Human renal proximal tubular epithelial cells (HK-2) were procured from Procell Life Science & Technology Co., Ltd. (Wuhan, Hubei, China; Product No. CL-0109) with accompanying official certificates for STR profiling and mycoplasma-free status. To ensure ongoing quality, we perform routine in-house testing using the BeyoDirect™ Mycoplasma qPCR Detection Kit (Beyotime, Shanghai, China; Product No. C0303S); all results have been negative, consistent with the kit’s criteria (sample Ct $>$35). The cells were cultured in HK-2 Cell Complete Medium (Procell, Wuhan, Hubei, China; Product No. CM-0109) at 37 °C in a humidified 5% CO₂ incubator. The medium was refreshed every three days, and the cells were passaged at a 1:4 ratio upon reaching 80% confluence. To establish the UA-injured cell model, HK-2 cells were seeded at an appropriate density and exposed to UA (MedChemExpress, MCE, Monmouth Junction, NJ, USA; Product No. HY-B2130) for 36 h. In order to explore the function of oxidative stress in the injury induced by UA, HK-2 cells were pre-treated. Specifically, they were incubated with the antioxidant N-acetylcysteine amide (NACA; 1 mM; Sigma-Aldrich, St. Louis, MO, USA; Product No. A0737) for 2 h before being co - exposed to UA (3 mM) for 36 h. P110 peptides and the TAT fragment, synthesized by Ontores Biotechnologies (Hangzhou, Zhejiang, China), were administered at a final concentration of 1 µM three hours prior to UA intervention.

The viability of cells was evaluated using the Cell Counting Kit-8 (CCK-8; Beyotime, Shanghai, China; Product No. C0037). Cells were plated in 96-well plates (2000 cells/well), subjected to compounds at different concentrations for specific time periods, and then each well was incubated with 10 µL of CCK-8 reagent. After incubation, absorbance was measured at 450 nm using a microplate reader (BioTek, Winooski, VT, USA).

Intracellular ROS levels were detected using 2^′,7^′-dichlorodihydrofluorescein diacetate (DCFH-DA; Beyotime, Shanghai, China; Product No. S1105M). Cells were incubated with DCFH-DA in serum-free medium for 30 minutes at 37 °C in the dark. After washing with PBS, the fluorescence intensity was quantified using a microplate reader (BioTek, Winooski, VT, USA) with excitation and emission wavelengths set at 495/529 nm.

Cell viability was evaluated by propidium iodide (PI; Servicebio, Wuhan, Hubei, China; Product No. G1021) staining. After treatments, cells were gently washed twice with PBS and stained with PI solution (10 µg/mL in PBS) for 10 minutes at room temperature in the dark. The nuclei were stained again with Hoechst 33342 (10 µg/mL; Thermo Fisher Scientific, Waltham, MA, USA; Product No. R37605). Fluorescent images were acquired using a Leica STELLARIS 5 confocal laser-scanning microscope (Leica Microsystems, Wetzlar, Germany).

According to the manufacturer’s guidelines, total RNA was extracted from cells using AG RNAex Pro Reagent (Accurate Biotechnology, Changsha, Hunan, China; Product No. AG21102). Complementary DNA (cDNA) was synthesized from 1 µg total RNA with the Evo M-MLV RT Premix (Accurate Biotechnology, Changsha, Hunan, China; Product No. AG11728). Quantitative PCR (qPCR) was carried out on a QuantStudio 3 Real-Time PCR System (Thermo Fisher Scientific, Waltham, MA, USA) using the SYBR® Green Premix Pro Taq HS qPCR Kit (Accurate Biotechnology, Changsha, Hunan, China; Product No. AG11718) and specific primers. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) served as an internal reference. Gene expression levels were calculated by the 2^{-Δ⁢Δ⁢CT} method. The sequences of the primers used were as follows: 5^′-CCAGTGAAGCCGGTCAATCT-3^′ (F, DRP1); 5^′-GCTGCCGTGACAGAAAACAC-3^′ (R, DRP1); 5^′-CTGAACGAGCTGGTGTCTGT-3^′ (F, FIS1); 5^′-GAACAGGGAAAGGACAGCGA-3^′ (R, FIS1); 5^′-AAAGATGGTGTGGCCGATGT-3^′ (F, SOD2); 5^′-CAAGCCAAACGACTTCCAGC-3^′ (R, SOD2); 5^′-GTGCCACCAAGTTCAAGCAG-3^′ (F, HO-1); 5^′-CACGCATGGCTCAAAAACCA-3^′ (R, HO-1); 5^′-TGCTTACACTTACGCTGCCAT-3^′ (F, NQO-1); 5^′-CCAGTGGTGATGGAAAGCAC-3^′ (R, NQO-1); 5^′-CGGGAAGGTGAAGCGCAATG-3^′ (F, MFN); 5^′-CTGTCCAGCTCTGTGGTGAC-3^′ (R, MFN); 5^′-CTGTGGCCTGGATAGCAGAA-3^′ (F, OPA1); 5^′-GCGAGGCTGGTAGCCATATT-3^′ (R, OPA1); 5^′-AATGGGCAGCCGTTAGGAAA-3^′ (F, GAPDH); 5^′-TCGTGTTGCACTGGTTAAAGC-3^′ (R, GAPDH).

Cytosolic and mitochondrial fractions were isolated using a Mitochondria Isolation Kit (Thermo Fisher Scientific, Waltham, MA, USA; Product No. 89874) following the manufacturer’s protocol. Briefly, the cells were washed with ice-cold PBS, resuspended in ice-cold mitochondrial isolation buffer, and homogenized. The homogenate was centrifuged at 800 $\times$g for 10 min at 4 °C to pellet the nuclei and unbroken cells. The supernatant was then centrifuged at 15,000 $\times$g for 10 min at 4 °C. The resulting supernatant constituted the cytosolic fraction and the pellet contained the mitochondrial fraction.

Total cellular protein was isolated by employing RIPA lysis buffer (Thermo Fisher Scientific, Waltham, MA, USA; Product No. 89901) that had been supplemented with protease and phosphatase inhibitors (Sigma-Aldrich, St. Louis, MO, USA; Product No. 78440). The concentration of the extracted protein was measured via the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA; Product No. 23227). Subsequently, equivalent quantities of the protein samples were subjected to separation through sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and then transferred onto polyvinylidene difluoride (PVDF) membranes. Membranes were blocked and probed with primary antibodies overnight at 4 °C in TBST (Tris-buffered saline with 0.1% Tween-20). The blocking agent and dilution buffer was 5% skim milk for all antibodies, except for the anti-p-DRP1 (Ser616) antibody, for which 5% BSA was used to minimize background. The primary antibodies used were as follows: anti-GAPDH (1:5000, Proteintech, Wuhan, Hubei, China; Product No. 60004-1-Ig), anti-DRP1 (1:2000, Proteintech, Wuhan, Hubei, China; Product No. 12957-1-AP), anti-p-DRP1 (Ser616; 1:1000, Abcam, Cambridge, UK; Product No. Ab314755), anti-COX IV (1:2000, Abcam, Cambridge, UK; Product No. Ab202554), anti-FIS1 (1:2000, Proteintech, Wuhan, Hubei, China; Product No. 10956-1-AP), anti-IL-1$\beta$ (1:1000, Cell Signaling Technology, Danvers, MA, USA; Product No. 12703), anti-IL-18 (1:1000, Cell Signaling Technology, Danvers, MA, USA; Product No. 67775), anti-BCL-2 (1:2000, Proteintech, Wuhan, Hubei, China; Product No. 12789-1-AP), anti-BAX (1:5000, Proteintech, Wuhan, Hubei, China; Product No. 60267-1-Ig), anti-CYT C (1:2000, Proteintech, Wuhan, Hubei, China; Product No. 10993-1-AP), anti-cleaved Caspase-3 (1:1000, Proteintech, Wuhan, Hubei, China; Product No. 25128-1-AP), and anti-cleaved Caspase-9 (1:1000, Cell Signaling Technology, Danvers, MA, USA; Product No.9509). After that, the PVDF membranes were incubated with a horseradish peroxidase (HRP)-conjugated secondary antibody (1:5000, Proteintech, Wuhan, Hubei, China; Product No. SA00001-1 and SA00001-2) for at room temperature 2 h. The protein bands were made visible using the Thermo iBright CL1000 imaging system (Thermo Fisher Scientific, Waltham, MA, USA) along with Pierce™ ECL Western reagent (Thermo Fisher Scientific, Waltham, MA, USA; Product No. 32209). The intensities of the bands were quantified using ImageJ software (version 1.46r, National Institutes of Health, Bethesda, MD, USA), and all the data were derived from a minimum of three independent experimental runs.

Protein-protein interactions between DRP1 and FIS1 were assessed using the Pierce™ Classic IP Kit (Thermo Fisher Scientific, Waltham, MA, USA; Product No. 88804). Briefly, the magnetic beads were washed and incubated with anti-FIS1 antibody or control IgG for 60 min. Cell lysates were introduced to antibody-bound beads and subjected to an overnight incubation at 4 °C under continuous rotation. Once the incubation period was completed, the beads were washed, and the immunoprecipitated complexes were then eluted. The existence of DRP1 within the FIS1 immunoprecipitates was examined through SDS-PAGE followed by Western blotting analysis.

Cells from each group were harvested by trypsinization, pelleted by centrifugation, and fixed in 2.5% glutaraldehyde (Servicebio, Wuhan, Hubei, China; Product No. G1102-100ML) for 3 h at 4 °C. Samples were then post-fixed in 1% osmium tetroxide in 0.1 M PBS (pH 7.4) for 2 h. After dehydration using a graded ethanol series, the pellets were embedded in epoxy resin. Ultrathin sections (50 nm) were prepared, stained with uranyl acetate and lead citrate and examined under a transmission electron microscope (Hitachi High Technologies, Tokyo, Japan).

Mitochondrial membrane potential was assessed using the potentiometric dye JC-1 (MCE, Monmouth Junction, NJ, USA; Product No. HY-15534). After treatments, cells were incubated with 20 µM JC-1 in complete medium for 20 minutes at 37 °C in a 5% CO₂ atmosphere. Cells were washed with PBS, and JC-1 fluorescence (monomeric form: green, Ex $\sim$510 nm/Em $\sim$527 nm; J-aggregates: red, Ex $\sim$585 nm/Em $\sim$590 nm) was visualized and quantified using a Leica STELLARIS 5 confocal microscope (Leica, Wetzlar, Germany). The ratio of red to green fluorescence intensity was calculated to determine $\mathrm{\Delta }$$\mathrm{\Psi }$m.

Total DNA was extracted from HK-2 cells using Qiagen DNeasy Blood and Tissue Kits (Product No. 69504; Hilden, Germany) according to standard methods. To determine mitochondrial DNA (mtDNA) copy number, mtDNA was quantified by normalizing the mitochondrial-encoded gene ND1 to the nuclear-encoded gene B2M using RT-PCR and the $\mathrm{\Delta }$$\mathrm{\Delta }$Ct method. The sequences of primer pairs were as follows: 5^′-CCCTAAAACCCGCCACATCT-3^′ (F, ND1); 5^′-GAGCGATGGTGAGAGCTAAGGT-3^′ (R, ND1); 5^′-GCTCGCGCTACTCTCTCTTT-3^′ (F, B2M); 5^′-TCATCCAATCCAAATGCGGC-3^′ (R, B2M).

The mtDNA copy number was normalized to that of B2M to calculate its relative value.

Concentrations of interleukin-1$\beta$ (IL-1$\beta$) and interleukin-18 (IL-18) in the supernatants of cell cultures were measured using commercially available ELISA kits (Elabscience, Wuhan, Hubei, China; Product No. E-EL-H0149 and E-EL-H0253) following the guidelines provided by the manufacturer. Briefly, supernatants were added to antibody-precoated wells and incubated for 1.5 hours at 37 °C. Once the washing step was completed, the biotinylated detection antibody and the streptavidin-biotin-peroxidase conjugate were added in a sequential manner. The reaction was stopped after incubation with the chromogenic substrate for 30 min. The absorbance at 450 nm was measured using a multifunctional microplate reader (BioTek, Santa Clara, CA, USA).

Data are presented as mean $\pm$ standard deviation (SD). Statistical analysis was performed using GraphPad Prism 8 (GraphPad Software, San Diego, CA, USA) with two-way analysis of variance (ANOVA) followed by Bonferroni’s test for multiple comparisons or Student’s t-test for comparison between two groups. All experiments were independently replicated at least thrice. p 0.05 was considered statistically significant.

To establish a hyperuricemic cell injury model, HK-2 cells were subjected to different concentrations of UA (0, 1, 2, 3, and 4 mM) for a duration of 36 h. The CCK-8 assay was employed to evaluate the viability of the cells. The data clearly demonstrated a dose-dependent decline in cell viability as UA concentration increased (Supplementary Fig. 1). Among the tested concentrations, treatment with 3 mM UA led to a reduction of cell viability by approximately 50%, which corresponded to the half-maximal inhibitory concentration. Therefore, this concentration was selected for subsequent experiments, as it effectively represents a condition where significant cell injury is induced while maintaining a measurable cell population for further analysis.

Next, to evaluate the effect of UA on cell membrane integrity, we compared the UA-treated group with the control group. Propidium iodide staining was used because PI can only enter cells with compromised membranes. The results showed a marked increase in the number of PI-positive cells in UA-treated groups (Fig. 1A,B). This substantial increase in PI-positive cells clearly indicated that the cell membranes in the UA-treated group were damaged, resulting in the loss of their normal barrier function.

Fig. 1.

UA triggers HK-2 cell injury, oxidative stress, and inflammatory response. (A) PI/Hoechst double-staining fluorescence images showing cell membrane damage in the UA-treated group (3 mM) (PI red, nuclei blue). Scale bar: 50 µm. (B) Quantitative analysis of PI-positive cells/signals. (C) Quantitative results of intracellular ROS levels detected by DCFH-DA fluorescence probe. (D) Protein expression levels of inflammatory factors IL-1$\beta$ and IL-18 detected by ELISA. UA, Uric acid; HK-2, Human renal proximal tubular epithelial cells; PI, Propidium iodide; ROS, Reactive oxygen species; DCFH-DA, 2’,7’-Dichlorodihydrofluorescein diacetate; IL-1$\beta$, Interleukin-1$\beta$; IL-18, Interleukin-18. The data are presented as the mean $\pm$ SD with a sample size of n = 3. Statistics were done using Student’s t-test. **p 0.01 and ***p 0.001.

In addition to cell membrane integrity, we investigated oxidative stress and inflammatory responses. ROS levels were measured, and the results showed a significant increase in the UA-treated group compared to that in the control group (Fig. 1C). Moreover, the expression of inflammatory factors such as IL-1$\beta$ and IL-18 was also significantly upregulated (Fig. 1D). Collectively, these findings confirmed the successful establishment of the UA-induced HK-2 cell injury model, as the observed impairment of cell membrane integrity, increased ROS levels, and upregulation of inflammatory factors are all characteristic features of cell injury.

To verify the pivotal role of oxidative stress in UA-mediated injury, we used the antioxidant, NACA. As shown in Supplementary Fig. 2A, UA treatment significantly reduced HK-2 cell viability, whereas NACA pretreatment (UA + NACA) markedly restored cell viability. NACA alone did not exhibit cytotoxicity. Consistent with the viability assay results, UA stimulation led to a substantial increase in intracellular ROS levels (Supplementary Fig. 2B). This oxidative burst was effectively suppressed by NACA pretreatment. These results demonstrate that UA-induced oxidative stress is the primary cause of cytotoxicity.

To further explore the mechanisms underlying UA-induced cell injury, we focused on mitochondrial dynamics, specifically the role of DRP1, a key regulator of mitochondrial fission. RT-qPCR was performed to examine the mRNA expression levels of DRP1 and FIS1. The results revealed that UA treatment significantly upregulated the mRNA expression of both DRP1 and FIS1 (Fig. 2A), suggesting that UA initiates mitochondrial fission at the transcriptional level.

Fig. 2.

UA promotes DRP1 activation and mitochondrial translocation. (A) RT-qPCR detection of DRP1 and FIS1 mRNA expression levels. (B) Western blot detection of total DRP1 and phosphorylated DRP1 (p-DRP1 Ser616) protein expression. (C,D) Quantitative analysis of DRP1 and p-DRP1 protein expression in (B). (E) Western blot detection of DRP1 enrichment in the mitochondrial fraction. (F,G) Quantitative analysis of DRP1 distribution in cytosolic and mitochondrial fractions. UA, Uric acid; DRP1, Dynamin-related protein 1; RT-qPCR, Reverse transcription quantitative polymerase chain reaction; FIS1, Mitochondrial fission 1 protein; p-DRP1, Phosphorylated DRP1. The data are presented as the mean $\pm$ SD with a sample size of n = 3. Statistics were done using Student’s t-test. *p 0.05, **p 0.01, and ^#not significant.

At the protein level, Western blot analysis was carried out to assess the expression of total DRP1 and its activated form, phosphorylated DRP1 at Ser616 (p-DRP1 Ser616). These results indicated that UA treatment significantly increased the expression of both total DRP1 and p-DRP1 Ser616. Notably, the increase in p-DRP1 Ser616 was more pronounced than that in total DRP1 (Fig. 2B–D). This differential increase suggests that UA not only enhances the overall expression of DRP1, but also promotes its activation, which is crucial for its function in mitochondrial fission.

Since the activation of DRP1 is known to mediate its translocation to the mitochondria, we investigated the subcellular distribution of DRP1. Cellular fractionation, a technique that separates cellular compartments based on their physical properties, was combined with Western blot analysis. This approach allowed quantification of DRP1 in the mitochondrial fraction. The outcomes indicated that UA treatment notably enhanced the enrichment of DRP1 in mitochondria (Fig. 2E–G). This finding provides direct evidence that UA treatment promotes the mitochondrial translocation of DRP1, which is likely to be of crucial importance in the mitochondrial fission event and contribute to UA-induced cell injury.

To explore the significance of the DRP1/FIS1 interaction in UA-induced cell injury, we used the specific peptide inhibitor P110. Co-IP assays were carried out, and the results clearly demonstrated that P110 treatment effectively inhibited the UA-induced protein interaction between DRP1 and FIS1 (Fig. 3A,B). This finding serves as an initial indication that P110 can disrupt relevant molecular interactions associated with cell injury.

Fig. 3.

P110 inhibits DRP1/FIS1 interaction and DRP1 mitochondrial translocation. (A) Co-IP analysis of the interaction between DRP1 and FIS1. (B) Quantitative analysis of DRP1 and FIS1 binding levels. (C) Western blot detection of DRP1 distribution in mitochondria. (D,E) Quantitative analysis of DRP1 protein levels in mitochondria and cytoplasm. Input: 10% total lysate; IgG, negative control; IP, anti-FIS1 immunoprecipitated; DRP1, Dynamin-related protein 1; FIS1, Mitochondrial fission 1 protein; Co-IP, Co-immunoprecipitation. The data are presented as the mean $\pm$ SD with a sample size of n = 3. Statistics were done using two-way ANOVA followed by Bonferroni’s test. *p 0.05, **p 0.01, and ^#not significant.

Subsequently, we investigated the impact of P110 on the intracellular distribution of DRP1. Western blot analysis revealed that P110 did not affect the total expression of DRP1. However, it significantly reduced the distribution of DRP1 in the mitochondrial fraction (Fig. 3C–E). This reduction in mitochondrial DRP1 levels indicates that P110 effectively blocks the mitochondrial translocation of DRP1, which is likely to be an important step in alleviating UA-induced cell injury.

In terms of cellular function, the P110 treatment had a significant positive effect on cell viability. As shown in Fig. 4A, P110 significantly improved the UA-induced reduction in cell viability, suggesting a cytoprotective effect. In the context of oxidative stress, P110 effectively counteracted UA-induced excessive ROS induced by UA stimulation (Fig. 4B). Moreover, it significantly upregulated the expression of antioxidant enzymes including SOD2, HO-1, and NQO1 (Fig. 4C). These changes in antioxidant enzyme expression imply enhanced cellular antioxidant defense capacity, which is crucial for cell survival under oxidative stress conditions.

Fig. 4.

P110 alleviates UA-induced cell injury. (A) CCK-8 assay detecting cell viability under P110 or TAT treatment in UA-induced cell injury. The red box indicates that treatment with 1 µM of P110 significantly improved cell viability. Additionally, 1 µM of P110 will be used in the subsequent experiments. (B) Detection of intracellular ROS levels by DCFH-DA fluorescence probe. (C) RT-qPCR detection of antioxidant enzymes SOD2, HO-1, and NQO1 expression. (D) Western blot detection of inflammatory factors IL-1$\beta$ and IL-18 protein expression. (E) Quantitative analysis of protein expression in (D). UA, Uric acid; CCK-8, Cell Counting Kit-8; TAT, transactivator of transcription; ROS, Reactive oxygen species; DCFH-DA, 2^′,7^′-Dichlorodihydrofluorescein diacetate; RT-qPCR, Reverse transcription quantitative polymerase chain reaction; SOD2, Superoxide dismutase 2; HO-1, Heme oxygenase-1; NQO1, NAD(P)H quinone dehydrogenase 1; IL-1$\beta$, Interleukin-1$\beta$; IL-18, Interleukin-18. The data are presented as the mean $\pm$ SD with a sample size of n = 3. Statistics were done using two-way ANOVA followed by Bonferroni’s test. *p 0.05, **p 0.01, ***p 0.001, and ^#not significant.

Additionally, P110 treatment had a notable effect on the inflammatory response. It markedly suppressed the upregulation of inflammatory factors such as IL-1$\beta$ and IL-18 (Fig. 4D,E). In contrast, the control peptide, TAT, did not produce these protective effects. Collectively, these results strongly suggest that P110 can effectively alleviate UA-induced oxidative stress, inflammatory responses, and cell injury by specifically inhibiting DRP1/FIS1 interaction.

To gain an in-depth understanding of the cytoprotective mechanism of P110, we focused on its regulatory effects on mitochondrial homeostasis. RT-qPCR was performed to examine the expression of mitochondrial fusion proteins. The results showed that P110 treatment significantly reversed UA-induced downregulation of the mitochondrial fusion proteins MFN2 and OPA1 (Fig. 5A). This restoration of fusion protein expression indicates that P110 may play a role in promoting mitochondrial fusion, which is an important aspect of mitochondrial dynamics.

Fig. 5.

P110 improves mitochondrial fusion and restores mitochondrial homeostasis. (A) RT-qPCR detection of the mRNA expression levels of mitochondrial fusion proteins MFN2 and OPA1. (B) Transmission electron microscopy images showing mitochondrial ultrastructure. Red arrows indicate mitochondria. Scale bar: 200 nm. (C) Quantitative analysis of mitochondrial length. (D) JC-1 staining fluorescence images showing mitochondrial membrane potential (red: aggregates, green: monomers). Scale bar: 20 µm. (E) Quantitative analysis of JC-1 red/green fluorescence ratio. (F) RT-qPCR detection of mtDNA expression. P110, Mitochondrial division inhibitor 1; RT-qPCR, Reverse transcription quantitative polymerase chain reaction; MFN2, Mitofusin 2; OPA1, Optic atrophy 1; JC-1, 5,5^′,6,6^′-Tetrachloro-1,1^′,3,3^′-tetraethylbenzimidazolylcarbocyanine iodide; mtDNA, Mitochondrial DNA. The data are presented as the mean $\pm$ SD with a sample size of n = 3. Statistics were done using two-way ANOVA followed by Bonferroni’s test. *p 0.05, **p 0.01, ***p 0.001, and ^#not significant.

Transmission electron microscopy was used to directly observe morphological changes in the mitochondria. The images revealed that P110 intervention markedly alleviated UA-induced mitochondrial fragmentation and effectively increased mitochondrial length (Fig. 5B,C). These morphological changes suggest that P110 can improve the mitochondrial structure, which is closely related to its function.

Furthermore, we evaluated the effects of P110 on $\mathrm{\Delta }$$\mathrm{\Psi }$m and mtDNA content. P110 treatment effectively restored the mitochondrial membrane potential (Fig. 5D,E) and increased mtDNA content (Fig. 5F). These results indicate that P110, through specifically inhibiting the DRP1/FIS1 interaction and reducing DRP1 mitochondrial translocation, can improve mitochondrial dynamics and maintain the structural and functional integrity of mitochondria.

Mitochondrial dysfunction is a key event in intrinsic apoptotic pathway. To determine whether P110 exerts its protective effects via this pathway, we examined mitochondria-mediated apoptosis-related indicators. Western blotting was performed to assess the expression of apoptosis-related proteins. The results showed that UA treatment significantly upregulated the expression of pro-apoptotic proteins including BAX, CYT C, cleaved Caspase-9, and cleaved Caspase-3, and concurrently downregulated the level of the anti-apoptotic protein BCL-2. However, P110 treatment reversed these abnormal expression patterns (Fig. 6A,B).

Fig. 6.

P110 suppresses cell apoptosis through the mitochondrial pathway. (A) Western blot detection of apoptosis-related proteins BAX, BCL-2, CYT C, cleaved Caspase-9, and cleaved Caspase-3 expression. (B) Quantitative analysis of protein expression in (A). (C) Cell PI fluorescence staining images showing apoptosis. Scale bar: 50 µm. (D) Quantitative statistics of apoptosis rate. BAX, BCL2-associated X protein; BCL-2, B-cell lymphoma 2; CYT C, Cytochrome C; PI, Propidium iodide. The data are presented as the mean $\pm$ SD with a sample size of n = 3. Statistics were done using two-way ANOVA followed by Bonferroni’s test. *p 0.05, **p 0.01, and ^#not significant.

To further confirm the anti-apoptotic effect of P110, apoptosis was detected using PI staining. The results showed that P110 treatment significantly reduced UA-induced apoptosis (Fig. 6C,D). These findings suggest that P110 attenuates mitochondria-mediated apoptosis by inhibiting DRP1/FIS1 interaction, thereby protecting against UA-induced renal tubular epithelial cell injury.

In summary, this study comprehensively demonstrated that P110 can specifically inhibit the DRP1/FIS1 interaction, which in turn leads to a series of beneficial effects, including the alleviation of oxidative stress, restoration of mitochondrial homeostasis, and inhibition of apoptosis, ultimately protecting cells from UA-induced injury.