Circadian gene BMAL1 ameliorates renal ischaemia-reperfusion injury in diabetic mice by enhancing mitophagy via the HIF-1/BNIP3 pathway

Table 1 highlights that db/db mice exhibited markedly elevated blood glucose levels and increased body weight compared to db/+ mice. These db/db mice displayed classic features of type 2 diabetes, including polydipsia, polyphagia, and polyuria.

As illustrated in Fig. 1, mice in the DS group exhibited elevated serum creatinine (Scr) levels compared to the NS group, alongside an increase in blood urea nitrogen (BUN) levels. Similarly, the NI/R group demonstrated higher Scr and BUN levels compared to the NS group, indicating more pronounced renal dysfunction. Notably, following renal I/R, the DI/R group showed a substantial rise in both Scr and BUN levels relative to the NI/R group. These results underscore that renal I/R induces renal functional impairment, with diabetes further aggravating the damage caused by I/R.

In Fig. 1D, hematoxylin and eosin (HE) staining of renal tissue revealed that in the NS group of mice, the renal tubular structure appeared intact, exhibiting regular morphology without obvious swelling, expansion, degeneration, or necrosis. Conversely, in the DS group, there was a small amount of tubular dilation, hyaline degeneration and necrosis. The I/R group exhibited significant renal tubular epithelial cell damage, evidenced by vacuolar degeneration, cell shedding, tubular dilation, interstitial edema, cast formation, and infiltration of inflammatory cells. These pathological alterations were markedly more severe in the DI/R group compared to the NI/R group. Furthermore, Paller score analysis of renal tubular injury revealed that the I/R group had a substantially higher injury score than the sham surgery group, with the DI/R group demonstrating elevated scores relative to the NI/R group.

To evaluate apoptosis levels in kidney tissues across different mouse groups, we utilized TdT-mediated dUTP Nick-End Labeling (TUNEL) staining. This analysis revealed a significant increase in TUNEL-positive cells in the kidneys of the I/R group compared to the sham operation group. Similarly, the DI/R group displayed a notable rise in TUNEL-positive cells relative to the NI/R group (Fig. 1E, F). Additionally, Western blot analysis demonstrated elevated expression of cleaved caspase-3 (CC3) in both the NI/R and DI/R groups (Fig. 1G, H). Collectively, these findings suggest that diabetes exacerbates kidney damage induced by I/R.

To elucidate the alterations in BMAL1 expression and mitophagy during diabetic kidney I/R, we assessed the levels of BMAL1 and mitophagy-related proteins in the kidneys of mice across different experimental groups (Fig. 2A). Western blot analysis demonstrated a notable reduction in BMAL1 expression in the kidney I/R group compared to the sham operation group, indicating a disruption in the core clock genes’ activity (Fig. 2B). Concurrently, there was a marked increase in microtubule-associated protein 1 light chain 3B (LC3B) II levels and a decrease in p62, reflecting enhanced autophagy (Fig. 2E, F). Additionally, we observed a significant decline in mitochondrial membrane proteins translocase of outer mitochondrial membrane 20 (TOMM20) and cytochrome c oxidase IV (COX IV), suggesting increased mitochondrial degradation (Fig. 2G, H). Furthermore, there was a substantial upregulation of HIF-1α and BNIP3, pointing to an elevated mitophagy process mediated by the HIF-1α/BNIP3 pathway in the kidneys following I/R (Fig. 2C, D).

Compared to the non-diabetic cohort, the diabetic group exhibited reduced expression levels of BMAL1, HIF-1α, BNIP3, and LC3B II, while p62, TOMM20, and COX IV levels were significantly elevated. These findings suggest that renal I/R induces a downregulation of BMAL1 and activates the HIF-1α/BNIP3-mediated mitophagy pathway. Furthermore, diabetes exacerbates the suppression of BMAL1 expression, leading to a concomitant decline in mitophagy. Therefore, we speculate that renal I/RI responsive activates mitophagy, while in a high/glucose environment, BMAL1 expression is additionally suppressed, and endogenous protective mechanisms are deactivated, leading to a decrease in mitophagy levels and further aggravation of renal damage.

To evaluate mitochondrial morphology and mitophagy activation, we employed transmission electron microscopy to examine kidney samples from each experimental group, and assessed mitochondrial number, area, and length (Fig. 2I-L). As illustrated in Fig. 2I-L, our results confirm the Western blot findings, revealing a significant increase in the number of autophagosomes and mitophagosomes in the kidneys of mice subjected to I/R, while the number of mitochondria decreased, indicating an enhanced mitophagic response to renal ischemia-reperfusion. Furthermore, compared to the NS group, the DS group exhibited marked lipid droplet accumulation in renal tubular epithelial cells, with a reduction in the number, area, and length of mitochondria. In the I/R group, mitochondrial integrity was significantly compromised, with a decrease in mitochondrial area and length, and rupture, swelling, and fragmentation of the cristae were observed. Additionally, the DI/R group showed a significant increase in the number of damaged mitochondria, further exacerbating mitochondrial injury.

Cell viability was evaluated using the Cell Counting Kit-8 (CCK-8) assay, while Western blotting was employed to quantify CC3 as a marker of apoptosis across experimental groups. Our results demonstrate that both HG and H/R conditions lead to cellular damage, decreased viability, and increased apoptosis. Particularly, H/R alone caused more severe cellular injury, with the HG + H/R group exhibiting the most significant damage (Fig. 3A, B, C). Mitochondrial function was assessed by measuring adenosine triphosphate (ATP) levels, reactive oxygen species (ROS) production, mitochondrial ROS and mitochondrial membrane potential (MMP) (Fig. 3D-J). Intracellular ROS and mitochondrial ROS levels were determined by the intensity of fluorescence from dichlorofluorescein (DCF) and MitoSox Red, with increased fluorescence indicating higher ROS levels. MMP alterations were visualized using JC-1 staining: red fluorescence denoted healthy mitochondria, while green fluorescence indicated a loss of MMP, with an increased green/red fluorescence ratio reflecting mitochondrial damage. Our findings reveal that both HG and H/R conditions result in diminished ATP levels and MMP, alongside elevated ROS levels. Notably, the HG + H/R condition resulted in the most pronounced decline in ATP and MMP, and the highest ROS levels. These results suggest that HG exacerbates H/R-induced mitochondrial dysfunction and damage in TCMK-1 cells, corroborating previous animal model studies. Additionally, we performed fluorescence colocalization analysis using Mito-Tracker Green and Lyso-Tracker Red to visualize mitochondria and lysosomes, respectively. The results showed an increase in mitochondria-lysosome fusion events in the hypoxia-reoxygenation group, while a decrease was observed in the high-glucose group, suggesting that mitophagy is inhibited under high-glucose conditions.

To investigate the effects of HG and H/R on cellular processes, we employed an in vitro model using TCMK-1 cells. Western blot analysis (Fig. 4) demonstrated that in cells subjected to NH/R, there was a significant increase in the expression levels of HIF-1α, BNIP3, and LC3B II compared to the NC group, while BMAL1, p62, TOMM20, and COX IV levels were notably decreased. This suggests that H/R induces a downregulation of BMAL1 and activates mitophagy in TCMK-1 cells. Conversely, in cells exposed to HG + H/R, the levels of BMAL1, HIF-1α, BNIP3, and LC3B II were diminished relative to the NH/R group, whereas p62, TOMM20, and COX IV levels were elevated. These results indicate that high glucose further inhibits BMAL1 expression and diminishes mitophagy, corroborating our animal experiment findings.

To elucidate the impact of BMAL1 on mitophagy through the HIF-1α/BNIP3 pathway under HG and H/R conditions, we transfected TCMK-1 cells with an BMAL1-OE lentiviral vector to generate stable BMAL1-overexpressing cells (OEB group), using the BMAL1-NC empty vector as a negative control (Vector group). As illustrated in Fig. 5, BMAL1 expression was markedly higher in the OEB group compared to the Vector group. This upregulation was accompanied by increased levels of HIF-1α, BNIP3, and LC3B II, while levels of p62, TOMM20, and COX IV were notably decreased. These findings indicate that BMAL1 overexpression augments mitophagy through the HIF-1α/BNIP3 signaling pathway.

To elucidate the impact of BMAL1 expression on TCMK-1 cells subjected to HG conditions during H/R injury, we evaluated alterations in cell viability, apoptosis, and mitochondrial function. Both HG and H/R treatments led to reduced cell viability and ATP production, alongside increased apoptosis. However, BMAL1 overexpression mitigated these effects, restoring cell viability and ATP levels while decreasing apoptosis (Figs. 6A, B, C, D). Additionally, elevated BMAL1 expression was associated with a reduction in ROS generation and attenuation of MMP damage induced by HG and H/R, as well as an increase in mitochondrial-lysosomal fusion events (Figs. 6E-L). These results suggest that BMAL1 overexpression ameliorates cellular injury and mitochondrial dysfunction in the context of HG and H/R exposure. These results suggest that under HG and H/R exposure, BMAL1 overexpression can alleviate cellular damage and mitochondrial dysfunction and partially restore the level of mitophagy.

To further explore whether BMAL1 overexpression enhances mitophagy through the HIF-1α/BNIP3 pathway, we treated cells with the HIF-1α inhibitor PX-478 (10 µM). As shown in Fig. 7, the PX-478-treated group exhibited a marked reduction in HIF-1α levels compared to both the HG + OEB and HG + OEB + H/R groups. This decrease in HIF-1α was accompanied by diminished levels of BNIP3 and LC3B II. In contrast, there was a notable increase in the expression of p62, TOMM20, and COX IV, suggesting that the inhibitory effect of PX-478 on HIF-1α impairs the BMAL1-induced mitophagy. It is worth noting that PX-478 has no significant effect on the expression of BMAL1.

To explore whether BMAL1 overexpression facilitates mitophagy through the HIF-1α/BNIP3 pathway to protect cells under high glucose and hypoxia-reoxygenation conditions, we re-evaluated cell viability, CC3 expression, and mitochondrial function. As illustrated in Fig. 8, cells in the PX-478 group exhibited reduced viability and ATP content, alongside increased ROS production, compromised MMP, decreased mitochondrial-lysosomal fusion events, and exacerbated apoptosis. These findings suggest that BMAL1 overexpression mitigates cellular damage induced by high glucose and hypoxia-reoxygenation by enhancing HIF-1α/BNIP3-mediated mitophagy.

The tubular epithelial cell in mouse (TCMK-1 cells), from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China), were cultured in Minimum Essential Medium (MEM, Hyclone, Beijing, China) containing 10% fetal bovine serum (FBS, Bioexplorer, USA) in the incubator with 5% CO2 at 37 °C.

In the H/R group, cells were first incubated in serum-free MEM medium for 12 h, followed by exposure to MEM medium supplemented with 10% FBS under hypoxic conditions (94% N2 + 5% CO2) for 24 h. Subsequently, the cells were transferred to a standard incubator with 5% CO2 for 12 h of reoxygenation. In contrast, the NC group cells were continuously maintained in a standard incubator. The HG group cells were cultured in MEM medium containing high glucose (30 mM).

The lentivirus was purchased from OBiO Technology(Shanghai). The lentiviral expression vector designated BMAL1-OE(pSLenti-EF1-P2A-Puro-CMV-ARNTL-3xFlag-WPRE) was used for BMAL1 gene (NM_001297719.2↗) delivery and stable overexpression. The empty vector designated BMAL1-NC (pSLenti-EF1-P2A-Puro-CMVMCS-3xFlag-WPRE) was used as a negative control. The MOI was 20. According to the manufacturer’s protocol, puromycin (50 µg/ml) was used to screen uninfected cells, and the surviving cells were further cultured and expanded. The successful overexpression of BMAL1 was subsequently validated through Western blot analysis.

The HIF-1α inhibitor PX-478 (S-2-amino-3-[4′-N, N,-bis(chloroethyl)amino]phenyl propionic acid N-oxide dihydrochloride, HY-10231, MedChemExpress, Shanghai, China) was prepared by dissolving 5 mg of PX-478 in 1 mL of dimethyl sulfoxide (DMSO; Sigma-Aldrich, USA) to create a stock solution. Subsequently, 40 µL of this stock solution was diluted into 50 mL of culture medium, resulting in a final concentration of 10 µM PX-478.

All animal experiments conducted in this study adhered to the Laboratory Animal Care guidelines set forth by Wuhan University and received approval from the Committee for the Use of Live Animals in Teaching and Research. Additionally, the study is reported in accordance with ARRIVE guidelines⁵³. We procured twelve eight-week-old male db/db (BKS.Cg-Dock7m+/+Leprdb/J) mice and twelve eight-week-old male non-diabetic (db/+) mice from Shulaibao (Wuhan) Biotechnology Co., Ltd. The mice were maintained at the Animal Experimental Center of Renmin Hospital, Wuhan University, under controlled environmental conditions, including a temperature of 22 ± 2 °C and regulated humidity. They were kept on a 12-hour light/dark cycle.

Bilateral renal I/R injury was induced in anesthetized mice. Briefly, mice were first weighed and then anesthetized with an intraperitoneal injection of pentobarbital sodium (50 mg/kg)⁵⁴. A mid-abdominal incision was made to expose the bilateral renal arteries, which were then clamped for 30 min to induce ischemia. After this period, blood flow was restored to the kidneys, and the mice were euthanized by using an excessive amount of anesthetic to collect blood and tissue samples for subsequent analyses. Mice with unclamped renal arteries served as controls for the no I/R condition.

For the in vivo studies, db/+ mice were allocated into two groups: the normal sham (NS group, n = 6) and the normal ischemia/reperfusion (NI/R group, n = 6). Similarly, db/db mice were divided into the diabetes sham (DS group, n = 6) and the diabetes ischemia/reperfusion (DI/R group, n = 6).

In the in vitro experiments, TCMK-1 cells were assigned to one of four conditions: negative control (NC group), high glucose control (HG group), negative control with hypoxia/reoxygenation (NH/R group), and high glucose with hypoxia/reoxygenation (HG + H/R group). Cells were further categorized based on the presence of BMAL1 overexpression lentivirus transfection into four distinct groups: high glucose with empty vector control (HG + Vector group), high glucose with BMAL1 overexpression (HG + OEB group), high glucose with empty vector control and hypoxia/reoxygenation (HG + Vector + H/R group), and high glucose with BMAL1 overexpression and hypoxia/reoxygenation (HG + OEB + H/R group). Additionally, the effect of PX-478 treatment was assessed by dividing cells into: high glucose with BMAL1 overexpression and no PX-478 (HG + OEB group), high glucose with BMAL1 overexpression and PX-478 (HG + OEB + PX-478 group), high glucose with BMAL1 overexpression and hypoxia/reoxygenation (HG + OEB + H/R group), and high glucose with BMAL1 overexpression, PX-478, and hypoxia/reoxygenation (HG + OEB + PX-478 + H/R group).

Cell viability was evaluated using the CCK-8 assay (CK04, Dojindo, Kumamoto, Japan) following the manufacturer’s protocol. Briefly, TCMK-1 cells were seeded in 96-well plates at 5 × 10³ cells per well. After exposure to hypoxia/reoxygenation, 10 µL of CCK-8 solution was added to each well, and cells were incubated for 3 h at 37 °C. Hydrogen peroxide (H₂O₂) was used as a positive control to induce cytotoxicity, with untreated cells serving as a negative control (Figure S9). Absorbance at 450 nm was subsequently measured with a microplate reader to determine cell viability.

Intracellular ROS in TCMK-1 cells were quantified using dichloro-dihydro-fluorescein diacetate (DCFH-DA). TCMK-1 cells were seeded in 6-well plates at 2 × 10⁵ cells per well. The cells in each well were incubated with 2 mL of 10 µM DCFH-DA (S0033, Beyotime, Shanghai, China) for 30 min at 37 °C in darkness, followed by three washes with serum-free medium. The same method was then employed to measure mitochondrial ROS, substituting DCFH-DA probe with MitoSOX Red (5 µM, MCE, USA). We applied treatments with H₂O₂ to increase ROS levels significantly, and used untreated samples as negative controls to demonstrate baseline ROS levels (Figure S9). Fluorescence images were captured using an Olympus fluorescence microscope (Tokyo, Japan). The relative fluorescence intensity was assessed using ImageJ software (Version 1.53).

ATP content was quantified using an ATP microplate assay kit (abs580117, Absin, Shanghai, China) following the manufacturer’s guidelines. A reduction in ATP levels serves as an indicator of cellular injury, reflecting potential mitochondrial dysfunction or impairment. Oligomycin was used as a positive control to inhibit ATP synthase and decrease ATP levels, alongside untreated samples as negative controls (Figure S9).

MMP was evaluated using a JC-1 MMP assay kit (C2003, Beyotime), following the manufacturer’s protocol. TCMK-1 cells were seeded in 6-well plates at 2 × 10⁵ cells per well. The cells in each well were treated with 1 ml of a JC-1 staining solution, prepared by diluting 5 µl of a 200X JC-1 stock solution with 1 ml of JC-1 staining buffer. This mixture was incubated for 20 min at 37 °C in the dark. Following incubation, cells were washed twice with JC-1 staining buffer. Carbonyl cyanide m-chlorophenyl hydrazone (CCCP) was used to induce mitochondrial depolarization as a positive control, with untreated cells serving as a negative control (Figure S9). Fluorescence microscopy was employed to assess the results. Elevated MMP is indicated by JC-1 aggregation within the mitochondrial matrix, resulting in red fluorescence, whereas diminished MMP is marked by the inability of JC-1 to aggregate, producing green fluorescence due to its monomeric form. MMP changes were quantified based on the relative intensity of red and green fluorescence.

After special treatment, the cells were labelled with mitochondria and lysosomes, which were tracked using commercial Mito-Tracker Green (C1048, Beyotime, Shanghai, China) and Lyso-Tracker Red (C1046, Beyotime, Shanghai, China), according to the manufacturer’s instructions.

SCr and BUN levels were quantified using a creatinine assay kit (EICT-100, BioAssay Systems) and a urea assay kit (DIUR-100, BioAssay Systems), respectively, following the manufacturer’s instructions.

Kidney samples were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned into 4 μm-thick slices. These sections were then stained with hematoxylin and eosin (HE). Tissue pathology was evaluated using an upright microscope, and tubular injury severity was quantified using the Paller renal tubular injury scoring system ²⁸. This system involves the random selection of 10 renal tubules per high-power field. Scores were assigned based on the following criteria: 1 point for tubular dilation, cell flattening, or swelling; 1 or 2 points for brush border injury or detachment; 2 points for the presence of casts; and 1 point for detached necrotic cells within the tubular lumen, provided these did not form casts or fragments.

For the detection of protein levels, the total protein was extracted using RIPA Lysis Buffer (GB2002, Servicebio, Wuhan, China) supplemented with a cocktail of protease inhibitor (GB2006, Servicebio, Wuhan, China), phosphatase inhibitor (GB2007, Servicebio, Wuhan, China), and phenylmethylsulfonyl fluoride (GB2008, Servicebio, Wuhan, China). The nuclear protein was isolated using NE-PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Scientific, 78833) to detect HIF-1α expression. Protein extracts from cells or renal cortical tissues were quantified using the BCA reagent (P0010S, Beyotime Biotechnology). Following separation by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), the proteins were transferred to a polyvinylidene difluoride membrane (Millipore, Billerica, MA, USA). The membrane was then blocked with 5% skim milk for 60 min. Subsequently, the membrane was incubated with primary antibodies, followed by horseradish peroxidase-conjugated secondary antibodies (AS003, AS014, Abclonal, Wuhan, China). The primary antibodies used were diluted as follows: HIF-1α (GB114936-100, Servicebio, Wuhan, China) at 1:1000, Histone H3 (ab1791, Abcam, Cambridge, FL, USA) at 1:5000, BMAL1 (14020, Cell Signaling Technology, Danvers, MA, USA) at 1:1000, LC3B (83506, Cell Signaling Technology, Danvers, MA, USA) at 1:1000, COX IV (4850, Cell Signaling Technology, Danvers, MA, USA) at 1:500, TOMM20 (42406, Cell Signaling Technology, Danvers, MA, USA) at 1:1000, p62 (39749, Cell Signaling Technology, Danvers, MA, USA) at 1:500, BNIP3 (ab109362, Abcam, Cambridge, FL, USA) at 1:1000, cleaved caspase 3 (CC3) (9661, Cell Signaling Technology, Danvers, MA, USA) at 1:1000, and β-actin (GB15001-100, Servicebio, Wuhan, China) at 1:3000. β-actin or Histone H3 was used as a loading control to ensure equal protein loading. Protein expression levels were quantified based on gray value analysis using ImageJ software (Version 1.53).

To assess apoptosis, paraffin-embedded renal sections were subjected to staining with a TUNEL kit (Roche, Basel, Switzerland), followed by counterstaining with 4′,6-diamidino-2-phenylindole (DAPI) for nuclear visualization. Apoptosis was quantified in accordance with the manufacturer’s guidelines. For each sample, ten random fields were examined to determine the apoptosis index.

Upon completion of the mouse modeling, the kidneys were harvested. Thin tissue slices, obtained from the interface of the renal cortex and medulla on ice, were immediately fixed in 2.5% glutaraldehyde. Following fixation, the samples were post-fixed in 1% osmium tetroxide, dehydrated through a graded ethanol series, and embedded in a hard resin. The embedded samples were then sectioned into ultra-thin slices of 80 nm. These sections were subjected to double staining with uranyl acetate and lead citrate, and subsequently examined using a transmission electron microscope (H-600, Hitachi) to evaluate mitochondrial morphology and quality. For all cases, 5–6 images were analyzed per sample and mitochondrial morphology per field were calculated. Each image represents 2–3 cells per field. Each cell contained ~ 50–100 mitochondria. Mitochondria length was obtained using the major axis as reported previously⁵⁵. TEM images were analyzed utilizing ImageJ software (Version 1.53).

Data are presented as mean ± SD. Statistical analysis was performed using GraphPad Prism software (Version 9.4.1, USA). Comparisons among multiple groups were conducted using one-way analysis of variance (ANOVA), with Tukey’s test employed for pairwise comparisons. A p-value of less than 0.05 was deemed statistically significant.