis length (normal ≈ 2–3; swollen = 1, indicating a more spherical morphology)).²⁷

Statistical analysis

All statistical analyses were performed using GraphPad Prism 9.0 software (GraphPad 9.0, San Diego, CA, USA). The results were expressed as mean ± standard deviation. One-way analysis of variance was used to compare p-values between the four groups, and Student’s t-test was used to compare data between two groups, with a p-value of less than 0.05 indicating statistical significance.

Results

LPS treatment induces ferroptosis in SV-HUC-1 cells

To evaluate the effect of LPS treatment on ferroptosis, we treated SV-HUC-1 cells with different concentrations of LPS (0, 1, 10, and 20 μg/mL) for 24 h. Cell viability at each concentration was determined using the CCK8 assay, and the results showed that LPS treatment inhibited SV-HUC-1 cell viability in a dose-dependent manner (Figure 1(a)). Additionally, we measured the GSH and MDA levels in each group and found that GSH levels decreased as LPS concentration increased, while MDA levels showed an opposite trend, increasing with higher LPS concentrations (Figure 1(b) and (c)). LPS treatment reduced cellular antioxidant capacity, accompanied by the accumulation of lipid peroxides. Western blot analysis demonstrated that LPS induction downregulated the expression of Nrf2 and two ferroptosis markers, SLC7A11 and GPX4, indicating that LPS treatment triggered ferroptosis in SV-HUC-1 cells (Figure 1(d)). To further determine whether LPS treatment induces apoptosis, we examined the expression levels of cleaved caspase-3, Bax, and Bcl-2 via western blot analysis. The results showed that treatment with 20 μg/mL LPS did not induce significant changes in the expression of these apoptosis-related proteins (Figure 1(e)).

Figure 1.

The effect of LPS treatment on ferroptosis in SV-HUC-1 cells. (a) The effect of LPS treatment on SV-HUC-1 cell viability was measured using the CCK-8 assay after treating the cells with different concentrations of LPS (0, 1, 10, and 20 µg/mL) for 24 h. (b) GSH levels in cells treated with different concentrations of LPS. (c) MDA levels in cells treated with different concentrations of LPS. (d) Western blot analysis of Nrf2, SLC7A11, and GPX4 expression in cells after stimulation with different concentrations of LPS and (e) Western blot analysis of cleaved caspase-3, Bax, and Bcl-2 expression in cells after stimulation with different concentrations of LPS. The above data are presented as mean value ± SD, ns means p > 0.05, *p < 0.05, **p < 0.01. GSH: glutathione; LPS: lipopolysaccharide; MDA: malondialdehyde; SV-HUC-1: SV40-immortalized human urothelial cells-1.

PA alleviates the decline in cell viability caused by LPS treatment

To investigate the effects of PA and Fer-1 on ferroptosis, SV-HUC-1 cells were first treated with different concentrations of PA (0, 20, 40, 60, and 80 μg/mL) and Fer-1 (0, 1, 2, 4, 6, 8, and 10 μM), respectively, and cytotoxicity was assessed using the CCK8 assay. The results showed that PA concentrations below 60 μg/mL had no significant impact on cell viability, whereas a marked reduction in cell viability was observed at 80 μg/mL (Figure 2(a)). Similarly, Fer-1 concentrations ≤8 μM exhibited no apparent cytotoxicity, but cell viability significantly declined when the concentration reached 10 μM (Figure 2(b)). Subsequently, we pretreated SV-HUC-1 cells with 20 μg/mL LPS for 24 h, followed by treatment with different concentrations of PA (0, 20, 40, and 60 μg/mL) and Fer-1 (0, 1, 2, 4, 6, and 8 μM) for another 24 h. Cell viability was measured using the CCK8 assay. Although LPS and PA treatment reduced cell viability, PA effectively improved the viability of LPS-stimulated SV-HUC-1 cells, indicating its rescue effect on LPS-induced cell death (Figure 2(c)). Furthermore, when LPS-stimulated SV-HUC-1 cells were treated with Fer-1, a significant increase in cell viability was observed at 4 μM Fer-1 compared with the control group. However, the protective effect of Fer-1 against LPS-induced loss of cell viability was markedly reduced at concentrations exceeding 4 μM. This reduction in efficacy may be attributed to lysosome-dependent cell death and nonspecific scavenging of ROS induced by high concentrations of Fer-1.^28,29 Therefore, 4 μM was chosen for subsequent experiments to minimize potential off-target effects (Figure 2(d)). Higher Fer-1 concentrations reduced cell viability, which we speculate may be due to drug toxicity (Figure 2(d)). Therefore, we used 60 μg/mL PA and 4 μM Fer-1 for subsequent treatment to minimize the potential off-target effects and ensure the reliability of the experimental results. Therefore, 60 μg/mL PA and 4 μM Fer-1 were selected for subsequent experiments.

Figure 2.

PA alleviates the decline in cell viability caused by LPS treatment. (a) Effect of different concentrations of PA on the viability of SV-HUC-1 cells: After treating the cells with different concentrations of PA (0, 20, 40, 60, and 80 μg/mL) for 24 h, cell viability was measured using the CCK8 assay. (b) Effect of different concentrations of Fer-1 on the viability of SV-HUC-1 cells. After treating the cells with different concentrations of Fer-1 (0, 1, 2, 4, 6, 8, and 10 μM) for 24 h, cell viability was measured using the CCK8 assay. (c) The effect of PA on cell viability after LPS treatment. SV-HUC-1 cells stimulated with LPS were treated with different concentrations of PA (0, 20, 40, and 60 μg/mL) for 24 h, and cell viability was measured using the CCK-8 assay and (d) the effect of Fer-1 on cell viability after LPS treatment: SV-HUC-1 cells stimulated with LPS were treated with different concentrations of Fer-1 (0, 1, 2, 4, 6, and 8 μM) for 24 h, and cell viability was measured using the CCK-8 assay. The above data are presented as mean value ± SD, ns means p > 0.05, *p < 0.05 compared with 0 μg/mL LPS. ^#p < 0.05 compared with the LPS group. LPS: lipopolysaccharide; PA: pachymic acid; SV-HUC-1: SV40-immortalized human urothelial cells-1.

PA alleviates LPS-induced cell damage by inhibiting ferroptosis

To determine whether PA can alleviate LPS-induced ferroptosis in SV-HUC-1 cells by inhibiting ferroptosis, we divided the cells into four groups: control, LPS, LPS+Fer-1, and LPS+PA. We measured MDA and GSH levels in each group. The results showed that MDA levels in the LPS group were significantly increased compared with those in the control group, while GSH levels were markedly reduced. The PA-treated group exhibited significantly higher GSH levels than the LPS group, accompanied by a marked reduction in MDA content. These findings suggest that PA effectively alleviates LPS-induced lipid peroxide accumulation while enhancing cellular antioxidant capacity. Notably, no statistically significant differences in GSH or MDA levels were observed between LPS+Fer-1 and LPS+PA groups (Figure 3(a) and (b)). To demonstrate LPS-induced ROS changes in the in vitro CG model, the cells were incubated with the fluorescent probe DCFH-DA, followed by the measurement of DCF fluorescence intensity using flow cytometry to assess intracellular ROS levels. To demonstrate LPS-induced ROS changes in the in vitro CG model, we assessed intracellular reactive ROS levels by measuring DCF fluorescence intensity using flow cytometry. The results revealed that ROS levels in the LPS group were significantly elevated compared with those in the control group. After 24 h of treatment with PA and Fer-1, ROS levels in both the LPS+Fer-1 and LPS+PA groups demonstrated a notable decrease (Figure 3(c)). Additionally, compared with the LPS group, the expression levels of Nrf2, SLC7A11, and GPX4 were upregulated in the LPS+PA and LPS+Fer-1 groups (Figure 3(d)). No statistically significant differences were detected between the LPS+PA and LPS+Fer-1 groups. These results indicate that PA can significantly alleviate LPS-induced ferroptosis in the cells.

Figure 3.

PA alleviates LPS-induced cell damage by inhibiting ferroptosis. (a) The intracellular GSH content in each group of cells was measured using a colorimetric method. (b) The intracellular MDA content in each group of cells was measured using a colorimetric method. (c) After staining the cells with DCFH-DA, the ROS fluorescence intensity in each group was analyzed via flow cytometry and (d) Western blot analysis of Nrf2, SLC7A11, and GPX4 expression in different cell groups. The above data are presented as mean value ± SD, ns means p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001. GSH: glutathione; LPS: lipopolysaccharide; MDA: malondialdehyde; PA: pachymic acid; ROS: reactive oxygen species.

The therapeutic effect of PA on LPS-induced CG in rats

We established a CG model in rats through intravesical instillation of LPS. After fixing bladder tissues and performing H&E staining on sections, microscopic examination revealed that the LPS group exhibited typical CG pathological features in bladder epithelium, including Brunn’s nest formation, epithelial layer edema, inflammatory cell infiltration, and lymphocytosis (Figure 4(a)). These pathological manifestations were consistent with the histopathological features of human CG,³⁰ confirming successful model construction. In contrast, both the LPS+Fer-1 and LPS+PA groups exhibited significantly alleviated inflammatory manifestations, disappearance of Brunn’s nests, and reduced epithelial edema. For the quantitative assessment of inflammation severity, three rat bladders from each group were selected, with five nonoverlapping fields per bladder undergoing pathological scoring. The results demonstrated that inflammation scores were significantly higher in the LPS group than in the control group, while PA and Fer-1 treatments significantly reduced the scores (Figure 4(b)). Further ELISA detection of bladder tissue inflammatory cytokines (IL-1β, IL-6, and TNF-α) revealed that the LPS group showed markedly elevated levels of inflammatory factors compared with the control group, whereas the LPS+Fer-1 and LPS+PA groups exhibited significant reductions (Figure 4(c) to (e)). These findings indicate that PA demonstrates significant therapeutic effects on LPS-induced CG, effectively alleviating CG-associated inflammatory responses.

Figure 4.

Therapeutic effect of PA on LPS-induced CG. (a) Histopathological sections of bladder tissues from each group were observed via H&E staining. The magnified regions revealed the epithelial layer of the bladder. In the LPS group, distinct formation of Brunn’s nests was observed, accompanied by epithelial edema and increased inflammatory cell infiltration. Following treatment with Fer-1 and PA, Brunn’s nests disappeared, epithelial edema was alleviated, and inflammatory cell infiltration was reduced (Scale bar = 100 μm). (b) At 400× magnification, histopathological features across multiple fields of view were observed. The severity of inflammation was scored and visualized as a heatmap, where columns represent three individual animals per group, and rows correspond to five distinct microscopic fields from each animal. Additionally, statistical analysis of the inflammatory scores was performed. (c) to (e) Levels of IL-1β, IL-6, and TNF-α in bladder tissues from each group of rats. The above data are presented as mean value ± SD, ns means p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001. CG: cystitis glandularis; H&E: hematoxylin and eosin; LPS: lipopolysaccharide; PA: pachymic acid.

PA inhibits ferroptosis and alleviates LPS-induced CG

To determine whether ferroptosis occurred in the rat groups, we measured the concentrations of MDA and GSH in bladder tissues using assay kits. The results were consistent with those observed in vitro, showing that the MDA concentration in the LPS group was significantly higher than that in the control group, while the GSH concentration was markedly lower. These findings indicate that the occurrence of CG is related to LPS-induced ferroptosis. The results for the LPS+Fer-1 and LPS+PA groups demonstrated that both Fer-1 and PA effectively reduced the MDA concentration in the CG rat model while increasing the GSH levels (Figure 5(a) and (b)). To further assess the changes in ROS levels in the bladder tissues of the rats, we performed frozen sectioning and incubated the tissues with the DHE probe solution. Fluorescence microscopy revealed that the fluorescence intensity in the LPS group was significantly higher than that in the control group, indicating that LPS stimulation led to ROS accumulation in the bladder tissues. In contrast, the fluorescence intensity in the LPS+Fer-1 and LPS+PA groups was markedly reduced compared with that in the LPS group (Figure 5(c)). TEM revealed that mitochondria in the bladder tissues of the LPS group exhibited ferroptosis-associated morphological alterations, including reduced cristae, shrunken membranes, and extensive swelling. However, Fer-1 and PA treatments significantly mitigated LPS-induced mitochondrial damage (Figure 5(d)). Further quantitative analysis demonstrated that compared with the control group, the LPS group showed a significant shortening of mitochondrial average length and a marked increase in the proportion of damaged mitochondria. In contrast, both the LPS+Fer-1 and LPS+PA groups displayed restored mitochondrial length and a significant reduction in the percentage of impaired mitochondria (Figure 5(e) and (f)). Additionally, western blot analysis revealed that the expression levels of Nrf2, SLC7A11, and GPX4 in the LPS group were lower than those in the control group, while the use of PA and Fer-1 reversed this trend (Figure 5(g)). Collectively, these findings demonstrate that PA alleviates LPS-induced CG by inhibiting ferroptosis, highlighting the pivotal role of ferroptosis in LPS-induced CG pathogenesis.

Figure 5.

PA inhibits ferroptosis and alleviates LPS-induced CG. (a) The intracellular GSH content in each group of rat bladder tissues was measured using a colorimetric method. (b) The intracellular MDA content in each group of rat bladder tissues was measured using a colorimetric method. (c) The DHE fluorescent probe staining results of frozen bladder tissue sections from each rat group were observed using an inverted fluorescence microscope. Quantitative analysis of fluorescence intensity in the sections was performed using ImageJ software (scale bar = 50 μm). (d) The mitochondrial structure of rat bladder tissue observed under transmission electron microscopy, with the mitochondria indicated by red arrows. (e) and (f). Mitochondrial length was quantitatively analyzed using ImageJ software, and the proportion of damaged mitochondria relative to the total mitochondrial count was manually assessed (mitochondrial length was quantified in 10 mitochondria, and the mitochondrial damage rate was assessed based on a minimum sample size of 40 mitochondria) and (g) Western blot analysis of Nrf2, SLC7A11, and GPX4 expression in rat bladder tissues from different groups. The above data are presented as mean value ± SD, ns means p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001. CG: cystitis glandularis; GSH: glutathione; LPS: lipopolysaccharide; MDA: malondialdehyde.

Discussion

With the widespread use of cystoscopy, the reported incidence of CG has been steadily increasing.¹ Clinically, CG is primarily characterized by recurrent urinary frequency, difficulty urinating, and visible hematuria. It remains controversial whether CG can lead to malignancy in clinical practice. In high-risk CG cases, the treatment approach is similar to that of papillary bladder tumors, with transurethral resection being the primary method, which imposes significant psychological and physical burdens on patients.³¹ Therefore, understanding the pathogenesis of CG and identifying new treatment methods are crucial. In this study, we demonstrated that PA effectively inhibits LPS-induced ferroptosis in bladder cells and tissues, reducing inflammation and improving the pathological damage associated with CG.

Chronic infection, particularly with E. coli (a common uropathogen), is a primary driver of CG. LPS, a key component of bacterial endotoxin, is widely used to construct inflammatory models. For instance, Liu et al.³² established a murine lung injury model through bronchial instillation of LPS, while Wang et al.³³ developed a periodontal inflammation cell model using LPS-stimulated RAW264.7 cells. To establish a CG cell model, we stimulated SV-HUC-1 cells with varying LPS concentrations and observed that treatment with 20 μg/mL LPS for 24 h significantly reduced cell viability, consistent with the findings of Wang et al.,³⁴ who successfully induced CG in SD rats via intravesical LPS instillation. In our study, rats treated with LPS displayed CG-defining histopathological features, including Brunn’s nests, diffuse inflammatory cell infiltration from the mucosa to the lamina propria, and pronounced edema. Histological scoring (based on edema, inflammatory cell density, and Brunn’s nest count) demonstrated significantly higher inflammation scores in the LPS group compared with those in the control group, validating the CG model.

In recent years, numerous active compounds derived from traditional Chinese medicine have demonstrated broad anti-inflammatory and antitumor properties. Studies have indicated that certain herbal extracts, such as Astragaloside IV and rosavin,^35,36 exhibit potent antiferroptotic effects by upregulating Nrf2 expression and activating ferroptosis-related proteins, including SLC7A11 and GPX4, thereby mitigating PM2.5 (fine particulate matter)-induced pulmonary injury. PA, an active compound derived from the traditional Chinese medicine Poria cocos, exhibits broad anti-inflammatory and antitumor properties.³⁷ Studies have suggested that PA mitigates inflammation by suppressing ferroptosis. For example, PA enhances Nrf2 signaling, modulates oxidative stress, and inhibits NF-κB-mediated inflammatory pathways, ameliorating alcoholic liver injury.³⁸ Notably, PA alleviates cyclophosphamide-induced CG in mice by lowering the levels of LDH and inflammatory cytokines.²⁶ To validate PA’s efficacy in LPS-induced CG, we first assessed its impact on cell viability via CCK-8 assays. PA and Fer-1 effectively reversed LPS-induced cytotoxicity. Higher Fer-1 concentrations (>4 μM) reduced viability, likely due to drug toxicity. In vivo, intraperitoneal PA administration reduced inflammatory cell infiltration, Brunn’s nest formation, and histopathological scores in CG rats. PA also normalized LPS-elevated TNF-α, IL-1β, and IL-6 levels, confirming their shared anti-inflammatory efficacy.

Mitochondria are dynamic organelles with dual-membrane structures that regulate metabolism, energy conversion, signaling, and programmed cell death.³⁹ As primary sources of ROS, mitochondria play a dual role: physiological ROS support metabolic signaling, while pathological ROS accumulation trigger oxidative stress, lipid peroxidation, and mitochondrial structural damage (e.g. cristae loss, swelling, and membrane wrinkling). This initiates a vicious cycle of impaired electron transport and amplified ROS production, culminating in ferroptosis.⁴⁰ TEM revealed LPS-induced mitochondrial swelling, rounding, and cristae reduction in rat bladders, whereas PA and Fer-1 restored mitochondrial integrity. Quantitative analysis further confirmed shortened mitochondrial length and elevated damaged mitochondria in the LPS group, both of which were reversed by PA and Fer-1. Furthermore, flow cytometry analysis combined with DHE fluorescence staining verified that LPS induces ROS overproduction in CG models, while both PA and the ferroptosis inhibitor Fer-1 significantly attenuated this oxidative stress response. These findings provide further evidence for the causal relationship between mitochondrial damage and excessive ROS production. Nevertheless, the precise mechanisms underlying mitochondrial dysfunction and ROS hyperaccumulation in CG remain to be elucidated.

Ferroptosis, an iron-dependent form of cell death driven by lipid peroxidation, is closely associated with inflammation. The hallmarks of ferroptosis include GSH depletion, GPX4 inactivation, and MDA accumulation—a marker of lipid peroxidation.⁴¹ LPS stimulation reduced intracellular GSH and elevated MDA in SV-HUC-1 cells in a dose-dependent manner. Treatment with PA restored GSH and suppressed MDA in vitro and in vivo, indicating enhanced antioxidant capacity. These findings underscore oxidative stress as a key pathological feature of CG and highlight the inhibition of ferroptosis as a therapeutic mechanism of PA.

Nrf2, a master regulator of oxidative stress responses, protects against inflammation and oxidative damage. Previous studies have demonstrated that PA activates Nrf2 to combat atherosclerosis.¹⁶ Nrf2 directly binds to antioxidant response elements in the promoters of SLC7A11 and GPX4, driving their expression. Knockout of Nrf2 exacerbates sepsis-associated encephalopathy by suppressing SLC7A11, GPX4, and GSH.⁴² SLC7A11 (xCT), a subunit of the cystine/glutamate antiporter system Xc⁻, imports cystine for GSH synthesis. Its downregulation depletes GSH, inactivates GPX4, and triggers ferroptosis.⁴³ We performed western blot analysis to detect the expression levels of Nrf2, SLC7A11, and GPX4 in cells and bladder tissues. The results revealed that low-concentration LPS (1 μg/mL) did not induce significant changes in the expression of these proteins in SV-HUC-1 cells, whereas high-concentration LPS (20 μg/mL) markedly suppressed their expression, indicating a dose-dependent induction of ferroptosis by LPS. This phenomenon may be associated with the dual regulatory nature of the Nrf2 pathway: under mild oxidative stress, Nrf2 is activated to exert protective effects, whereas severe stimulation leads to impaired nuclear translocation of Nrf2 and suppression of downstream target gene expression.⁴⁴ Further detection of ferroptosis-related protein expression in cells and rat bladder tissues demonstrated that in both in vitro and in vivo models, the LPS group exhibited significantly reduced protein levels of Nrf2, SLC7A11, and GPX4 compared with the control group. However, treatment with PA and Fer-1 resulted in a notable upregulation of these proteins in treated cells and bladder tissues relative to the LPS group. Notably, across both models, no statistically significant differences were observed between the LPS+Fer-1 and LPS+PA groups in terms of Nrf2, SLC7A11, and GPX4 expression. These findings indirectly suggest that oxidative stress and ferroptosis are effective therapeutic targets for alleviating CG, although the specific mechanisms remain to be fully elucidated.

Our study demonstrated that LPS promotes the pathological progression of CG by inducing antioxidant system imbalance (manifested as GSH depletion and MDA accumulation) in SV-HUC-1 cells and rat bladder tissues, leading to ferroptosis-dependent cellular damage. PA inhibits CG-associated ferroptosis by activating the expression of Nrf2, SLC7A11, and GPX4, thereby restoring cellular redox homeostasis, improving CG pathological phenotypes, and alleviating CG inflammation. However, this study has several limitations that warrant attention. First, as a lipophilic triterpenoid, the poor water solubility of PA may restrict its in vivo bioavailability, thereby potentially affecting its therapeutic efficacy. Furthermore, current studies lack a systematic evaluation of PA’s chronic toxicity and pharmacokinetics, necessitating further research to establish a safe dosing window.⁴⁵ Our animal experiments did not systematically evaluate the dose–response relationship of PA at varying doses in CG models, necessitating future establishment of gradient dosing models to determine the optimal therapeutic window. Second, although PA demonstrates therapeutic potential, its clinically applicable dosage remains undefined, and rigorous head-to-head efficacy comparisons with standard therapies such as D-mannose and antibiotics are imperative. Furthermore, based on the findings by Zhang et al.,⁴⁶ we speculate that PA exerts its therapeutic effect on CG by acting on Nrf2. However, this proposed mechanism requires further validation using conditional gene knockout models combined with multiomics analyses, which will form a focus of our future research. The mechanistic scope warrants expansion; whether the anti-CG effects of PA involve alternative signaling pathways should be examined using conditional gene knockout models integrated multiomics analyses. Finally, whether LPS-induced CG involves other types of cell death requires further investigation. The molecular bridge connecting CG progression to bladder carcinogenesis remains uncharacterized, underscoring the need to investigate PA’s dual regulatory effects within tumorigenic microenvironments using advanced disease models.

Conclusion

Our findings demonstrate that ferroptosis plays a critical role in LPS-induced CG. PA effectively ameliorates CG-associated inflammation by suppressing ferroptosis (Figure 6), suggesting that targeting ferroptosis may represent a novel therapeutic strategy for CG. PA shows promising potential as a novel therapeutic candidate for CG.

Figure 6.

PA inhibits ferroptosis in CG by upregulating Nrf2, SLC7A11, and GPX4 expression (By Figdraw).

Supplemental Material

sj-pdf-1-imr-10.1177_03000605251396755 - Supplemental material for Pachymic acid alleviates lipopolysaccharide-induced cystitis glandularis by inhibiting ferroptosis

Supplemental material, sj-pdf-1-imr-10.1177_03000605251396755 for Pachymic acid alleviates lipopolysaccharide-induced cystitis glandularis by inhibiting ferroptosis by Yongbo Tang, Haiwei Hu, Shuangyan Chen, Yueyang Gong and Bo Ge in Journal of International Medical Research

Footnotes

Acknowledgments

Not applicable.

Authorship contribution statement

Yongbo Tang and Bo Ge designed the experiments;Yongbo Tang performed experiments and collected data;Yongbo Tang and Haiwei Hu processed the data;Yongbo Tang,Shuangyan Chen,and Yueyang Gong analyzed and interpreted the results;Yongbo Tang,Shuangyan Chen,and Yueyang Gong prepared the draft manuscript;Yongbo Tang and Haiwei Hu visualized the draft manuscript;Bo Ge supervised and edited the article;Bo Ge provided final approval of the version to be published.

Compliance with ethical standards

All animal experiments were approved by the Experimental Animal Ethics Committee of Guilin Medical University,approval number: GLMC20241012. Date: 08 January 2024.

Data availability statement

The datasets used or analyzed during the current study are available from the corresponding author on reasonable request.

Declaration of conflicting interests

All authors declare that they have no conflicting interest.

Funding

This research was supported by The National Natural Science Foundation of China (No.82460158),Guangxi Key Research and Development Program (No.AB25069043),Guangxi Province Natural Science Foundation (No. 2023GXNSFAA026169),Guangxi Clinical Key Specialty Construction (Cultivation) Project,and The Science and Technology Plan of Guilin (No.20230135-1-1).

ORCID iD

Bo Ge

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