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Introduction

Acute pancreatitis (AP) is characterized by pancreatic acinar cell death and persistent local or systemic inflammation. The incidence of the disease is increasing and the mortality is high (1). Currently, the complex pathogenesis of AP is still unclear. Studies have confirmed that mitochondrial Ca2+ overload is associated with acinar cell death in AP (2,3). However, most of the current studies regarding the pathogenesis of AP primarily focus on acinar cells, while only a few studies focus on pancreatic ductal epithelial cells (PDECs), which is another important cell type closely related to AP.

PDECs play a key role in maintaining pancreatic homeostasis and avoiding substantial pancreatic inflammation (4-6). Studies have shown that transient high pressure, intracellular Ca2+ overload, and cytoskeletal changes of PDECs can aggravate AP (7-9). In addition, PDECs are an important component of the pancreatic ductal mucosal barrier (PDMB) and play an important role in preventing abnormal activation of trypsinogen and the reflux of digestive enzymes to the pancreas (5,10). Furthermore, impaired ductal fluid secretion and a lower intraluminal pH lead to zymogen activation and acinar cell dysfunction (11,12). Therefore, the decrease of PDMB function caused by PDECs damage may be the main cause of pancreatic injury and AP occurrence. Studying PDECs will help to uncover the underlying mechanisms of the development of AP.

A previous study from our group has identified intracellular Ca2+ overload in PDECs in AP (9). As one of the most important intracellular signal transduction molecules, Ca2+ is involved in almost every physiological activity within cells, including mitophagy. Mitophagy is the selective autophagy of mitochondria whereby dysfunctional mitochondria are transported to lysosomes for degradation (13) and it has a crucial role in mitochondrial quality and quantity control as well as cell survival. Mitochondrial Ca2+ overload is related to the mitochondrial structure, mitochondrial reactive oxygen species, mitochondrial membrane potential (MMP), and ATP production, which all contribute to the initiation of mitophagy (14).

Recent studies have shown that mitophagy within pancreatic acinar cells plays an important role in the pathogenesis and development of AP. This includes both PTEN-induced putative kinase 1 (PINK1)/Parkin-activated mitophagy and vacuole membrane protein 1-related mitophagy (15,16). However, whether similar mechanisms exist in PDECs remains to be elucidated. Intracellular Ca2+ overload in acinar cells leads to mitochondrial depolarization through the opening of the mitochondrial permeability transition pore (17) and these damaged mitochondria can be removed by mitophagy. However, the relationship between mitochondrial Ca2+ overload and mitophagy in PDECs during AP has not been investigated.

The mitochondrial calcium uniporter (MCU), as a highly selective Ca2+ uptake channel, transports Ca2+ into the mitochondrial matrix along an electrochemical gradient (18). As such, the MCU plays a crucial role in mitochondrial Ca2+ homeostasis and mitochondrial energy metabolism. There has been convincing evidence to suggest a close relationship between the MCU and mitophagy. Guan et al (19) found that upregulation of the MCU inhibited mitochondrial fusion and mitophagy through the calpain/optic atrophy type 1 signaling pathway, leading to myocardial ischemia-reperfusion injury. Chen et al (20) found that proliferation stimulated by apelin-13 in vascular smooth muscle cells was regulated by mitophagy that was caused by MCU-associated mitochondrial Ca2+ uptake. The MCU may be a key protein that links mitochondrial Ca2+ overload and mitophagy. A previous study by our group indicated that MCU expression and mitochondrial Ca2+ were increased in the rat model of AP (21). However, the exact mechanism by which MCU regulates mitophagy through mitochondrial Ca2+ regulation in AP remains to be elucidated.

Caerulein (CAE), an analog of cholecystokinin, has been widely used in vitro and in vivo to establish models of AP (22). Our previous studies found that the change in protein expression of autophagy marker, the apoptosis and pyroptosis protein expression of NLPR3, cleaved caspase-1 and cleaved caspase-3, the tight junction protein expression of ZO-1 and occludin in HPDE6-C7 cells treated with CAE in vitro were consistent with the rat model of AP (8,9,23,24). In the present study, it was hypothesized that MCU-associated mitochondrial Ca2+ overload promotes mitophagy by regulating the PINK1/Parkin pathway in AP. To test this hypothesis, CAE-treated HPDE6-C7 cells were used to imitate the pathological changes of PDECs in AP. In addition, ruthenium red (RR) is an inorganic polycationic dye that has been reported as an MCU inhibitor (25). It was first used to stain specific plant material and was subsequently found to be a voltage-sensitive calcium channel inhibitor (26,27). Therefore, RR was used as an MCU-specific inhibitor to assess the effect of MCU on mitophagy in the present study.

Materials and methods

Cell culture

Normal human pancreatic ductal epithelial cells (HPDE6-C7) were obtained from WHELAB (https://www.whelab.com/pro/580.html; cat. no. C1248; passage 3) and cultured in Dulbecco's Modified Eagle's Medium (DMEM; cat. no. C11995500BT; Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% fetal bovine serum (cat. no. S711-001S; Shanghai Shuangru Biotechnology Co., Ltd.), 100 U/ml penicillin, and 0.1 mg/ml streptomycin (cat. no. P1400; Beijing Solarbio Science & Technology Co., Ltd.). Cells were placed in an incubator with sterile humidified air with 5% CO2 at 37˚C. Cells were treated at the same time with CAE (100 nmol/l; cat. no. EY5113; Shanghai Amquar Biological Technology Co., Ltd.) and RR (10 µmol/l; cat. no. 00541; MilliporeSigma) for 24 h. CAE and RR were both dissolved in phosphate-buffered saline for the experiments. Cells were divided into four groups according to the type of treatment: CAE, RR, CAE+RR, and control.

Flow cytometry

Following treatment with RR (1, 5, 10, 50 and 100 µmol/l) for 24 h, the rate of apoptosis in HPDE6-C7 cells was detected using the annexin V-FITC/PI kit [cat. no. AT101; Multisciences (Lianke) Biotech Co., Ltd.], following the manufacturer's instructions. In brief, after digestion with an accutase solution, the cells were resuspended in binding buffer, then incubated with 5 µl of annexin V-FITC and 10 µl of PI for 5 min at 25˚C in an incubator. Cell apoptosis rates were calculated as the percentage of early + late apoptotic cells and measured by flow cytometry (FACSVerse, BD Biosciences) and scatter diagrams were visualized using FlowJo software version 10.8.1 (FlowJo LLC).

Cell Counting Kit (CCK-8) assay

HPDE6-C7 cells were plated in a 96-well plate and cultured for 6 h. After treatment with RR (1, 5, 10, 50 and 100 µmol/l) for 24 h at 37˚C, the cell medium was replaced with a mixture of fresh medium (90%) and CCK-8 reagent (10%). Cells were incubated for 1 h at 37˚C in the dark, and the absorbance was measured at 450 nm.

Western blotting

Protein was isolated from HPDEE6-C7 cells using RIPA lysis buffer (cat. no. R0020; Beijing Solarbio Science & Technology Co., Ltd.) supplemented with 1 mmol/l PMSF, and the protein concentration was measured by a BCA Protein Assay Kit (cat. no. P0012; Beyotime Institute of Biotechnology). Equivalent amounts of protein (50 µg/well) were separated by 10 or 15% SDS-PAGE. The gels were electroblotted onto PVDF membranes (162-0219; Bio-Rad Laboratories, Inc.). Next, membranes were placed in nonfat milk (cat. no. P0216; Beyotime Institute of Biotechnology) for 40 min at room temperature and incubated at 4˚C for 16-20 h with primary antibodies, including: MCU (cat. no. 14997S; Cell Signaling Technology, Inc.; 1:1,000), LC3 (cat. no. ab48394; Abcam; 1:1,000), SQSTM1/p62 (cat. no. ab109012; Abcam; 1:15,000), and translocase of the outer mitochondrial membrane complex subunit 20 (cat. no. TOMM20; ab186735; Abcam; 1:2,000), PINK1 (cat. no. ab300623; Abcam; 1:1,000), Parkin (cat. no. 14060-1-AP; Proteintech Group, Inc.; 1:1,000), β-Actin (cat. no. 4970S; Cell Signaling Technology, Inc.; 1:1,000), and GAPDH (cat. no. ab9485; Abcam; 1:2,500). The following day, the membranes were incubated with the DyLight 800-conjugated secondary anti-rabbit antibody for 1 h at room temperature (cat. no. 5151; Cell Signaling Technology, Inc.; 1:10,000). Proteins were visualized using the Odyssey infrared imaging system (LI-COR, Inc.), and the gray value of the protein band was analyzed using ImageJ software (version 1.53e; National Institutes of Health).

Transmission electron microscopy (TEM)

HPDE6-C7 cells were immersed in electron microscope fixation fluid for 2.5 h at 4˚C for fixation and stained in 1% osmium acid for 2 h at room temperature. After dehydration, cells were embedded in epoxy resin at 37˚C overnight. Epoxy resin samples were heated at 60˚C for 48 h, and slices (80 nm) were obtained using an ultramicrotome (EM UC7; Leica Microsystems GmbH). After counterstaining sequentially with 2% uranium acetate and 2.6% lead citrate for 8 min at room temperature, images were observed by TEM (HT7700; Hitachi High-Technologies Corporation).

Measurement of mitochondrial membrane potentials (MMP)

MMP changes in HPDE6-C7 cells were measured with an MMP assay kit and JC-1 (cat. no. C2006; Beyotime Institute of Biotechnology). Briefly, cells were loaded with the JC-1 working solution (X1) at 37˚C for 20 min. After washing with pre-cooled staining buffer, cells were immediately observed using fluorescence microscopy (Olympus Corporation). Fluorescent images were analyzed using ImageJ software (version 1.53e; National Institutes of Health).

Measurement of mitochondrial Ca2+

Levels of mitochondrial Ca2+ in HPDE6-C7 cells were identified using Rhod-2 AM (cat. no. 40776ES50, Shanghai Yeasen Biotechnology Co., Ltd.) and Mito-Tracker Green (MTG; cat. no. C1048; Beyotime Institute of Biotechnology). After washing with D-Hank's balanced salt solution (cat. no. H1046; Beijing Solarbio Science & Technology Co., Ltd.), cells were loaded with the Rhod-2 AM working solution (4 µmol/l) at 37˚C for 30 min. Subsequently, cells were loaded with MTG (200 nmol/l) at 37˚C for 30 min and immediately observed with a fluorescence microscope (Olympus Corporation). Fluorescent images were analyzed using ImageJ software (version 1.53e; National Institutes of Health).

Fluorescence colocalization assay of mitochondria and lysosomes

To visualize the formation of autolysosomes during mitophagy, MTG and Lyso-Tracker Red (LTR; cat. no. C1046; Beyotime Institute of Biotechnology) were used to label the mitochondria and lysosomes, respectively. Cells were cultured in confocal dishes (BS-15-GJM, Biosharp) and stained in medium containing 200 nmol/l MTG for 30 min and 75 nmol/l LTR for 15 min at 37˚C. Confocal fluorescent images were obtained using a laser scanning confocal microscope (TCS SP8 STED; Leica Microsystems GmbH) with a x63 oil immersion objective.

Immunofluorescence

At room temperature, HPDE6-C7 cells were fixed in 4% paraformaldehyde for 15 min, then permeabilized with 0.1% Triton X-100 for 5 min. After treatment with blocking buffer for 30 min, cells were incubated at 4˚C overnight with primary antibodies, including LC3 (cat. no. ab48394; Abcam; 1:1,000), Parkin (cat. no. 14060-1-AP; Proteintech Group, Inc.; 1:500), and TOMM20 (cat. no. 66777-1-Ig; Proteintech Group, Inc.; 1:500). Subsequently, cells were incubated with Alexa Fluor 555 (cat. no. 4413S; Cell Signaling Technology, Inc.; 1:1,000) and Alexa Fluor 488 (cat. no. 550036, Chengdu Zen-Bioscience Co., Ltd.; 1:1,000)-conjugated secondary antibodies for 1 h at room temperature in the dark. Fluorescent images were obtained using a fluorescence microscope (Carl Zeiss AG).

Statistical analysis

All experiments were repeated at least three times. Data were presented as the mean ± standard deviation, and SPSS 26 software (IBM Corp.) was used to perform all statistical analyses. Pearson's correlation coefficient (PCC) was determined as a statistic for quantifying colocalization using the ImageJ plugin Colocalization Finder. Student's t-test or one-way analysis of variance followed by Tukey's test was used to determine the statistical significance between indicated groups. P<0.05 was considered to indicate a statistically significant difference. Cartograms were generated using GraphPad Prism 9 software (Dotmatics).

Results

RR inhibited MCU expression in CAE-treated HPDE6-C7 cells

RR has been reported as a specific inhibitor of MCU (28). To identify a working nontoxic concentration of RR for the current study, the cytotoxic effect of RR in HPDE6-C7 cells was evaluated using a CCK-8 assay and flow cytometry. The results of the CCK-8 assay revealed decreased cell viability with higher concentrations of RR (50 or 100 µmol/l; Fig. 1A). Flow cytometry also confirmed that apoptosis was increased in cells exposed to 50 or 100 µmol/l RR (Fig. 1B and C). Thus, the maximum nontoxic concentration of RR (10 µmol/l) was used in subsequent experiments. Additionally, western blotting revealed that CAE increased the expression level of MCU and RR treatment restrained MCU expression in CAE-treated cells (Fig. 1D and E).

Figure 1 Effects of RR on the viability and apoptosis of HPDE6-C7 cells, and MCU expression in CAE-treated HPDE6-C7 cells. HPDE6-C7 cells were treated with RR (1, 5, 10, 50 and 100 µmol/l) for 24 h. (A) CCK-8 assay was used to detect cell viability. (B and C) Cell apoptosis was detected by flow cytometry. Q1: Necrotic cells, Q2: Late-apoptotic cells, Q3: Early-apoptotic cells, Q4: Live cells. HPDE6-C7 cells were treated with 10 µmol/l RR and 100 nmol/l CAE for 24 h. (D) Western blot and (E) semiquantitative analysis of MCU expression. Data were presented as the mean ± standard deviation. **P<0.01, ***P<0.001 vs. control. All experiments were repeated at least three times. RR, ruthenium red; MCU, mitochondrial calcium uniporter; CAE, caerulein.

Inhibition of MCU attenuated CAE-induced mitochondrial Ca2+ accumulation and MMP decline in HPDE6-C7 cells

The effect of CAE on mitochondrial Ca2+ levels and MMP in HPDE6-C7 cells was explored using fluorescence microscopy. The results of PCC indicated that the Rhod-2-loaded Ca2+ were strongly correlated with the Mito-Tracker (Fig. 2C). Therefore, it can be speculated that the Rhod-2-loaded Ca2+ may come from mitochondria. The results indicated that treatment of HPDE6-C7 cells with CAE overtly increased mitochondrial Ca2+ accumulation (Fig. 2A and B). Furthermore, cells were loaded with JC-1 to measure the MMP. The red/green fluorescence density was decreased in CAE-treated cells, indicating that treatment with CAE induced MMP depolarization (Fig. 2D and E). Notably, RR effectively attenuated mitochondrial Ca2+ accumulation and restored MMP in CAE-treated cells. These data demonstrated that CAE promotes MCU-associated mitochondrial Ca2+ overload and MMP decline.

Figure 2 RR treatment inhibited mitochondrial Ca2+ accumulation and reduced MMP decline in HPDE6-C7 cells. HPDE6-C7 cells were treated with 10 µmol/l RR and 100 nmol/l CAE for 24 h. (A and B) Changes in mitochondrial Ca2+ were detected by labeling with Rhod-2 AM and MTG and observed under a fluorescence microscope (magnification, x400; scale bar, 50 µm). (C) PCC was used to calculate the correlation between Rhod-2 AM and MTG. No correlation: PCC=0; weak: PCC <0.4; moderate: 0.4 ≤PCC <0.7; strong: 0.7 ≤PCC<1; complete or perfect: PCC=1. (D and E) Changes in MMP were detected by staining with JC-1 and observation under a fluorescence microscope (magnification, x200; scale bar, 50 µm). Data were presented as the mean ± standard deviation. **P<0.01, ***P<0.001 vs. control. All experiments were repeated at least three times. RR, ruthenium red; MMP, mitochondrial membrane potential; CAE, caerulein; MTG, Mito-Tracker Green; PCC, Pearson's correlation coefficient.

Inhibition of MCU attenuated CAE-induced mitophagy in HPDE6-C7 cells

The effect of MCU on mitophagy in AP was investigated by western blotting, immunofluorescence, fluorescence probes and TEM. The results of western blotting indicated that CAE improved the LC3-II/I ratio and reduced the expression of p62 and TOMM20. In turn, the LC3-II/I ratio was reduced and the expression of p62 and TOMM20 were increased by RR treatment in CAE-treated cells (Fig. 3A and B). In addition, the results of the immunofluorescence assays showed that the colocalization of LC3 and TOMM20 was increased in the CAE-treated cells, but this increased colocalization was decreased by treating the cells with RR (Fig. 3C and D). Moreover, the fluorescence images showed that the colocalization of mitochondria and lysosomes was increased in CAE-treated cells, which indicate an increased fusion of mitophagosomes and lysosomes. However, RR treatment reduced the overlap of mitochondrial and lysosomal staining (Fig. 4A and B). Furthermore, autophagosomes during mitophagy and impaired mitochondria were also detected by TEM and RR treatment reduced the formation of mitophagosomes and damaged mitochondria (Fig. 4C). These results demonstrated that elevated mitophagy in CAE-treated cells can be reduced by RR treatment.

Figure 3 RR treatment inhibited CAE-induced mitophagy in HPDE6-C7 cells. HPDE6-C7 cells were treated with 10 µmol/l RR and 100 nmol/l CAE for 24 h. (A and B) Expression of microtubule-associated proteins 1A/1B LC3B, p62 and TOMM20 were detected by western blotting. (C and D) TOMM20 (a mitochondrial marker) colocalization with LC3 was observed by fluorescence microscopy (magnification, x1,000; scale bar, 10 µm). The PCC was used to calculate the colocalization between LC3 and TOMM20. No correlation: PCC=0; weak: PCC <0.4; moderate: 0.4 ≤PCC <0.7; strong: 0.7 ≤PCC<1; complete or perfect: PCC=1. Data were presented as the mean ± standard deviation. *P<0.05, **P<0.01, ***P<0.001 vs. control. All experiments were repeated at least three times. RR, ruthenium red; CAE, caerulein; TOMM20, translocase of the outer mitochondrial membrane complex subunit 20; PCC, Pearson's correlation coefficient.

Figure 4 RR treatment reduced CAE-induced mitophagosome formation in HPDE6-C7 cells. (A and B) Mitochondrial colocalization with lysosomes was observed by fluorescence confocal microscopy (magnification, x630; scale bar, 10 µm). The PCC was used to calculate the colocalization between mitochondria and lysosomes. No correlation: PCC=0; weak: PCC <0.4; moderate: 0.4 ≤PCC <0.7; strong: 0.7 ≤PCC<1; complete or perfect: PCC=1. (C) Mitophagy and damaged mitochondria in HPDE6-C7 cells were observed by transmission electron microscopy. Black arrows indicate normal mitochondria. Red arrows indicate damaged mitochondria and the blue arrow indicates mitophagy (magnification, x15,000; scale bar, 1.0 µm). Data were presented as the mean ± standard deviation. **P<0.01 vs. control. All experiments were repeated at least three times. RR, ruthenium red; CAE, caerulein; TOMM20, translocase of the outer mitochondrial membrane complex subunit 20; PCC, Pearson's correlation coefficient; MTG, Mito-Tracker Green; LTR, Lyso-Tracker Red.

Inhibition of MCU attenuated PINK1/Parkin pathway activation in HPDE6-C7 cells

The effect of MCU on the PINK1/Parkin pathway in AP was investigated by western blotting and immunofluorescence. The results of western blotting showed that CAE increased the expression of PINK1 and Parkin, but this increased expression could be reversed by treating the cells with RR (Fig. 5A and B). Moreover, the immunofluorescence of Parkin and TOMM20 indicated that the expression of Parkin was increased in CAE-treated cells, demonstrating that CAE promoted the translocation of Parkin into the mitochondrial membrane. As expected, this effect was alleviated by RR treatment (Fig. 5C).

Figure 5 RR treatment decreased the expression of PINK1 and Parkin in CAE-treated HPDE6-C7 cells. HPDE6-C7 cells were treated with 10 µmol/l RR and 100 nmol/l CAE for 24 h. (A and B) Expression of PINK1 and Parkin was detected by western blotting. (C) Immunofluorescence of Parkin and TOMM20 (a mitochondrial marker) was observed by fluorescence microscopy (magnification, x200; scale bar, 50 µm). Data were presented as the mean ± standard deviation. **P<0.01, ***P<0.001 vs. control. All experiments were repeated at least three times. RR, ruthenium red; PINK1, putative kinase 1; CAE, caerulein; TOMM20, translocase of the outer mitochondrial membrane complex subunit 20.

Discussion

Previous studies have confirmed that mitophagy is involved in the pathogenesis of AP (15,29,30). In addition, MCU-associated mitochondrial Ca2+ overload plays a key role in AP (31,32). However, the effect of MCU on mitophagy in AP remains ambiguous. The present study was the first to investigate the regulatory role of MCU on mitophagy in AP, to the best of the authors' knowledge. The data indicate that activated MCU may promote mitophagy by regulating the PINK1/Parkin pathway in PDECs in AP in vitro.

Studies have demonstrated that mitochondrial Ca2+ overload plays a crucial role in the initiation of acinar cell death in AP (2,3,17). As an essential channel for mitochondrial Ca2+ uptake, the MCU may play an important role during mitophagy in AP. The results in the present study showed that CAE increased MCU overexpression, mitochondrial Ca2+ accumulation, and MMP depolarization. RR is a potent inhibitor of MCU (33). In the present study, the inhibition of the MCU by RR efficiently attenuated mitochondrial Ca2+ accumulation and restored MMP depolarization. These results demonstrated that reducing MCU-dependent mitochondrial Ca2+ uptake could improve MMP depolarization in PEDCs in AP. Notably, RR could not decrease the mitochondrial Ca2+ level in normal cells in the present study. Similarly, in our previous study of AP in rats, RR inhibited mitochondrial Ca2+ overload in the AP group, but did not reduce mitochondrial Ca2+ in the control group (21). Moreover, in Jiang et al (34), RR inhibited mitochondrial Ca2+ overload promoted by aconitine, but did not reduce mitochondrial Ca2+ in mouse hippocampal neuron HT22 cells. These phenomena may be due to the fact that RR does not interfere with the cytoplasmic Ca2+ signal propagation to the mitochondria in normal cells (27). In addition, the gating of MCU by the mitochondrial calcium uptake 1 may have a barrier effect on RR in normal cells (35).

Mitochondrial dysfunction and mitophagy have been extensively studied in acinar cells in AP (17,36,37). However, the presence of mitophagy in PDECs in AP and its underlying mechanisms remain unclear. LC3-II as a molecular marker of autophagy represents the relative number of autophagosomes and p62 as an autophagy substrate is negatively correlated with autophagy (38-40). A reduction in the expression of TOMM20, which is a translocase of the outer mitochondrial membrane 20, indicates that impaired mitochondria may be removed by mitophagy. Our previous study found that CAE induced autophagy activation in the early stage of AP (8). RR mainly inhibits mitochondrial Ca2+ and is recognized as a potent inhibitor of MCU. Therefore, RR may have a significant effect on mitophagy, which is one of the important forms of autophagy. In the present study, the elevated expression of LC3-Ⅱ and the decreased expression of p62 in CAE-treated cells suggested the presence and active process of autophagy. In addition, the increased colocalization of a mitochondrial marker (TOMM20) and LC3 as well as the decreased expression of TOMM20 in the present study suggested the increased level of mitophagy in CAE-treated cells. These results were supported by both the TEM and fluorescence microscopy results, where mitophagosomes were observed using TEM, and an increased colocalization of mitochondria and lysosomes was observed using fluorescence microscopy. These results indicated the presence of excessive mitophagy in PEDCs in this in vitro model of AP. Notably, according to the expression of TOMM20 and autophagy markers, changes in mitophagosomes and colocalization of TOMM20 and LC3 in CAE+RR-treated cells, RR was able to inhibit excessive mitophagy in PEDCs in AP. Similarly, Liu et al (41) found that the inhibition of the MCU attenuated excessive mitophagy and autophagic cell death by cadmium in liver cells. Yu et al (42) found that the inhibition of the MCU by Ru360 can inhibit hyperactivated mitophagy and protect neurocytes from ischemia/reperfusion injury. Another study has suggested that depolarized mitochondria are the initiating factor for mitophagy (43). In the present study, the MCU inhibitor RR decreased MCU expression, reduced mitochondrial Ca2+ overload and inhibited mitochondrial depolarization in PDECs in AP. Therefore, it can be hypothesized that the MCU can promote mitochondrial Ca2+ uptake and mitochondrial depolarization and further lead to mitophagy. However, whether RR exerts effects on other types of autophagy has not been reported and needs further study.

The PINK1/Parkin pathway plays an important role in the removal of dysfunctional mitochondria. First, PINK1 accumulates and is phosphorylated in the outer membrane of impaired mitochondria. Subsequently, phosphorylated PINK1 mediates Parkin translocation to the outer mitochondrial membrane and activates its ubiquitin ligase activity, causing the ubiquitination of a range of mitochondrial proteins and the formation of autophagosomes (13,43,44). Zhang et al (15) found that PINK1 and Parkin were elevated in the CAE-AP mouse model as well as the arginine-SAP mouse model at 24 h, but were decreased in the arginine-SAP mouse model at 48 and 72 h, suggesting that mitophagy is activated in the early stages of AP but inhibited in the late stages. The data in the present study showed that CAE promoted the expression of PINK1 and Parkin in HPDE6-C7 cells. In addition, the immunofluorescence results showed an increased translocation of Parkin to the mitochondrial outer membrane in CAE-treated cells. These changes were all reduced when the cells were treated with RR. Together, these results demonstrated that the MCU may promote mitophagy by regulating the PINK1/Parkin pathway in PDECs in AP. However, one limitation of the present study was the lack of manipulation experiments directly involving the PINK/Parkin pathway, such as the use of PINK1 inhibitors and knockout/knockdown of PINK1 or Parkin.

Either impaired mitophagy or excessive mitophagy has an important effect on cell death (45,46). The current study supported the hypothesis that mitochondrial Ca2+ overload by the MCU causes the accumulation of depolarized mitochondria and activates excessive mitophagy in PDECs in AP. Excessive mitophagy can lead to an imbalance in oxidative phosphorylation, oxidative stress and eventual cell death (47). Therefore, the present study provided a possible mechanism by which autophagic cell death of PDECs may disrupt the PDMB and aggravate AP. However, an important limitation of the present study was that cell death inhibitors, such as 3-MA (for autophagic cell death) (48), z-VAD (for apoptosis) (49) and Nec-1 (for necrosis) (50), were not used to detect the presence of autophagic cell death.

In addition to the inhibitor RR, the MCU activator spermine was used in the rat model of AP in our previous study (51). The results of the pre-experiment showed that spermine at different concentrations (2.5, 5.0, and 10.0 mg/kg) had no effect on the pathological score of pancreatic tissue and the amylase levels in the rat model of AP. In addition, RR effectively inhibits the MCU but also inhibits a range of other channels, including the two-pore-domain K+ channels (tandem pore domain acid-sensitive K+ channel 3, Twik-related K+ channel 2 and Twik-related arachidonic acid-stimulated K+ channel), transient receptor potential receptor vanilloid subfamily, calcium homeostasis modulator channels, ryanodine receptor, and Piezo channels (25,52). Therefore, a superior way to investigate the role of MCU would be to use MCU knockout mice or transfection of MCU short interfering RNA. Future work will assess the regulatory effect of the MCU on mitophagy in AP using the above methods.

In conclusion, the present study suggested that MCU-associated mitochondrial Ca2+ overload may induce mitophagy by regulating the PINK1/Parkin pathway in PDECs in AP in vitro. MCU-associated mitochondrial Ca2+ overload and mitophagy are attractive therapeutic targets for AP in the future.

Acknowledgements

Not applicable.

Funding

Funding: The present study was supported by grants from the National Natural Science Foundation of China (grant no. 81970558) and a Scientific Research Project of the Guangxi Administration of Traditional Chinese Medicine (grant no. GXZYA20220227).

Availability of data and materials

The data generated in the present study may be requested from the corresponding author.

Authors' contributions

YL and HYY were the main contributors to the present study. YL performed experimental work, analyzed data and wrote the manuscript. HYY revised the manuscript. GDT and HYY conceived the present study and obtained funding. NM and YYQ designed the experimental procedures. MTX, XLX and LL performed cell culture and collected experimental data. YL and GDT confirm the authenticity of all the raw data. All authors read and approved the final manuscript.

Ethics approval and consent to participate

Not applicable.

Patient consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

References