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Activated DRP1 promotes mitochondrial fission and induces glycolysis in ATII cells under hyperoxia

Respiratory Research volume 25, Article number: 443 (2024) Cite this article

Abstract

Backgroud

Recent studies have reported mitochondrial damage and metabolic dysregulation in BPD, but the changes in mitochondrial dynamics and glucose metabolic reprogramming in ATII cells and their regulatory relationship have not been reported.

Methods

Neonatal rats in this study were divided into model (FIO2:85%) and control (FIO2: 21%) groups. Lung tissues were extracted at 3, 7, 10 and 14 postnatal days and then conducted HE staining for histopathological observation. We assessed the expression of mitochondria dynamic associated proteins and glycolysis associated enzymes in lung tissues, primary ATII cells and RLE-6TN cells. Double immunofluorescence staining was used to confirm the co-localization of DRP1 and ATII cells. Real-time analyses of ECAR and OCR were performed with primary ATII cells using Seahorse XF96. ATP concentration was measured using an ATP kit. We treated RLE-6TN cells at 85% hyperoxia for 48 h with mitochondrial fission inhibitor Mdivi-1 to verify the role of DRP1 in regulating glucose metabolic reprogramming.

Findings

We found that hyperoxia causes ATII cells’ mitochondrial morphological change. The expression of DRP1 and p-DRP1 increased in lung tissue and primary ATII cells of neonatal rats exposed to hyperoxia. Glycolysis related enzymes including PFKM, HK2, and LDHA were also increased. Hyperoxia inhibited ATP production in ATII cells. In RLE-6TN cells, we verified that the administration of Mdivi-1 could alleviate the enhancement of aerobic glycolysis and fragmentation of mitochondria caused by hyperoxia.

Interpretations

Hyperoxia exposure leads to increased mitochondrial fission in ATII cells and mediates the reprogramming of glucose metabolism via the DRP1 signaling pathway. Inhibiting the activation of DRP1 signaling pathway may be a promising therapeutic target for BPD.

Introduction

Bronchopulmonary dysplasia (BPD) is a common respiratory complication in preterm infants. The incidence of BPD in preterm infants whose birth weight < 1500 g can reach up to 30% [1]. Infants with BPD face high mortality rates, prolonged hospital stays, heavy economic and social burdens, and long-term adverse outcomes in various systems, including the respiratory and nervous systems [2, 3]. To lower the incidence of BPD in preterm births and ameliorate poor prognosis, it is crucial to identify effective preventive, therapeutic, and predictive markers for BPD. Inflammation, oxidative stress, vascular malformation, malnourishment, and microecological imbalance are components of the complicated pathophysiology of BPD [4]. Dysregulated glucose, amino acid, and lipid metabolism, collectively known as metabolic reprogramming, may lead to the onset and progression of BPD [5]. Currently, research on metabolic reprogramming primarily focuses on tumors, whereas research on premature BPD is limited and mainly focuses on abnormalities in glucose metabolism. Under aerobic conditions, cells derive energy via glycolysis in the cytoplasm followed by oxidative phosphorylation in mitochondria. Under hypoxic conditions, cells predominantly use the glycolytic pathway for energy production instead of relying on oxygen-dependent oxidative phosphorylation. However, certain disease situations can alter cellular metabolism pathways.

The “Warburg effect” or “aerobic glycolysis” stand for the aberrant metabolic process that was first observed by Warburg, wherein tumor cells prone to undergo glycolysis in the cytoplasm even in aerobic settings [6]. Fanos et al. discovered that in the BPD group, urine lactic acid increased and gluconic acid decreased compared to that in the non-BPD group [7]. Kandasamy et al.’s study showed that mitochondrial oxygen consumption by umbilical vein endothelial cells was lower in infants with BPD than in preterm infants without BPD [8]. Animal experiments have demonstrated that hyperoxia-induced glycolysis and pentose phosphate pathways increase, whereas the rate of ATP production in neonatal mouse endothelial cells decreases [9, 10]. Collectively, these results suggest that BPD is characterized by an abnormal intracellular glucose metabolic pathway, accompanied by decreased mitochondrial aerobic respiration, which may be associated with mitochondrial damage.

The pathological basis for the occurrence and development of BPD is injury to type II alveolar epithelial (ATII) cells, which play a critical role in the secretion, regeneration, and differentiation of alveoli to maintain pulmonary homeostasis [11]. ATII cells have a dense population of mitochondria, which are vital organelles in ATP energy metabolism, biosynthesis, and signal transduction. The functional state of mitochondria is dependent on the dynamic balance between mitochondrial generation, fusion, fission and the degradation of injured mitochondria by mitochondrial autophagy, commonly referred to as mitochondrial dynamics. Changes in mitochondrial dynamics are key determinants of the onset and progression of lung diseases [12]. Mitochondria regulate their morphology, size, and quantity via continuous fusion and division, promoting membrane and content mixing among mitochondria and preserving high-quality mitochondrial DNA [13]. Gebb et al. observed that high oxygen levels lead to enhanced levels of mitochondrial reactive oxygen species and DNA damage in rat lung tissues [14]. The expression level of mitochondrial fission protein, DRP1, in the lung tissue of neonatal rats with BPD was significantly higher than that in the normoxia group, suggesting an increased level of mitochondrial division [15]. In vitro experiments have shown that the expression of the mitochondrial fusion proteins MFN1 and MFN2 in the RLE-6TN cell line decreases considerably under hyperoxic stimulation, whereas the DRP1 expression considerably increases [16].

In addition, post-translational modifications of DRP1, such as phosphorylation at Ser616 (p-DRP1), are important mechanisms regulating mitochondrial fission [17]. Mitochondrial division inhibits the tricarboxylic acid (TCA) cycle, consequently hindering energy generation. Additionally, the enhancement of the glycolysis pathway also leads to a decrease in energy generation. Therefore, in this study, we initiated our investigation with a focus on mitochondrial dynamic changes in ATII cells within the BPD model and hypothesized that DRP1 promotes glycolysis, thereby inducing the reprogramming of intracellular glucose metabolism during the pathogenesis of BPD.

Materials and methods

Antibodies and reagents

Anti-DRP1 (D6C7) rabbit monoclonal (8570, Cell Signalling Technology, Danvers, Massachusetts, USA), anti-Phospho-DRP1 (Ser616) polyclonal (3455, Cell Signalling Technology, Danvers, Massachusetts, USA), anti-MFN1 polyclonal (13798-1-AP, Proteintech, Wuhan, China), anti-MFN2 polyclonal (12186-1-AP, Proteintech, Wuhan, China), anti-OPA1 polyclonal (ab42364, Abcam, Cambridge, UK), anti-PFKM polyclonal (55028-1-AP, Proteintech, Wuhan, China), anti-hexokinase 2 polyclonal (22029-1-AP, Proteintech, Wuhan, China), anti-LDHA-specific polyclonal (19987-1-AP, Proteintech, Wuhan, China), mitochondrial fission factor (Mff) monoclonal (66527-1-Ig, Proteintech, Wuhan, China), anti-SPB (F-2) monoclonal (sc-133143, Santa Cruz Biotechnology, Dallas, Texas, USA), and anti-beta actin monoclonal antibodies (66009-1-lg, Proteintech, Wuhan, China) were used. Mdivi-1 was purchased from Sigma (M0199, Sigma-Aldrich, Missouri, USA). 2-DG was purchased from MCE (HY-13966, MedChemExpress, Shanghai, China).

Hyperoxia exposure animal model

Newborn Sprague Dawley rats in the model group (FiO2 = 0.85) were subjected to a hyperoxia environment within 12 h after birth, while the control group (FiO2 = 0.21) was treated to a normoxia environment, in accordance with our previously published protocols. Soda lime was used to keep the CO2 content at < 0.5%, while silica gel was utilized to remove water vapor from the oxygen tank. To minimize the influence of variations in nursing ability, the maternal rats were exchanged among cages every 24 h and were utilized to feed the newborn rats. Every day, the cage was opened for thirty minutes, and food and clean drinking water were provided. It was configured with a 12 h day/night cycle. Water and food were available at all times. At 3, 7, 10 and 14 d after the exposure to hyperoxia or normoxia, neonatal rats were anesthetized with isoflurane (R510-22, RWD, Shenzhen, China) inhalation anesthesia and then euthanized. The whole lungs were collected aseptically by opening the chest. The right lower lungs were preserved in paraformaldehyde for subsequent HE staining, whereas the other lung lobes were used for ATII cells isolation. Mature SD rats weighing 220–250 g were purchased from the BEIJING HFK BIOSCIENCE CO.LTD. The Ethics Committee of Animals at China Medical University approved and supervised all animal experiments (2023PS932K) [18].

Lung histology

Right lower lung tissues were fixed with 4% paraformaldehyde for 24 h, dehydrated with a graded series of ethanol, vitrified with xylene, and embedded in paraffin. Sections were cut (5 μm) and stained with HE. An optical microscope (80i, Nikon, Tokyo, Japan) was used to observe morphological changes. Three fields in each section were randomly selected for analysis. Using ImageJ software, alveolar development was evaluated by calculating the radial alveolar count (RAC), mean liner intercept (MLI), mean alveolar diameter (MAD), and alveolar wall thickness.

Isolation and purification of ATII cells

ATII cells were extracted from newborn rats on 3, 7, 10, and 14 d after birth. The lung tissues of newborn rats were sliced with scissors, perfused with normal saline, and then digested with 0.25% trypsin-EDTA (25200056, Gibco, Waltham, Massachusetts, USA) at 37 °C. Digestion was suspended after 25 min, and samples were filtered via 200 μm mesh filters. Following centrifugation at 800 rpm for 5 min, the cell pellet was resuspended for digestion in 0.1% collagenase I at 37 °C for 25 min (C8140, Solarbio, Beijing, China). The cells were harvested via centrifugation at 800 rpm for 5 min, resuspended in DMEM/F12 including penicillin/streptomycin (KGM12500-500, KeyGEN BioTECH, Jiangsu, China) and 10% fetal bovine serum (FSP500, Excell, Zhejiang, China), and incubated at 37 °C. Every 45 min, the culture dish was swapped out to eliminate the fibroblasts and unattached cells. Finally, foreign cells, such as fibroblasts and macrophages, were removed using an IgG-coated plate for differential attachment. Cells were harvested after 12–24 h for subsequent experiments [19].

Hyperoxia exposure cell model and treatment

Normoxia group RLE-6TN rat type II lung epithelial cells (BNCC337708, BeNa Culture Collection, Beijing, China) were cultured in RPMI 1640 medium supplemented with penicillin/streptomycin (KGM31800-500, KeyGEN BioTECH, Jiangsu, China) and 10% fetal bovine serum (164210-50, Procell, Wuhan, China) at 37 °C in the presence of 21% O2 and 5% CO2 for 24, 48, and 72 h. When the cell density in the normoxia group reached 70–80%, they were subjected to hyperoxia (85% O2 and 5% CO2) for 24, 48, and 72 h in the hyperoxia group [20]. For the Mdivi-1 treatment, cells were assigned to the Mdivi group at random and treated with Mdivi-1 (5 µM) for 48 h under hyperoxic conditions, with the culture medium being changed every 24 h.

Cell viability analysis

Cell Counting Kit-8 (CCK-8, HY-K030, MCE, New Jersey, USA) was used to assess cell viability. RLE-6TN cells were seeded in two 96-well plates and cultured for 24 h. Subsequently, one plate of cells was treated with varying concentrations (100, 500 nM, and 1, 5, 10, 50, and 100 µM) of Mdivi-1 for 48 h under hyperoxic conditions. Each well was then filled with 10 µL of diluted enhanced CCK-8 reagent in 100 µL of serum-free media. The 96-well plates were subsequently incubated for 2 h at 37 °C. Microplate reader (Cytation 5, Biotek, Vermont, USA) was used to determine optical density (OD) at 450 nm. The formula for cell viability (%) was as follows: (OD of treatment cells − blank OD) / (OD of control cells − blank OD) × 100.

Western blot analysis

RIPA buffer (P0013B, Beyotime, Shanghai, China), PMSF (ST507-10 ml, Beyotime, Shanghai, China), protease inhibitor cocktail (K1007, Ape×bio, San Diego, California, USA), and phosphatase inhibitor cocktail (K1015, Ape×bio, San Diego, California, USA) mixture was used to extract total protein from lung tissues, primary ATII, and RLE-6TN cells treated with Mdivi-1 following exposure to normoxia or hyperoxia. After sodium dodecyl sulfate-polyacrylamide gel electrophoresis, separated proteins were transferred onto a polyvinylidene fluoride membrane (PVDF). 5% skim milk was used to block the PVDF membrane and incubated for 16 h at 4 ℃ with various antibodies [anti-DRP1 (D6C7) rabbit monoclonal (1:1000), anti-Phospho-DRP1 (Ser616) polyclonal (1:1000), anti-MFN1 polyclonal (1:500), anti-MFN2 polyclonal (1:5000), anti-OPA1 polyclonal (1:1000), anti-PFKM polyclonal (1:1000), anti-hexokinase 2 polyclonal (1:3000), anti-LDHA-specific polyclonal (1:3000), and anti-beta actin monoclonal antibodies (1:40000)]. Thereafter, the PVDF membrane was incubated with a secondary antibody for 2 h (SA00001-1, SA00001-2, Proteintech, Wuhan, China) and washed three times in Tris-buffered saline containing 0.1% Tween-20 (TBST). Finally, protein bands were detected using enhanced chemiluminescence (180–501, Tanon, Shanghai, China). Using Image J software, band intensities were normalized to the beta-actin signal.

Metabolic flux analysis

By following the manufacturer’s instructions, the Seahorse XF96 analyzer (Agilent Technologies, Santa Clara, California, USA) was used to measure the levels of oxidative phosphorylation (OXPHOS) and metabolic flux. The cell culture plate was opened on a clean table, and 80 µL of cells were added to each well, with an inoculation density of 10,000 cells/well for ATII cells and 5000 cells/well for RLE-6TN cells. The medium was given to the background well, and the plate was placed on a clean workbench for 1 h. Thereafter, it was cultured 20 h in a 5% CO2 cell incubator at 37 °C to allow the cells to adhere to the surface. Before the assay, the cell density in each well was approximately 80%. Meanwhile, the probe plates were hydrated and placed for 20 h in a CO2-free cell incubator at 37 ℃. The Seahorse XF96 analyzer was turned on for at least 5 h prior to the experiment. The next day, the plate was put in the CO2-free cell incubator at 37 °C after the detection media had been added to the cell growth medium to reach a final volume of 180 µL per well. Subsequently, the probe plate was removed, and the drug was added to the dosing port. Thereafter, the plate was placed into the Seahorse XF96 analyzer for calibration, and the cell plate was used for detection. For the glycolysis stress test, the detection medium was prepared by mixing the Seahorse XF DEME medium with glutamine. Drugs, including 10 mM glucose, 1 µM oligomycin, and 50 mM 2-DG, from the XF Glycolysis Stress Test Kit were given according to the manufacturer’s instructions. With regard to the mitochondrial stress test, the detection medium was mixed with Seahorse XF DEME medium, glutamine, glucose, and pyruvate. Drugs such as 1.5 µM oligomycin, 0.5 µM FCCP, and 0.5 µM rotenone/antimycin A (Rot/AA) (rot/AA) from the XF Cell Mito Stress Test Kit were used on the basis of the manufacturer’s instructions. In addition to normal ATII and RLE-6TN cells, we also tested RLE-6TN cells treated with Mdivi-1.

Mitochondrial and membrane potential measurements

In 6-well plates, cells were planted at a density of 50,000 cells/well. The mitochondrial membrane potential was measured using an enhanced JC-1 assay kit (C2003S, Beyotime, Shanghai, China) according to manufacturer’s instructions. JC-1 was diluted in JC-1 dye working solution (1:200) and then applied to the plates. Cells were incubated for approximately 20 min at 37 °C and subsequently observed via fluorescence microscopy (ECLIPSE Ts2, Nikon, Tokyo, Japan). Mitochondria in normal cells have a greater membrane potential, and JC-1 can create orange-red fluorescent J-aggregates outside of the mitochondrial membrane, while sensor dyes appear as green fluorescent monomers in cells that had damaged mitochondria [21]. Red fluorescence indicates significant mitochondrial polarization, while JC-1 monomer green fluorescence indicates depolarized areas. MitoTracker Red CMXRos (M9940, Solarbio, Beijing, China) was used to label mitochondria according to manufacturer’s instructions. MitoTracker was diluted using DMEM/F12 medium supplemented with serum to 200 nM and given to the ATII cells, incubated for 30 min at 37 ℃, and subsequently observed using confocal microscopy (LSM880, Zeiss AG, Oberkochen, Germany).

Transmission electron microscopy

Approximately 1 mm3 of fresh lung tissue from the left lung was promptly obtained following thoracotomy for transmission electron microscopy (TEM). 2.5% glutaraldehyde electron microscope fixation solution was used to fix lung tissue at room temperature for 2 h in the dark and subsequently transferred to 4 ℃ for 24 h of preservation. Thereafter, it was dehydrated through a graded series of 50–100% ethanol before embedding. After obtaining ultrathin slices (50–60 nm), they were double-stained for 30 min at room temperature using lead citrate and uranyl acetate. Samples of lung tissue from the model group at 3, 7, 10, and 14 d postnatal were examined using a TEM (JEM1400PLUS, Japan Electron, Japan) at 100 kV. At 10,000× magnification, the quantity and morphological changes of mitochondria were noted in ATII cells that had peculiar lamellar bodies.

Double immunofluorescence staining

For tissue immunofluorescence, after dehydration, phosphate-buffered saline (PBS) was used to wash paraffin sections three times for 5 min each. Microwave repair was performed using citric acid for 7 min, and then cooling to room temperature. The cells were washed thrice with PBS for 5 min. The membrane was blocked with 5% bovine serum albumin (ST025-20g, Beyotime, Shanghai, China) at room temperature for 1 h. After drying, cells were incubated for about 16 h with anti-SPB (1:100, sc-133143, Santa Cruz Biotechnology, Dallas, Texas, USA) and anti-DRP1 (D6C7) rabbit monoclonal antibodies (1:100, 8570, Cell Signalling Technology, Danvers, Massachusetts, USA) at 4 °C, and then followed by incubation with Cy3 goat anti-rabbit IgG (H+L) (1:100, K1209, Ape×Bio, San Diego, California, USA) and Alexa 488 goat anti-mouse IgG (H+L) antibodies (1:100, K1204, Ape×Bio, San Diego, California, USA) for about 2 h at room temperature. Finally, DAPI (AR1176, BOSTER, Wuhan, China) was used to stain nuclei, and used confocal microscopy (LSM880, Zeiss AG, Oberkochen, Germany) to capture fluorescent images.

For cellular immunofluorescence, ATII cells at 7, 10, and 14 d were seeded at 500,000 cells/mL in 6-well plates for detection. Cells were then washed with PBS and fixed with 4% paraformaldehyde for about 20–30 min. The cells were permeabilized and blocked with 0.5% Triton (9002-93-1, Solarbio, Beijing, China) and serum for 10 min at room temperature for 1 h. Thereafter, anti-SPB and anti-DRP1, or anti-Phospho-DRP1 (Ser616) polyclonal (3455, Cell Signalling Technology, Danvers, Massachusetts, USA) and Mff monoclonal antibodies (1:100, 66527-1-Ig, Proteintech, Wuhan, China) were given to the cells and incubated for 16 h at 4 °C, followed by incubation for 2 h with a fluorescent secondary antibody at room temperature. DAPI was used to stain the nuclei, and used confocal microscope to capture fluorescent images.

ATP test kit

ATII (3, 7, 10, and 14 d) or RLE-6TN cells were seeded at 100,000 cells/well in 6-well plates. ATP production was measured by an ATP test kit (S0027, Beyotime, Shanghai, China). The cells were lysed with 200 µL of ATP-detected lysate in each well and centrifuged at 12000g at 4 ℃ 5 min. The supernatant was stored for subsequent determination. The ATP-detected lysate diluted ATP standard solution to the appropriate concentration gradients: 0.01, 0.03, 0.1, 0.3, 1, 3, and 10 µM. The ATP test liquid was configured by diluting the ATP test reagent with the ATP test reagent diluent at a proportion of 1:4. The test was performed in a 96-well plate; each well was added with 100 µL of ATP detection working solution and incubated at room temperature for 5 min. Thereafter, 20 µL of samples or standard products were given to each well and subsequently tested with a multimode reader (Cytation 5, Biotek, Vermont, USA).

Metabolomics detection

The cell precipitates of the model and control groups were collected, frozen in liquid nitrogen, and submitted to the Shanghai iProteome Company for testing (project number: PDZX-2024-0005). For analysis and identification, high-resolution LMS based on the Orbitrap principle was used to detect metabolites in samples, which were qualitatively and quantitatively matched with retention time, molecular mass, secondary spectrum, and other metabolite information in the local database.

Statistical analyses

We used GraphPad Prism version 9.0 (GraphPad Software) for data analysis. All data are presented as the mean ± standard deviation (SD). The Shapiro–Wilk test was used to determine the normality of the data sets. Pearson’s correlation was used to perform correlation analysis. To compare multiple groups, we used a two-way analysis of variance (ANOVA) analysis of variance. With regard to all comparisons, P < 0.05 was considered significant.

Results

Hyperoxia exposure leads to histological changes in the lungs

After neonatal rats were exposed to normoxic or hyperoxic conditions for 3, 7, 10, and 14 d, lung tissues were collected, and alveolar development was evaluated. In the control group, the quantity of alveoli rose, MAD and MLI decreased, alveolar ridge structure increased, and alveolar septum gradually thinned. In the model group, the number of alveoli decreased, MAD and MLI increased, alveolar structure was simplified, and the alveolar septum gradually thickened (Fig. 1A). At 3 d postnatal, alveolar RAC did not reveal any statistically significant changes between the control and model groups. However, 7 d after birth, the MAD and MLI were larger in the model group, and the alveolar ridge structure was blunt and reduced. The values of RAC were significantly low in the model group (P < 0.05). Similarly, the alveolar septum thickness was not significantly different between the groups at 3 d postnatal, whereas at 7, 10, and 14 d after birth, the lung septum thickness significantly increased in the model group (P < 0.05, Fig. 1B–E). These results suggested that lung histological changes and alveolar growth were delayed after hyperoxia exposure.

Fig. 1
figure 1

Hyperoxia exposure leads to histological changes in the lungs. A HE staining of the lung tissues of neonatal rats was observed at 3, 7, 10, and 14 d after birth via microscopy (×400 magnification; scale bar, 50 μm; n = 6). B–E Changes in lung morphology were evaluated using RAC values, MLI, MAD, and alveolar wall thickness. The mean ± SD is used to express the data. * P < 0.05, ** P < 0.01 compared with control group

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Mitochondrial morphological disorders in ATII cells exposed to hyperoxia

Mitochondria of primary ATII cells in control and model groups were examined via TEM, and ultrastructure was compared. In the control group, the basic cell substructure appeared normal, with compact mitochondria and numerous lipid lamellar bodies in ATII cells (white arrows). Furthermore, the mitochondrial cristae gradually became denser, and the mitochondrial volume gradually increased. Three days after birth, under hyperoxic conditions, the mitochondrial structure started to exhibit signs of disruption. Seven days after birth, the mitochondrial size of the model group significantly decreased, and mitochondrial counts significantly increased (P < 0.05). The mitochondrial cristae disappeared, the double-membrane structure of mitochondria was destroyed, and mitochondria swelled, fractured, and became more split. Structural impairment of the lamellar corpuscle was also observed in the model group (yellow arrows; Fig. 2A–C). The staining results of ATII cells revealed a mitochondrial transformation from long and tubular to short and fragmented in the model group (yellow arrows), contrasted to that in the control group (white arrows), particularly at 14 d after birth (Fig. 2D). In conclusion, ATII cells in the model group showed a progressive increase in mitochondrial damage over time, characterized by increasing swelling and fragmentation, which became more pronounced as time elapsed. Hyperoxia exposure also led to membrane potential measurement (MMP) reduction in ATII cells. MMP, a sensitive indicator of mitochondrial damage and autophagy activation, was measured using fluorescence intensity in ATII cells to assess the damage to mitochondrial function in the model group at different times [22]. To examine the changes in mitochondrial membrane potential, ATII cells were subjected to JC-1 staining in both the model and control groups at 3, 7, 10, and 14 d after birth. The findings demonstrated that the membrane potential of the model group significantly dropped from 7 days after birth, in comparison to the control group, with JC-1 polymer (red) gradually decreased, and JC-1 monomer (green) gradually increased (Fig. 2E, F).

Fig. 2
figure 2

Morphological changes of mitochondria in ATII cells after hyperoxia exposure. A Structural changes in mitochondria were observe using transmission electron microscopy, and images show mitochondrial morphological changes in ATII cells (yellow/white arrow; ×30 000 magnification; scale bar, 0.5 μm; n = 3). B, C Changes in mitochondrial size and cell count. D MitoTracker Red CMXRos staining showing that model group’s mitochondria were short and fragmented (yellow/white arrow; ×800 magnification; scale bar, 25 μm). E The membrane potential of the model group evidently decreased 7 d after birth, the JC-1 polymer (red) gradually decreased, and the JC-1 monomer (green) gradually increased (×400 magnification; scale bar, 50 μm; n = 3). F Analyses the ration of JC-1 polymer (red) to JC-1 monomer (green) in primary ATII cells at 3, 7, 10, and 14 d after birth in the control and model groups. The mean ± SD is used to express the data. * P < 0.05, ** P < 0.01 compared with the control group

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Increased expression of mitochondrial dynamics-associated proteins in hyperoxia-exposed lung tissues and ATII cells

The mitochondrial outer membrane fusion proteins MFN1 and MFN2 were expressed at lower levels in the lung tissue of the model group at 7 and 14 d after birth, respectively, compared to the control group, with a significant decrease at 7 d after birth (P < 0.05). The expression of the inner membrane fusion protein OPA1 was reduced at 3 and 7 d after birth, followed by an elevation at 10 and 14 d after birth, particularly noticeable at 10 d after birth, similar to the pattern observed for MFN1 and MFN2 at 10 d after birth, suggesting a potential compensatory elevation. The fission proteins DRP1 and p-DRP1 in the model group increased 3 d after birth and were significantly elevated 7 d after birth (P < 0.05) in lung tissue (Fig. 3A–F). In primary ATII cells, the expression levels of MFN1, MFN2, and OPA1 in the model group significantly declined at 3 and 7 d after birth (P < 0.05). However, at 10 and 14 d after birth, the reduction in MFN1 and MFN2 was not evident, and the OPA1 expression marginally increased. The fission protein DRP1 in the model group significantly rose at 3 d after birth (P < 0.05), whereas p-DRP1 significantly increased 7 d after birth (Fig. 4A–F). According to these findings, the model group had a higher frequency of mitochondrial fission compared to the control group. SPB is a biomarker for the identification of ATII cells [23]. Double staining for DRP1 and SPB was performed on lung tissue sections and ATII cells to evaluate DRP1 expression. These two proteins are primarily distributed in the cytoplasm. DRP1 and SPB co-localized, and the fluorescence of DRP1 gradually increased over time in the model group contrasted to that in the control group (white arrows, Figs. 3G and 4G). Expression level of p-DRP1 and the mitochondrial outer membrane receptor Mff, which can recruit activated DRP1 to the mitochondria and promote mitochondrial division, was also observed [24]. The fluorescence intensities of p-DRP1 and Mff gradually increased over time (white arrows, Fig. 4H).

Fig. 3
figure 3

Changes in the expression of mitochondrial dynamics-associated proteins in lung tissues of neonatal rats with bronchopulmonary dysplasia. A–F Western blotting and analyses of MFN1, MFN2, OPA1, DRP1, and p-DRP1 at 3, 7, 10, and 14 d after birth in the lung tissues of the control and model groups (n = 3). G Representative immunostaining of lung sections from the control and model groups using anti‑DRP1 and anti‑SPB. Yellow puncta denote co-localization (white arrows, ×400 magnification; scale bar, 50 μm; n = 3). The mean ± SD is used to express the data. *P < 0.05, **P < 0.01 vs. control

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Fig. 4
figure 4

Changes in the expression of mitochondrial dynamics-associated proteins in primary ATII cells of rats with bronchopulmonary dysplasia. A–F Western blotting and analyses of MFN1, MFN2, OPA1, DRP1, and p-DRP1 in primary ATII cells at 3, 7, 10, and 14 d after birth in the control and model groups (n = 3). G, H Representative immunostaining of ATII cell slides from the control and model groups using anti‑DRP1 and anti‑SPB or anti‑p-DRP1 and anti-Mff antibodies. Yellow puncta denote co-localization (white arrows, ×400 magnification; scale bar, 50 μm; n = 3). The mean ± SD is used to express the data. *P < 0.05, **P < 0.01 vs. control

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Increased expression of glucose metabolic reprogramming-associated proteins in hyperoxia-exposed lung tissue and ATII cells

In the model group of lung tissue, the expression levels of glycolysis-associated enzymes, including PFKM, hexokinase 2 (HK2), and LDHA, three key rate-limiting enzymes in glycolysis, increased compared to those in the control group 7 d after birth (P < 0.05). However, 3 d after birth, the PFKM and LDHA expression decreased, suggesting a potential compensatory reduction (Fig. 5A–D). In primary ATII cells, the expression of PFKM, HK2, and LDHA increased under hyperoxic conditions 3 d after birth; however, their expression was significantly elevated 7 d after birth (P < 0.05). These results suggested that under hyperoxic conditions, glucose tends to undergo glycolysis rather than OXPHOS in the model group (Fig. 5E–H).

Fig. 5
figure 5

Changes in glucose metabolism in lung tissues and ATII cells. A–D Western blotting and analyses of HK2, PFKM, and LDHA in the lung tissues of the control and model groups at 3, 7, 10, and 14 d after birth (n = 3). E–H Western blotting and analyses of HK2, PFKM, and LDHA in primary ATII cells in the control and model groups at 3, 7, 10, and 14 d after birth (n = 3). I, J Representative graph of ECAR output of ATII cells and responses to glucose, 2‑DG and oligomycin, and ECAR of glycolysis performed using Seahorse XF96. K, L Representative graph of the OCR output of ATII cells and responses to FCCP, oligomycin, rotenone/antimycin A, and mitochondrial OCR of maximal respiration performed using Seahorse XF96. The assay was conducted on one plate with at least three replicates. M ATP concentrations in primary ATII cells at 3, 7, 10, and 14 d after birth in the control and model groups (n = 3). The mean ± SD is used to express the data. *P < 0.05, **P < 0.01 vs. control

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Hyperoxia leads to increased glycolysis and decreased OXPHOS in ATII cells

Real-time analyses of the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) were performed using ATII cells isolated from lung tissues via the Seahorse XF96 metabolic extracellular flux analyzer. The model group’s anaerobic metabolism levels were assessed throughout time and compared to those of the control group, and the ECAR of glycolysis increased 3 d after birth. However, it significantly increased at 10 and 14 d after birth (P < 0.05) (Fig. 5I, J). Furthermore, the mitochondrial OCR of maximal respiration was elevated from 3 d after birth and significantly increased 7 d after birth in the model group in contrast to that in the control group (P < 0.05) (Fig. 5K, L). This finding suggested that mitochondrial dysfunction may occur earlier than aerobic glycolysis. ATP concentrations in ATII cells separated from the BPD model were significantly reduced 3 d after birth. ATP was primarily produced by mitochondrial OXPHOS in cells, with low levels of ATP indicating a decrease in OXPHOS (P < 0.05) (Fig. 5M). After the administration of 2-DG (50 mg/kg.d for 10 d since 5 d after birth), the expression level of HK2, PFKM, LDHA in ATII cells was lower than that in the model group (P < 0.05, Fig. 6A, B).The histological changes in the lungs at 14 d after birth showed that the alveolar structure was improved in the group of given 2-DG contrast to the model group. The thickened alveolar septum and reduced RAC value in model group were alleviated (P < 0.05, Fig. 6C–E).

Fig. 6
figure 6

Histological changes of lungs after given 2-DG to model group. A, B Western blotting and analyses of HK2, PFKM, and LDHA in the ATII cells of the control, model, and 2-DG given groups (n = 3). C HE staining of the lung tissues of neonatal rats was observed at 14 d after birth via microscopy (×400 magnification; scale bar, 50 μm; n = 3). D, E Changes in lung morphology were evaluated using RAC values, and alveolar wall thickness. The mean ± SD is used to express the data. *P < 0.05, **P < 0.01 vs. control

Full size image

Expression of DRP1 and glucose metabolic reprogramming proteins in hyperoxia-exposed RLE-6TN cells

In the model group of RLE-6TN cells, the expression levels of MFN1, MFN2, and OPA1 was noted decreasing, alongside an elevation in the expression level of DRP1, particularly noticeable at 48 h after exposure to hyperoxia (P < 0.05) (Fig. 7A–E). The expression levels of glycolysis-associated enzymes, including PFKM, HK2, and LDHA, were higher than the control group, particularly at 48 h after exposure to hyperoxia (P < 0.05). However, 72 h after hyperoxia exposure, the expression level of LDHA decreased, which may be associated with the cell state (Fig. 7F–I).

Fig. 7
figure 7

Changes of mitochondrial dynamics and glucose metabolic reprogramming in RLE-6TN cells exposed to hyperoxia. A–E Western blotting and analyses of MFN1, MFN2, OPA1, and DRP1 in RLE-6TN cells at 24, 48, and 72 h after hyperoxia exposure (n = 3). F–I Western blotting and analyses of HK2, PFKM, and LDHA in RLE-6TN cells at 24, 48, and 72 h after exposed to hyperoxia (n = 3). J Volcanic map of differential metabolites. Red, upregulated differential metabolites; blue, downregulated differential metabolites; and gray, undifferentiated metabolites. K, L Circos and histograms of differential metabolites, KEGG pathway enrichment *P < 0.05, **P < 0.01 vs. control

Full size image

Metabolite changes in RLE-6TN cells under hyperoxia conditions

Non-target metabolomic detection of RLE-6TN cells was completed in both the model and control groups. The volcanic map of differential metabolites showed that 46 metabolites were upregulated and 139 were downregulated, including sodium gluconate, uridine diphosphate glucose, glucaron, and other glucose metabolites (P < 0.05) (Fig. 7J). The different metabolites were enriched in the KEGG pathway, and the circos and histogram exhibited changes in sugar metabolism-related pathways, such as sucrose and galactose (P < 0.05) (Fig. 7K, L). The Seahorse XF96 metabolic extracellular flux analyzer was also used to perform real-time analyses of the ECAR and OCR values of RLE-6TN cells under hyperoxia conditions. The ECAR of glycolysis increased after 24 h of hyperoxia exposure and was particularly elevated at 48 h (P < 0.05). The mitochondrial OCR of maximal respiration was significantly elevated at 48 h in the model group compared to that in the control group (P < 0.05) (Fig. 8A–D).

Fig. 8
figure 8

Effect of Mdivi-1 administration on RLE-6TN cells exposed to hyperoxia. A–D Representative graphs of the ECAR and OCR outputs of RLE-6TN cells at 24, 48, and 72 h after hyperoxia exposure. E Cell viability curve of RLE-6TN cells after the treatment with Mdivi-1. F, G Western blotting and analyses of DRP1, PFKM, HK2, and LDHA in RLE-6TN cells at 48 h after exposed to hyperoxia and the addition of Mdivi-1 (n = 3). H, I Representative graph of ECAR and OCR output of RLE-6TN cells at 48 h after exposed to hyperoxia and the addition of Mdivi-1. J ATP concentration at 48 h after hyperoxia exposure and addition of Mdivi-1. K MitoTracker Red CMXRos staining showed that mitochondrial fragmentation improved after the addition of Mdivi-1 48 h under hyperoxic conditions (white arrows, ×800 magnification; scale bar, 25 μm). The mean ± SD is used to express the data. *P < 0.05, **P < 0.01 vs. control

Full size image

Concentration selection of Mdivi via the CCK8 assay

As the most significant changes in RLE-6TN cells were observed at 48 h after hyperoxia exposure, 48 h was the time selected for intervention administration to confirm the regulatory relationship between mitochondrial damage and metabolic reprogramming at the cellular level. Under normoxic conditions, RLE-6TN cells were treated with Mdivi-1 to inhibit mitochondrial division. The CCK8 assay was used to determine the optimal concentration of Mdivi-1. By making the cell viability curve showed that the optimal concentration of Mdivi-1 was 5 μm, with no significant difference in cell viability compared to the control (Fig. 8E).

Inhibition of mitochondrial fission downregulates the expression of glucose metabolic reprogramming-associated proteins

After the administration of Mdivi-1, the expression level of DRP1 in RLE-6TN cells 48 h after hyperoxia exposure was lower than that in the control group (P < 0.05). The expression levels of glycolysis-associated enzymes, including PFKM, HK2, and LDHA, at 48 h were significantly downregulated compared to the control group (P < 0.05) (Fig. 8F, G). Therefore, we also selected 48 h to confirm metabolic changes. Following Mdivi-1 administration, the observed increase in ECAR and decrease in OCR were alleviated (Fig. 8H, I). ATP levels were elevated, indicating an increase in OXPHOS (P < 0.05) (Fig. 8J). MitoTracker Red CMXRos staining showed that the addition of Mdivi-1 improved mitochondrial fragmentation in the model group (white arrows, Fig. 8K).

Discussion

Long-term oxygen inhalation is harmful to neonates, especially preterm infants, and increases the risk of oxygen toxicity. BPD is the most prevalent complication of hyperoxia [25]. However, the pathogenesis of BPD remains unexplored [26]. Recently, changes in the metabolic patterns of BPD have been observed; however, the specific molecular mechanisms are not well established [8, 27, 28]. Mitochondrial dysfunction can affect the onset and development of BPD via morphological abnormalities, mitochondrial oxidative stress, mitochondrial biogenesis, and mitochondrial dynamics [29]. Mitochondrial dysfunction can also affect cell metabolism [30]. In our study, we examined the alterations in mitochondrial dynamics and discussed their role in regulating glucose metabolism under hyperoxic conditions to explore the mechanism of ATII cell injury in an animal model of BPD and a hyperoxia cell model.

Under normal physiological conditions, the division and fusion of mitochondria are maintained in a relatively balanced state. When division is excessive or fusion is reduced, mitochondrial fragmentation increases, resulting in abnormal morphology, quantity, and function. Mitochondrial fragmentation is the origin of several diseases, including Parkinson’s disease, cardiovascular disease, and diabetes; however, its role in BPD remains unclear. The mitochondrial division proceeds sequentially via DRP1 activation, fission site labeling, ring spirochaete assembly, GTP hydrolysis, and DRP1 spiral contraction, finally severing mitochondria. DRP1, localized in the cytosol, is crucial for regulating mitochondrial fission by interacting with proteins on the mitochondrial outer membrane, including fission 1 (Fis1), Mff, and mitochondrial dynamic proteins of 49 and 51 KDa (MiD49 and MiD51), leading to mitochondrial fission. Among them, Mff can accumulate directly at mechanically induced contractions on the mitochondria, which plays a core role in DRP1 recruitment in mitochondrial fission in mammalian cells. Fis1 acts as an adapter for the yeast-specific peripheral membrane proteins, which are then recruited and assembled with the yeast DRP1 homologous Dnm1, is not appear to be necessary for mitochondrial division in mammalian cells [31,32,33]. Similar to the studies of Dai et al. and Ma et al., we observed changes in mitochondrial dynamics in the animal models of BPD and hyperoxia cell models [15, 34]. In our study, DRP1 and p-DRP1 expression was elevated under hyperoxic conditions in lung tissue and primary ATII cells. DRP1 has many post-translational modifications, including phosphorylation, SUMOylation, acetylation, and O-GlcNAcylation. Phosphorylation of DRP1 is the most prominent and extensively studied [35]. When fission is initiated, DRP1 is recruited to the outer mitochondrial membrane and phosphorylated at serine 616, causing mitochondrial fragmentation [36]. Mff recruits activated DRP1 to transposition to the mitochondrial outer membrane [37]. Our results from immunofluorescence double staining demonstrated a significant increase in the binding of p-DRP1 to Mff after activation, driving mitochondrial division.

Western blotting and Seahorse XF96 metabolic extracellular flux analysis revealed that the change in DRP1 expression preceded the increase in glycolysis under hyperoxic conditions. This was confirmed by administering Mdivi-1 to RLE-6TN cells exposed to hyperoxic conditions. The addition of Mdivi-1 reduced ECAR and OCR values, increased ATP production, and alleviated mitochondrial fragmentation, consistent with the findings of Ding et al. [38]. Metabolic reprogramming is complex, involving dysregulation of glucose, amino acid, and lipid metabolism, which can occur simultaneously and interact with each other [39]. The increase in glycolysis and decrease in the TCA cycle during glucose metabolic reprogramming leads to a reduce in ATP production. The enhancement of mitochondrial fission also inhibits ATP production, which may be a potential mechanism. Our results also showed that hyperoxia inhibited ATP production in ATII cells in the BPD model. Reprogramming of energy metabolism may be an important mechanism in BPD. In this study, we found that the elevated expression of HK2 preceded that of PFKM and LDHA in lung tissue, indicating that the activation of the HK2 might play a key modulator in glucose metabolic reprogramming. HK2 can also bind to voltage-dependent anion channels in the mitochondrial outer membrane and inhibit the opening of the mitochondrial permeability transition pore [40]. Although the role of the DRP1-HK2 signaling pathway has been reported in other diseases, its role in BPD remains unclear [41]. In addition to changes in glucose metabolism-related enzymes, glycometabolism-related products were altered by metabolomic detection.

Our study had some limitations. First, Mdivi-1 was not administered in vivo. Second, this study primarily discussed the relationship between DRP1 and glucose metabolic reprogramming under hyperoxic stimulation and also detected p-DRP1 expression, but we did not explore the specific mechanism between glucose metabolic reprogramming and DRP1. Future studies will focus on the effects of hyperoxia and DRP1, exploring other post-translational modifications and their influence.

Conclusion

In this study, we demonstrated that the exposure of lung epithelial cells to hyperoxia may influence mitochondrial fission, consequently impacting glucose metabolism and leading to glucose metabolic reprogramming via activating the DRP1 signaling pathway. Subsequent deciphering of the molecular mechanisms underlying hyperoxia-associated lung injury could aid in the development of future strategies to target and protect ATII cells from injury in BPD models.

Data availability

No datasets were generated or analysed during the current study.

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Acknowledgements

We are truly appreciative Key Laboratory of Health Ministry for Congenital Malformation for their technical support in finishing this research.

Funding

This research was supported by grants from the Key R&D Guidance Plan Projects in Liaoning Province (2020JH1/10300001), National Natural Science Foundation of China (No. 82071688, No. 82471752).

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  1. Danni Li and Jianhua Fu contributed equally.

  2. Tong Sun and Haiyang Yu contributed equally.

Authors and Affiliations

  1. Department of Pediatrics, Shengjing Hospital of China Medical University, Shenyang, Liaoning, China

    Tong Sun, Dingning Zhang, Danni Li & Jianhua Fu

  2. Department of Neurology, Shengjing Hospital of China Medical University, Shenyang, Liaoning, China

    Haiyang Yu

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Contributions

Conceptualization: JH Fu. Data Curation: T Sun, DN Li, DN Zhang. Formal Analysis: T Sun, HY Yu. Funding acquisition: JH Fu. Supervision: JH Fu. Writing original Draft: T Sun, HY Yu. Writing: review and editing: JH Fu and DN Li.

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Sun, T., Yu, H., Zhang, D. et al. Activated DRP1 promotes mitochondrial fission and induces glycolysis in ATII cells under hyperoxia. Respir Res 25, 443 (2024). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12931-024-03083-8

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Respiratory Research

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