MSC-Mediated Mitochondrial Transfer Promotes Metabolic Reprogramming in Endothelial Cells and Vascular Regeneration in ARDS

doi:10.21203/rs.3.rs-4813289/v1

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MSC-Mediated Mitochondrial Transfer Promotes Metabolic Reprogramming in Endothelial Cells and Vascular Regeneration in ARDS

https://doi.org/10.21203/rs.3.rs-4813289/v1

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Background

Acute Respiratory Distress Syndrome (ARDS) involves extensive pulmonary vascular endothelial injury. Mitochondrial damage plays a critical role in this endothelial injury. While mesenchymal stem cells (MSCs) are being explored as a cellular therapy for ARDS, their role in repairing mitochondrial damage in endothelial cells remains unclear. This study investigates the potential of MSCs to repair mitochondrial damage in ARDS lung endothelial cells through mitochondrial transfer and elucidates the underlying mechanisms.

Methods

This study established ARDS mouse models and cellular models of mitochondrial damage in pulmonary endothelial cells. Initially, we observed the ability and mechanisms of MSCs to transfer mitochondria to lung endothelial cells both in vivo and in vitro. Subsequently, we investigated how this mitochondrial transfer by MSCs affects the repair of mitochondrial and endothelial damage, as well as its impact on vascular regeneration in ARDS. Finally, we elucidated the mechanisms by which MSC-mediated mitochondrial transfer promotes vascular regeneration in ARDS. Various cell biology techniques, including flow cytometry, immunofluorescence staining, and confocal microscopy, were utilized for experimental observations.

Results

MSCs used tunneling nanotubes (TNTs) to transfer mitochondria to pulmonary endothelial cells. The endothelial cells internalized these mitochondria through dynamin-dependent clathrin-mediated endocytosis. The mitochondrial transfer increased mitochondrial complex I expression, reduced ROS production and apoptosis, and promoted cell proliferation in endothelial cells. The reparative effects of MSCs diminished when their mitochondrial transfer ability was inhibited. MSC-mediated mitochondrial transfer activated the tricarboxylic acid (TCA) cycle and citrate-dependent fatty acid synthesis in endothelial cells, leading to the release of pro-angiogenic factors and promoting vascular regeneration. Inhibiting TCA or fatty acid synthesis in endothelial cells significantly reduced MSC-promoted vascular regeneration.

Conclusion

MSCs transfer mitochondria to ARDS lung endothelial cells, activating the TCA cycle and fatty acid synthesis, which promotes endothelial cell proliferation and the release of pro-angiogenic factors, thereby enhancing vascular regeneration. These findings offer a promising therapeutic approach for repairing mitochondrial damage and promoting vascular regeneration in ARDS.

Acute Respiratory Distress Syndrome

Mesenchymal Stem Cells

Mitochondrial Transfer

Pulmonary Microvascular Endothelial Cells

Tunneling Nanotubes

Acute Respiratory Distress Syndrome (ARDS) represents a significant challenge in the field of critical care medicine, characterized by a high incidence and mortality rate, imposing a substantial burden on patients and healthcare systems worldwide. According to global epidemiological data, the incidence of ARDS is showing an increasing trend year by year, with millions of new cases emerging globally each year, and the mortality rate often exceeds 30%[1, 2]. Epidemiological data from China indicates that the hospital mortality rate of ARDS is 34%, and even higher for severe ARDS, reaching up to 60%[3]. Over the past two decades, clinical treatment progress for ARDS has remained at the level of organ support, such as lung-protective ventilation strategies and prone positioning ventilation, lacking effective treatment targeting the pathogenesis and pathological changes of ARDS[4]. Therefore, finding therapeutic methods that can block the pathogenesis of ARDS and alleviate its pulmonary pathological damage has significant clinical application value in improving the prognosis of patients with ARDS.

Mesenchymal Stem Cells (MSCs), characterized by their pluripotency, self-renewal, and immunomodulatory capabilities, have emerged as a promising therapeutic strategy in the treatment of ARDS. Firstly, Recent studies have highlighted the role of MSCs in modulating both innate and adaptive immune responses in ARDS, leading to a reduction in lung injury attributable to excessive inflammation[5, 6]. Secondly, These cells have been shown to enhance pathogen clearance and mitigate both direct and inflammatory damages[7]. Thirdly, MSCs facilitate the repair of alveolar epithelial and pulmonary endothelial cells, thereby restoring the integrity of the alveolar-capillary barrier and improving pulmonary fluid clearance[8]. Lastly, their potential to differentiate into damaged lung tissues positions MSCs as a pivotal agent in promoting lung repair in ARDS[9]. Our preliminary investigations have demonstrated that MSCs can significantly reduce inflammatory lung damage in ARDS mouse models, decrease pulmonary capillary permeability, and ameliorate lung pathology. However, the mechanisms by which MSCs repair ARDS lung injury have not yet been fully elucidated.

Mitochondrial damage plays a crucial role in the pathogenesis and progression of ARDS, being intimately associated with aspects such as energy metabolism, oxidative stress, cell apoptosis, and oxygen sensing[10, 11]. Mitochondria are vital organelles within cells, primarily responsible for energy production, regulating cellular metabolism, and maintaining intracellular homeostasis. In the pathophysiology of ARDS, mitochondrial damage in alveolar epithelial cells and microvascular endothelial cells is directly involved in the disease process[12]. Specifically, mitochondrial damage can lead to disturbances in cellular energy metabolism, weakening the cell's adaptability to stress and exacerbating the progression of ARDS[13]. Changes in mitochondrial membrane permeability trigger the release of mitochondrial DNA, provoking an intensified inflammatory response by the immune system[14]. Alterations in the protein channels of the mitochondrial membrane directly result in apoptosis and necrosis of endothelial and epithelial cells. Moreover, mitochondria are involved in oxygen sensing and cell signaling, mitochondrial damage can lead to dysregulation of cellular signaling pathways[15].

Mitochondrial transfer refers to the process where mitochondria are transported from one cell to another. This phenomenon was first revealed in a study published in 2006, which demonstrated that ρ0 cells lacking mitochondrial DNA could restore mitochondrial function by accepting mitochondria transferred from adjacent cells[16]. Recent studies have shown that mitochondrial transfer can occur between cells through various mechanisms, including transfer via extracellular vesicles, establishment of tunneling nanotubes (TNTs) for mitochondrial transfer, and release of wandering mitochondria by donor cells for capture by recipient cells[17, 18]. Upon receiving healthy mitochondria, the recipient cells experience a restoration of impaired mitochondrial function, stabilizing their energy metabolism and maintaining the functionality of various organs under disease conditions[17].

MSCs can repair mitochondrial damage in ARDS through mitochondrial transfer. Since Islam et al. first discovered that MSCs could transfer mitochondria to ARDS alveolar epithelial cells, restoring their aerobic metabolism[19], the mechanism of mitochondrial transfer has gained increasing attention in the field of ARDS injury repair. Recent studies have shown that MSCs can transfer mitochondria to alveolar macrophages in ARDS, enhancing their phagocytic ability and increasing the efficiency of pathogen clearance[20, 21]. Additionally, MSCs can transfer mitochondria to T cells, increasing the proportion of regulatory T cells and exerting anti-inflammatory effects[22, 23]. This study explored whether MSCs transfer mitochondria to ARDS pulmonary endothelial cells and elucidated the potential role and mechanism of mitochondrial transfer in promoting the regeneration of ARDS pulmonary vasculature. Our results confirmed that MSCs could transfer mitochondria to ARDS pulmonary endothelial cells, stimulating the Tricarboxylic Acid (TCA) cycle in endothelial cells, activating citrate-dependent fatty acid synthesis, and promoting vascular regeneration in ARDS. This finding provides new insights for the repair of pulmonary injury in ARDS.

Establishment of Animal and Cell Models

Husbandry of C57BL/6J Mice

In this study, male C57BL/6J mice (Strain number N000013), aged 5–8 weeks, were procured from GemPharmatech (Nanjing, China). Upon arrival, the mice were housed in a specific pathogen-free (SPF) environment, where the ambient temperature was maintained at 22 ± 2°C with a relative humidity of 50 ± 10%, adhering to a 12-hour light/dark cycle. The mice were kept in standard ventilated cages, with 3–5 mice per cage. Access to food and water was provided ad libitum. Throughout the experimental period, the welfare and health status of the animals were routinely monitored by a professional team. The Animal Experimental Ethics Committee of Southeast University approved these experiments (Approval number 20200226001), which were conducted in compliance with Chinese legislation concerning the use of experimental animals.

Establishment of the ARDS Mouse Model

An ARDS mouse model was established using a chemical induction method. Male C57BL/6J mice, aged 5–8 weeks and weighing between 20–25 grams, were used for the experiments. The mice were acclimatized for at least one week prior to the experiment. For the modeling procedure, the mice were first anesthetized with 2%-3% isoflurane inhalation. Subsequently, tracheal intubation was performed, followed by the intratracheal injection of 5 mg/kg body weight of LPS (Beyotime, cat. no. ST1470) to establish the ARDS model. After the establishment of the model, the mice were returned to clean cages for recovery. Over the next 24 hours, the development of ARDS was assessed by monitoring parameters such as respiratory rate, changes in body weight, and behavioral activity.

Intravenous Injection of MSCs in Mice

MSCs were administered to the experimental mice via tail vein injection. Initially, C57BL/6J mouse bone marrow-derived MSCs (OriCell, cat. no. MUBMX-01001) were purchased. The experimental mice were fasted a day before the MSCs injection, while ensuring an adequate supply of water. On the day of injection, after establishing the ARDS mouse model and 4 hours post-modeling, the mice were secured on a specialized tail vein injection stand. MSCs were diluted in sterile saline to a concentration of 1×10^6 cells/200 µL and then slowly injected into the mouse tail vein. Care was taken to avoid the formation of air bubbles during the injection. After the injection, the mice were returned to their cages for recovery, with close observation to ensure no apparent discomfort or abnormal behavior.

Culture, Passaging, and Cryopreservation of MSCs

Bone marrow-derived MSCs from C57BL/6J mice were expanded in vitro. The culture medium consisted of DMEM/F-12 (Gibco, cat. no. 11320033), 10% fetal bovine serum (Gibco, cat. no. 10099141C), and 1% penicillin-streptomycin (Gibco, cat. no. 15140148). Cells were cultured in a humidified incubator at 37°C with 5% CO₂. When cells reached 80–90% confluency, passaging was performed.

For passaging, cells were first washed twice with PBS (Procell, cat. no. PB180327) and then treated with 1 mL of 0.25% trypsin-EDTA solution (Gibco, cat. no. 25200114) in a 25T flask. Cells were incubated at 37°C for 1–2 minutes until detachment from the flask bottom. Immediately, an equal volume of complete culture medium was added to neutralize trypsin activity. The cell suspension was transferred to a 15 mL centrifuge tube and centrifuged at 300×g for 5 minutes. The supernatant was discarded, and the cells were resuspended in fresh culture medium and seeded into new flasks for continued cultivation.

When MSCs reached the appropriate passage number for cryopreservation, they were first collected following the passaging steps. Cells were resuspended at a concentration of 1×10^6 cells/mL in cryopreservation solution (NCM Biotech, cat. no. C40100). The cell suspension was aliquoted into pre-chilled cryovials, which were then placed in a cryogenic box in a -80°C freezer overnight before transferring to liquid nitrogen for long-term storage.

Culture, Passaging, and Cryopreservation of MPMECs

We cultured an immortalized mouse pulmonary microvascular endothelial cell line (MPMECs), whose proliferative capacity, morphological characteristics, genetic stability, and expression of endothelial cell markers have been confirmed in previous studies[24]. The culture medium comprised of basic DMEM/F-12, 5% fetal bovine serum, 1% endothelial cell growth supplement (ScienCell, cat. no. 1052), 1% penicillin-streptomycin, 90U/mL heparin (Sigma-Aldrich, cat. no. H3149), and 92mg/L D-valine (Sigma-Aldrich, cat. no. V1255). Cells were maintained in a humidified incubator at 37°C with 5% CO2, with fresh medium replacement every 1–2 days. Passaging was performed when cells reached 70%-80% confluence.

For passaging MPMECs, cells were first washed twice with PBS and then treated with 1mL of 0.25% trypsin-EDTA solution diluted tenfold in PBS. Cells were incubated at 37°C for 1–2 minutes until detachment from the flask bottom. Subsequently, an equal volume of culture medium containing FBS was added to neutralize trypsin. The cell suspension was transferred to a centrifuge tube and centrifuged at 300×g for 5 minutes. The supernatant was discarded, and the cells were resuspended in fresh culture medium and seeded into new flasks for continued cultivation.

For cryopreservation of MPMECs, cells were first collected following the passaging protocol. Cells were resuspended at a concentration of 1×10^6 cells/mL in cryopreservation solution. The cell suspension was aliquoted into pre-chilled cryovials. The vials were then placed in a cryogenic box in a -80°C freezer overnight before transferring to liquid nitrogen for long-term storage.

Mitochondrial Damage Induction with Rotenone

In this study, rotenone (MedChemExpress, cat. no. HY-B1756) was employed to induce mitochondrial damage, simulating conditions of mitochondrial dysfunction in ARDS. Rotenone is a naturally occurring compound known to impair mitochondrial respiration by Complex I of the mitochondrial electron transport chain. Cells intended for the experiment were cultured to an appropriate density at 37°C and 5% CO2 prior to treatment. A working solution of rotenone was prepared. The rotenone powder was first dissolved in dimethyl sulfoxide (DMSO) (MedChemExpress, cat. no. HY-Y0320) to create a high-concentration stock solution (1 mM). For experimental use, this stock solution was further diluted to the desired final concentration (100nM) in the culture medium. During treatment, the culture medium containing rotenone was directly added to the cell culture dishes, gently swirled to ensure uniform mixing. The cells were stimulated for 4 hours. After the treatment, the medium containing rotenone was removed, and the cells were washed with PBS, followed by replacement with fresh culture medium for subsequent experiments.

Method Details

Flow Cytometry Analysis of Mitochondrial Transfer from MSCs to MPMECs

MSCs and MPMECs were cultured separately until reaching 70–80% confluency, and mitochondria in MSCs were labeled with MitoTracker Deep Red FM (Invitrogen, cat. no. M22426). Briefly, MitoTracker Deep Red FM was dissolved in DMSO to make a 1 mM stock solution and then diluted with complete culture medium to a working concentration of 200 nM (2 µL of stock solution added to 10 mL of complete culture medium). The culture medium was removed from the flasks, replaced with pre-warmed working solution, and the cells were incubated for 30 minutes. Afterward, the cells were washed with PBS and fresh culture medium was added.

For co-culture experiments, 1×10^5 mitochondria-prelabeled MSCs were co-cultured with MPMECs for 24 hours. Post co-culture, the cells were treated with 0.25% trypsin-EDTA for digestion, then collected by centrifugation and washed with PBS. Prior to flow cytometry, CD31 antibody (BD Pharmingen, cat. no. 553373) was used to specifically label surface proteins of MPMECs. Dual labeling with MitoTracker Deep Red FM and CD31 antibody allowed for the distinction between the two cell types in flow cytometry and the detection of MSCs' mitochondria in MPMECs.

Flow Cytometry Analysis of Mitochondrial Transfer from MSCs to MPMECs in ARDS Mouse Model

An ARDS mouse model was established, and MSCs were cultured to 70–80% confluence and labeled with MitoTracker Deep Red FM for mitochondrial tracking. Four hours after establishing the ARDS mouse model, these mitochondria-prelabeled MSCs were administered via tail vein injection to the ARDS model mice. Twenty-four hours later, the mice were euthanized, and their lung tissues were rapidly harvested. Under sterile conditions, the lung tissues were subjected to mechanical and enzymatic digestion to prepare a single-cell suspension, which was then collected by centrifugation and washed with PBS.

MPMECs were specifically labeled using a fluorescently-tagged CD31 antibody, which targets unique surface proteins of these cells. Flow cytometry was employed to distinguish between endothelial and non-endothelial cells, and to detect the presence of MSC mitochondria within MPMECs. The fluorescence intensity of MitoTracker Deep Red FM within the MPMEC population and the proportion of cells were analyzed to assess the degree of mitochondrial transfer from MSCs to MPMECs.

Immunofluorescence Staining of Mouse Lung Tissue to Detect Mitochondrial Transfer from MSCs to MPMECs in ARDS

Mitochondria of in vitro cultured MSCs were labeled with MitoTracker dye (MitoTracker Deep Red FM). Four hours post-establishment of the ARDS mouse model, these pre-labeled mitochondria MSCs were injected into the ARDS model mice via tail vein. Twenty-four hours after ARDS modeling, the mice were euthanized, and their lung tissues were rapidly extracted and immediately frozen at -80°C. Frozen sections were then prepared, slicing the lung tissues into 5 µm thick sections.

The lung tissue sections underwent immunofluorescence staining, with primary antibody incubation using a CD31 antibody (Abcam, cat. no. ab182981) for labeling MPMEC surface markers. Excess primary antibody was washed off, followed by incubation with a secondary antibody, Alexa Fluor 488 (Abcam, cat. no. ab150077), for specific labeling of MPMECs. Finally, the sections were observed and imaged using a fluorescence microscope. The co-localization of MSCs' MitoTracker labeling (red) with MPMECs' CD31 labeling (green) was analyzed to determine whether MSC mitochondria were transferred to MPMECs.

Confocal Microscopy Observation of Mitochondrial Transfer and Tunneling Nanotubes Between MSCs and MPMECs

Initially, MSC mitochondria were labeled using MitoTracker Deep Red FM. MPMEC mitochondria were labeled with MitoTracker Green FM (Invitrogen, cat. no. M7514). Briefly, MPMECs were cultured in confocal dishes to 50% confluency. MitoTracker Green FM was dissolved in DMSO to make a 1 mM stock solution and then diluted with complete culture medium to a working concentration of 100 nM (1 µL of stock solution added to 10 mL of complete culture medium). The culture medium was removed from the confocal dishes, replaced with pre-warmed working solution, and the cells were incubated for 30 minutes. Subsequently, the cells were washed with PBS and fresh culture medium was added. Mitochondria-labeled MSCs were then added to the confocal dishes containing MPMECs and co-cultured for 12 hours. During this period, MSCs and MPMECs might exchange mitochondria via tunneling nanotubes. Under the confocal microscope, the co-localization of MSC mitochondria (red fluorescence) and MPMEC mitochondria (green fluorescence) was observed, along with the morphology of tunneling nanotubes.

Fluorescence Microscopy Observation of Mitochondrial Transfer from MSCs to MPMECs in Isolated Culture

In this study, we employed a Transwell system to isolate and culture MSCs and MPMECs, and observed the mitochondrial transfer from MSCs to MPMECs using fluorescence microscopy. Initially, MPMEC mitochondria were labeled with MitoTracker Deep Red FM, and MPMECs were cultured in the lower chamber of the Transwell system. MSC mitochondria were labeled with MitoTracker Green FM, and these pre-labeled MSCs were seeded in the upper chamber of the Transwell system. After 24 hours of isolated co-culture, allowing for the transfer of mitochondria from MSCs to MPMECs, fluorescence microscopy was performed. Appropriate fluorescence excitation and emission wavelengths were adjusted to observe the MPMECs in the lower layer of the Transwell system. The co-localization of red and green fluorescence within MPMECs confirmed the transfer of mitochondria from MSCs to MPMECs during isolated culture.

Preparation of Single-Cell Suspension from Mouse Lung Tissue

Initially, mice were euthanized following full anesthesia, and the lungs were rapidly excised and placed in pre-cooled PBS. Under sterile conditions, the lung tissues were thoroughly washed in PBS to remove blood. The washed tissues were then transferred to a sterile cutting board, where they were minced into 1–2 mm pieces using surgical scissors and tweezers. These tissue pieces were transferred to a digestion buffer containing collagenase (Beyotime, cat. no. ST2303) and DNase (Beyotime, cat. no. D7073), and incubated at 37°C with gentle shaking for 1–2 hours to dissociate the tissue and release individual cells.

Following digestion, the mixture was filtered through a 100 µm cell strainer to remove undigested tissue fragments. The collected cell suspension was then centrifuged (300×g for 5 minutes), and the supernatant was discarded. Subsequently, the cell suspension was treated with red blood cell lysis buffer (Beyotime, cat. no. C3702) to eliminate red blood cells. After 5 minutes, the suspension was centrifuged again, the supernatant was discarded, and the cells were resuspended in PBS warmed to room temperature.

Western Blot Analysis of Mitochondrial Complex I Expression

Initially, total protein was extracted from the target cells or tissues, and the protein concentration was determined using a BCA Protein Assay Kit (Beyotime, cat. no. P0010). The protein samples were then mixed with SDS loading buffer and boiled at 95°C for 10 minutes to denature the proteins. Subsequently, the samples (15 µg) were loaded onto an SDS-PAGE gel for electrophoretic separation. After electrophoresis, the proteins were transferred to a PVDF membrane, which was then blocked with 5% BSA solution to prevent non-specific binding, and incubated at room temperature for 1 hour. The membrane was then incubated overnight at 4°C with a specific primary antibody against mitochondrial Complex I (Abcam, cat. no. ab110245).

The next day, the membrane was washed three times with TBS containing 0.1% Tween-20 for 10 minutes each to remove excess primary antibody. This was followed by incubation with the corresponding HRP-conjugated secondary antibody (Beyotime, cat. no. A0208) at room temperature for 1–2 hours. After the incubation with the secondary antibody, the membrane was washed again and then treated with chemiluminescent substrate to detect the protein signal. To ensure consistency in protein loading, the membrane was also probed with an antibody against the internal control protein β-actin (Beyotime, cat. no. AF0003).

Flow Cytometry Analysis of CFSE Mean Fluorescence Intensity in MPMECs

In this study, flow cytometry was utilized to detect CFSE staining in MPMECs to evaluate cell proliferation and division. Initially, MPMECs were cultured to 50% confluency, and CFSE (Invitrogen, cat. no. C34570) was added to the culture at a concentration of 5 µM. The cells were then incubated at 37°C with 5% CO2 for 15–20 minutes. During this period, CFSE permeated the cell membrane and bound to intracellular amino acids, forming a stable fluorescent complex. After incubation, cells were washed with PBS and resupplied with complete culture medium. Subsequently, 1×10^5 MSCs were added to the MPMEC culture system according to the experimental groups, and after 24 hours of co-culture, the treated cells were collected, washed with PBS, labeled with CD31 antibody to mark MPMECs, and prepared for flow cytometry analysis. In the flow cytometer, appropriate lasers and detectors were set to differentiate MPMECs while simultaneously measuring the mean fluorescence intensity of CFSE in MPMECs. Data were analyzed using FlowJo software, and cell proliferation was assessed by calculating the mean fluorescence intensity of CFSE.

Flow Cytometry Analysis of ROS Mean Fluorescence Intensity in MPMECs

Flow cytometry was employed to assess the levels of reactive oxygen species (ROS) in MPMECs. MPMECs were cultured to 50% confluency and subjected to cell treatments according to experimental groups, with the MSC-treated experimental group receiving 1×10^5 MSCs. After 24 hours of co-culture with MSCs, the cells were collected. A specific ROS assay kit (Elabscience, cat. no. E-BC-K138-F) was used to treat the cell samples. Briefly, the ROS probe DCFDA was used for ROS detection, dissolving DCFDA at 10 µM in serum-free culture medium, and incubating MPMECs with this solution for approximately 20–30 minutes at 37°C. During incubation, DCFDA penetrated the cells and was converted into a fluorescent compound under the influence of ROS. After incubation, cells were washed with PBS to remove uninternalized or unreacted DCFDA. Subsequently, cells were labeled and distinguished as MPMECs using a CD31 antibody. Flow cytometry analysis was performed to analyze the fluorescence signal, detecting and assessing the mean fluorescence intensity of ROS in CD31-positive cells.

Flow Cytometry Analysis of Cell Apoptosis in MPMECs

Flow cytometry was utilized to assess apoptosis in MPMECs. Prior to the experiment, MPMECs were cultured to an appropriate density and subjected to experimental treatments, with 1×10^5 MSCs added to the MSC-treated group. After cell treatment, an Annexin V/PI Apoptosis Detection Kit (Elabscience, cat. no. E-CK-A211) was used for labeling and analysis. Firstly, the treated cells were collected and washed with PBS to remove serum and dead cells from the culture medium. The cell concentration was adjusted to 1×10^6 cells/mL, and then cells were resuspended in staining buffer containing Annexin V-FITC and PI, gently mixed to avoid excessive agitation. The cells were incubated in the dark at room temperature for 15–20 minutes. Following staining, MPMECs were labeled with CD31, and then analyzed using flow cytometry.

Using flow cytometry software, CD31 positive MPMECs were selected, and cells were categorized into early apoptotic cells (Annexin V positive/PI negative), late apoptotic or necrotic cells (Annexin V positive/PI positive), and viable cells (Annexin V negative/PI negative). Based on these classifications, the level of apoptosis in MPMECs could be quantitatively analyzed.

Hematoxylin and Eosin (H&E) Staining and Lung Injury Scoring of Mouse Lung Tissue

In this study, mouse lung tissues were stained with Hematoxylin and Eosin (H&E) to assess the extent of lung injury. Firstly, after experimental treatment, mice were euthanized, and lung tissues were rapidly excised. The tissues were fixed in 4% paraformaldehyde solution for at least 24 hours to preserve the integrity of the tissue structure. After fixation, the lung tissues underwent dehydration, clearing, and paraffin infiltration, followed by embedding in paraffin blocks. The paraffin-embedded lung tissues were sectioned into 4–5 µm thick slices using a microtome and placed on slides. The slides were then dried at 60°C for about 30 minutes to enhance adhesion to the slides. Subsequently, H&E staining was performed. The sections were deparaffinized, hydrated through a series of graded ethanol solutions, stained with hematoxylin for 3–5 minutes to color the nuclei, rinsed with running water, and stained with eosin for 1–2 minutes to color the cytoplasm and other tissue structures. The sections were then dehydrated in ascending ethanol series, cleared, and mounted. After staining, images were observed and captured using an optical microscope.

The pathological injury of lung tissue was scored using the Smith scoring method. Lung edema, alveolar and interstitial inflammation, alveolar and interstitial hemorrhage, atelectasis, and hyaline membrane formation were semi-quantitatively analyzed with a score ranging from 0 to 4; where no injury scored 0, lesion area < 25% scored 1, 25%-50% scored 2, 50%-75% scored 3, and lesion filling the field of view scored 4. The total lung injury score was the sum of these items, with the average score calculated from 10 high-power fields per animal.

Quantitative Real-Time PCR Analysis

The quantitative Real-Time PCR (qRT-PCR) technique was employed to analyze the mRNA expression levels of specific genes. Initially, total RNA was extracted from samples such as cells and tissues using an RNA extraction kit (Takara, cat. no. 9109), following the manufacturer’s guidelines. The concentration and purity of the extracted RNA were determined using a spectrophotometer to ensure RNA quality. Prior to qRT-PCR, the extracted total RNA was transcribed into cDNA using reverse transcriptase and specific primers as per the instructions of the reverse transcription kit (Takara, cat. no. RR047A). Upon completion of reverse transcription, the cDNA was used as a template for qRT-PCR analysis. The qRT-PCR reaction was carried out in a total volume of 10 µL, following the reaction system setup as described in the kit instructions (Takara, cat. no. RR420A). To quantify the expression levels of target genes, the internal reference gene β-actin was used as a control. The expression level changes of the target genes were calculated using the relative quantification method (2^-ΔΔCt method), by comparing the threshold cycle numbers (Ct values) of the target genes with that of the internal reference gene.

Enzyme-Linked Immunosorbent Assay

Enzyme-linked immunosorbent assay (ELISA) technology was utilized for the quantitative analysis of specific protein expression levels. An appropriate ELISA kit was selected based on the target protein. All procedures were conducted strictly following the instructions provided by the kit manufacturer.

Immunohistochemistry and HALO Quantitative Analysis

Immunohistochemistry was employed to localize and quantitatively analyze specific proteins in mouse lung tissues. Following experimental treatment, mice were euthanized, and lung tissues were rapidly excised. The excised tissues were fixed in 4% paraformaldehyde solution for 24 hours, then dehydrated, cleared, and embedded in paraffin. The paraffin-embedded tissues were sectioned into continuous 4–5 µm thick slices and placed on slides. The sections underwent deparaffinization and hydration, followed by antigen retrieval using microwave treatment to expose protein epitopes. Subsequently, the sections were treated with 3% hydrogen peroxide for 10 minutes to block endogenous peroxidase activity. The sections were then blocked with 5% BSA at room temperature for 30 minutes to prevent non-specific binding. Next, the sections were incubated with primary antibodies specific to the target protein, typically overnight at 4°C. The following day, after washing the sections, they were incubated with the corresponding secondary antibodies for 1 hour. Color development was then performed using a diaminobenzidine (DAB) chromogen system, where DAB reacts with hydrogen peroxide to produce a brown precipitate, marking positive signals. Finally, the nuclei were counterstained with hematoxylin and the sections were mounted.

For quantitative analysis of the immunohistochemically stained sections, HALO image analysis software was used. Images of the sections under the microscope were imported into the HALO software, and the software's built-in algorithms were employed to quantitatively analyze the positive signals of the specific protein. This analysis included assessment of the intensity of positive staining, distribution range, and the number of positive cells. By comparing these parameters across different experimental groups, quantitative evaluation of the expression differences of specific proteins in lung tissues was conducted.

ATP Quantitative Analysis in MPMECs

The ATP levels in MPMECs were quantitatively analyzed using an ATP Assay Kit (Elabscience, cat. no. E-BC-F002). Initially, MPMECs were cultured to the required density and subjected to appropriate treatments as per experimental requirements. After treatment, cells were lysed using cell lysis buffer to release ATP. To ensure efficient lysis, the lysed cells were incubated on ice for 10–15 minutes and subjected to repeated pipetting or gentle vortexing for thorough contact with the lysis buffer. Following the instructions of the ATP assay kit, the cell lysate was centrifuged (10000×g, 4°C, for 5 minutes) to remove cell debris. The supernatant was then used for ATP measurement.

In a 96-well plate, 100 µL of sample and ATP detection reagent were added and mixed well, followed by incubation at room temperature for 5–10 minutes. The luminescence intensity was measured using a luminometer, and the ATP concentration in the samples was calculated based on a standard curve. Each sample was assayed in triplicate to ensure the accuracy and reproducibility of the results.

Immunofluorescence Analysis of FAS Protein in MPMECs

Immunofluorescence staining was used to analyze the expression of FAS protein in MPMECs. Initially, MPMECs were cultured on microscope-compatible slides and treated according to experimental groups. Upon reaching appropriate confluency, the cells were fixed with 4% paraformaldehyde solution (Biosharp, cat. no. BL539A) for 20 minutes to preserve cellular morphology and structure. After fixation, cells were washed three times with PBS to remove the fixative. The cells were then permeabilized with 0.1% Triton X-100 (Beyotime, cat. no. P0096) for 10 minutes to allow antibody penetration. Following permeabilization, the cells were again washed with PBS.

To prevent non-specific binding, cells were blocked with 1% BSA (BioFroxx, cat. no. 4240GR100) and incubated for 30 minutes. Subsequently, the cells were incubated overnight at 4°C with a specific primary antibody against FAS protein (Servicebio, cat. no. GB12089-100). The next day, cells were washed with PBS to remove unbound primary antibody, followed by incubation with a fluorescently-labeled secondary antibody (Abcam, cat. no. ab150120) for 1 hour. After secondary antibody incubation, cells were washed again with PBS. Finally, for nuclear staining, DAPI fluorescent dye (Beyotime, cat. no. C1005) was applied. The expression and localization of FAS protein were observed and captured using a fluorescence microscope.

Mouse Lung Wet-to-Dry Weight Ratio

To assess the extent of pulmonary edema, we measured the wet-to-dry weight ratio of the mouse lungs. Initially, mice were subjected to the respective experimental treatments. After the completion of treatments, the mice were euthanized, and their total body weight was accurately measured. The thoracic cavity was then immediately opened to excise the lung tissues. Upon extraction, the lung tissues were first gently rinsed at room temperature with saline to remove blood and adherent materials from the lung surface, followed by gentle dabbing with filter paper to remove surface moisture. Subsequently, the wet weight of the lungs was quickly and accurately measured using a precision electronic balance to ensure accuracy. The lung wet-to-dry weight ratio was calculated by dividing the wet weight of the lungs by the total body weight of the mouse, expressed in mg/g.

Evans Blue Permeability Assay in Mouse Pulmonary Microvasculature

To assess the permeability of the pulmonary microvasculature in mice, an Evans Blue dye assay (Solarbio, cat. no. IE0280) was conducted. Prior to the experiment, mice underwent the designated treatments. At specific time points during the experiment, Evans Blue dye was injected via the tail vein at a dosage of 30 ug/g body weight. After the dye injection, it was allowed to circulate in the system for 30 minutes, ensuring adequate time for the dye to bind to plasma albumin and reach equilibrium. Following the injection period, mice were euthanized, and lung tissues were rapidly excised. The lung tissues were first gently rinsed at room temperature with saline to remove surface blood and adherent materials. Subsequently, 100 mg of minced lung tissue was fixed in 1 mL of methanol for 24 hours to extract Evans Blue dye from the tissue. The absorbance of the solution was then measured at a wavelength of 620 nm using a spectrophotometer to quantify the Evans Blue content in the lung tissues. The content of Evans Blue in the lung tissues was calculated by comparing the measured absorbance values with a known concentration Evans Blue standard curve and expressed in µg/mL wet weight.

Statistical Analysis

Data analysis was conducted using GraphPad Prism (Version 9.0.2). The statistical significance between the experimental and control groups was determined through the use of independent sample t-tests for pairwise comparisons and one-way ANOVA for analyses involving multiple groups. Post-hoc analysis for significant ANOVA results was carried out using Tukey's test to compare means between groups. A threshold of p < 0.05 was established for statistical significance, ensuring that findings were rigorously evaluated for their reliability and validity within the context of our research objectives.

Alleviation of Endothelial Injury in ARDS by MSCs through Mitochondrial Transfer

To validate the hypothesis that MSCs can alleviate endothelial damage in ARDS through mitochondrial transfer, we first verified the capability of MSCs to transfer mitochondria to MPMECs. MPMECs damaged with rotenone were co-cultured with MSCs labeled with MitoTracker Deep Red for 24 hours. Flow cytometry analysis confirmed the presence of MitoTracker Deep Red-labeled mitochondria in MPMECs (Fig. 1A), suggesting mitochondrial transfer from MSCs to the damaged endothelial cells. Furthermore, when MSCs were subjected to rotenone-induced mitochondrial damage, a significant reduction in mitochondrial transfer to MPMECs was observed (Fig. 1B). Subsequent in vivo experiments were performed to confirm mitochondrial transfer from MSCs to endothelial cells in an ARDS mouse model. Four hours after ARDS induction, MSCs labeled with MitoTracker Deep Red were administered intravenously. After 24 hours, lung tissues were harvested for frozen sectioning, and endothelial cells were identified using CD31 immunostaining. Immunofluorescence analysis revealed the presence of MitoTracker Deep Red-labeled mitochondria within CD31-positive endothelial cells, indicating mitochondrial transfer from MSCs to endothelial cells in ARDS mice (Fig. 1C). Additionally, single-cell suspensions prepared from the mouse lung tissues and stained with CD31 showed approximately 39% of endothelial cells had received mitochondria from MSCs, as determined by flow cytometry (Fig. 1D).

We further explored the reparative role of MSC-mediated mitochondrial transfer on endothelial damage. Animal studies demonstrated that intravenous injection of MSCs increased the expression of mitochondrial complex I in lung tissues of ARDS mice (Fig. 1E and 1F), suggesting a potential mitigative effect of MSCs on mitochondrial damage in ARDS. Co-culturing MSCs with mitochondrially damaged MPMECs also resulted in increased mitochondrial complex I expression in these cells. This effect was diminished when mitochondrial transfer was inhibited by damaging the mitochondria of MSCs with rotenone (Fig. 1G and 1H), indicating that mitochondrial transfer is a key mechanism through which MSCs repair mitochondrial damage in MPMECs. Additionally, MSCs were found to reduce ROS production (Fig. 1I) and decrease apoptosis (Fig. 1J) in mitochondrially damaged MPMECs, but after inhibiting mitochondrial transfer by MSCs, their effect in reducing ROS and apoptosis in MPMECs was significantly diminished, further supporting the therapeutic role of MSCs in repairing mitochondrial damage through transfer mitochondria. Moreover, a comparative assessment of lung tissue pathology in ARDS mice treated with MSCs versus those with inhibited mitochondrial transfer revealed a reduced efficacy in repairing lung tissue pathology when mitochondrial transfer was impeded (Fig. 1K and 1L). Overall, these findings substantiate the efficacy of MSCs in mitigating endothelial and pulmonary damage in ARDS through mitochondrial transfer.

MSCs Promote Endothelial Cell Proliferation and Vascular Regeneration in ARDS Lung Tissue via Mitochondrial Transfer

To investigate whether MSCs could facilitate the proliferation of mitochondrially impaired MPMECs through mitochondrial transfer, we co-cultured MSCs with damaged MPMECs for 24 hours. Flow cytometry revealed a reduction in CFSE fluorescence intensity in mitochondrially damaged MPMECs (Fig. 2A), and an upregulation of the proliferation gene Ki67 in MPMECs was observed (Fig. 2B). These findings suggest that MSCs can promote proliferation in mitochondrially damaged MPMECs. This proliferative effect was significantly reduced when MSCs with mitochondria damaged by rotenone were used, indicating that mitochondrial transfer from MSCs is crucial for the proliferation of damaged MPMECs. Further experiments using Transwell chambers to isolate MSCs from damaged MPMECs for 24 hours showed that MSCs could still significantly reduce CFSE fluorescence intensity in MPMECs (Fig. 2C) and increase the expression of the cell proliferation gene Ki67 (Fig. 2D). This suggests that MSCs can promote MPMEC proliferation even when physically separated, likely through mitochondrial transfer. When MSCs were treated with rotenone to impair mitochondrial function, the enhancement of MPMEC proliferation was notably decreased. Additionally, animal experiments showed that intravenous injection of MSCs increased the expression of the proliferation gene Ki67 in lung tissues of ARDS mice (Fig. 2E). However, this effect was significantly diminished when the mitochondrial function of MSCs was impaired by rotenone, confirming the role of MSC-mediated mitochondrial transfer in promoting endothelial cell proliferation.

We also examined whether MSCs could promote vascular regeneration in ARDS through mitochondrial transfer. Co-culturing mitochondrially damaged MPMECs with MSCs for 24 hours significantly increased the expression of vascular growth factors VEGF and HGF mRNA in MPMECs (Fig. 2F and 2G). This effect was markedly reduced when MSCs with impaired mitochondria (induced by rotenone) were used, suggesting that mitochondrial transfer from MSCs enhances the release of pro-angiogenic factors from MPMECs. Using Transwell chambers to isolate MSCs from MPMECs, we observed a similar upregulation in the expression of VEGF and HGF mRNA in MPMECs (Fig. 2H and 2I). However, when MSCs were treated with rotenone to inhibit mitochondrial transfer, the expression levels of these angiogenic factors significantly decreased. Further animal studies revealed that intravenous injection of MSCs increased the expression of VEGF and HGF mRNA in lung tissues of ARDS mice (Fig. 2J and 2K). This increase was notably less pronounced when MSCs with impaired mitochondria were used. Moreover, ELISA analysis of cell culture supernatants showed that MSCs could enhance the release of VEGF and HGF from mitochondrially damaged MPMECs (Fig. 2L and 2M), and this capability was significantly diminished when MSCs with damaged mitochondria were used. Finally, immunohistochemical analysis of lung tissues from ARDS mice treated with MSCs revealed improved vascular integrity compared to untreated ARDS mice. This reparative effect on lung tissue vasculature was significantly reduced when MSCs with impaired mitochondria were used (Fig. 2N and 2O), confirming that MSCs promote endothelial cell secretion of vascular growth factors and vascular regeneration in ARDS through mitochondrial transfer.

MSCs Transfer Mitochondria to MPMECs via TNTs

Previous studies have reported that MSCs can transfer mitochondria to ARDS alveolar epithelial cells via TNTs[25, 26]. However, it remains unclear whether MSCs utilize TNTs for mitochondrial transfer to endothelial cells in ARDS. To investigate this, we pre-stained mitochondria of MPMECs with MitoTracker Green and added MSCs with mitochondria pre-labeled with MitoTracker Deep Red to the MPMEC culture system for a 12-hour co-culture. Confocal microscopy observations revealed the formation of TNTs between MSCs and MPMECs, through which MSCs transferred mitochondria to MPMECs (Fig. 3A). Building upon these findings, we further examined the impact of inhibiting TNT formation on the efficiency of mitochondrial transfer from MSCs to MPMECs. MSCs with pre-labeled mitochondria were co-cultured with MPMECs for 24 hours, and Cytochalasin B was added at various concentrations to inhibit TNT formation. We observed a significant reduction in the efficiency of mitochondrial transfer with the inhibition of TNTs, and no significant difference in the inhibitory effect on mitochondrial transfer was noted between 500 nM, 1000 nM, and 2000 nM concentrations of Cytochalasin B (Fig. 3B). Therefore, 500 nM Cytochalasin B was selected for subsequent experiments to inhibit TNT formation.

We also assessed the impact of TNT inhibition on ROS production and apoptosis in MPMECs. Co-culturing mitochondrially impaired MPMECs with MSCs for 24 hours, flow cytometry indicated that MSCs significantly reduced ROS production and apoptosis in MPMECs. However, the addition of Cytochalasin B to inhibit TNT formation markedly decreased MSCs' ability to reduce ROS generation and MPMEC apoptosis (Fig. 3C and 3D), suggesting that MSCs transfer mitochondria to mitochondrially damaged MPMECs via TNTs, thereby exerting their reparative effects. Furthermore, we explored the influence of inhibiting TNT formation between MSCs and MPMECs on the expression of mitochondrial fusion genes Mfn1 and Mfn2. Co-culturing MSCs with damaged MPMECs for 24 hours resulted in a significant increase in the expression of Mfn1 and Mfn2 in MPMECs, indicating mitochondrial fusion post-mitochondrial transfer from MSCs. In contrast, the addition of Cytochalasin B to inhibit TNT formation significantly decreased the expression of Mfn1 and Mfn2 in MPMECs (Fig. 3E and 3F), thereby confirming that MSCs can transfer mitochondria to mitochondrially damaged MPMECs via TNTs.

MPMECs Uptake Mitochondria from MSCs via Dynamin-Dependent Clathrin-Mediated Endocytosis

Previous studies have shown that MSCs can transfer mitochondria to recipient cells via extracellular vesicles[20]. To investigate whether MSCs transfer mitochondria to MPMECs under isolated conditions, we employed a Transwell chamber system. MPMECs with mitochondria pre-stained with MitoTracker Deep Red were cultured in the lower chamber, while MSCs with mitochondria pre-labeled with MitoTracker Green were cultured in the upper chamber of the Transwell. After 24 hours, fluorescence microscopy revealed co-localization of both mitochondrial stains in MPMECs (Fig. 4A), suggesting mitochondrial transfer from MSCs to MPMECs even when physically separated. However, the mechanism through which MPMECs internalize mitochondria from MSCs was unclear. We discovered that under isolated culture conditions, the mitochondrial transfer from MSCs to MPMECs could be inhibited by dynasore, a dynamin-dependent clathrin-mediated endocytosis inhibitor (Fig. 4B). This finding suggests that MPMECs might uptake mitochondria from MSCs through dynamin-dependent endocytosis. Further experiments using the Transwell chamber assessed the impact of inhibiting endocytosis on ROS production and apoptosis in MPMECs. We found that isolated co-culture of MSCs with mitochondrially damaged MPMECs reduced ROS production and apoptosis in MPMECs, but the reparative effects of MSCs were diminished when MPMEC endocytosis was inhibited by dynasore (Fig. 4C and 4D). Additionally, we continued to investigate the effect of inhibiting endocytosis in MPMECs on the expression of mitochondrial fusion genes Mfn1 and Mfn2. Isolated co-culture of MSCs with damaged MPMECs increased the expression of mitochondrial fusion genes in MPMECs. However, inhibiting endocytosis in MPMECs significantly reduced the expression of Mfn1 and Mfn2 (Fig. 4E and 4F), confirming that MPMECs internalize mitochondria from MSCs via dynamin-dependent clathrin-mediated endocytosis.

Mitochondrial Transfer from MSCs to MPMECs Enhances Fatty Acid Synthesis, Facilitating Vascular Regeneration in ARDS

As previously mentioned, MSCs can promote vascular regeneration in ARDS through mitochondrial transfer, though the underlying mechanisms remain unclear. Given the role of mitochondria as cellular powerhouses, we first examined whether MSCs alter the energy metabolism of MPMECs via mitochondrial transfer. After co-culturing MSCs with mitochondrially damaged MPMECs, we observed an increase in ATP production in MPMECs. However, this increase was not diminished when mitochondrial transfer was inhibited by treating MSCs with rotenone (Fig. 5A), suggesting that the MSCs’ role in ATP production in MPMECs is independent of mitochondrial transfer. Previous studies have demonstrated the critical role of fatty acid synthesis in angiogenesis. We investigated whether MSCs regulate lipid metabolism in MPMECs through mitochondrial transfer. After a 24-hour co-culture of MSCs with mitochondrially damaged MPMECs, PCR analysis showed a significant increase in the expression of key enzymes in fatty acid synthesis, including FAS, ACC, and ACLY, in MPMECs. This increase was markedly reduced when mitochondrial transfer from MSCs was inhibited (Fig. 5B, 5C, and 5D), indicating that MSCs modulate fatty acid synthesis in MPMECs through mitochondrial transfer. Further animal experiments confirmed that intravenous injection of MSCs enhanced the expression of these key enzymes in fatty acid synthesis in lung tissues of ARDS mice. This enhancement was significantly decreased when mitochondrial transfer was inhibited by damaging the mitochondria of MSCs (Fig. 5E, 5F, and 5G), suggesting that MSCs regulate fatty acid metabolism in ARDS lung tissues through mitochondrial transfer.

To further validate that MSCs mediate vascular regeneration by regulating lipid metabolism in MPMECs, we inhibited fatty acid synthesis in MPMECs using C75, a fatty acid synthase inhibitor, and observed its effects on the angiogenic factors VEGF and HGF. Initially, the inhibitory effect of C75 on fatty acid synthesis was confirmed through immunofluorescence detection of FAS expression post 12-hour co-culture of MSCs with mitochondrially damaged MPMECs. MSCs were found to promote FAS expression and cell proliferation in MPMECs; however, addition of C75 significantly inhibited both FAS expression and MPMEC proliferation (Fig. 5H). Building upon this, we observed that co-culturing MSCs with MPMECs enhanced the expression of angiogenic factors VEGF and HGF mRNA, while the addition of C75 significantly suppressed their expression (Fig. 5I and 5J). This indicates that MSCs can mediate the angiogenic activity of MPMECs through the regulation of fatty acid synthesis. Furthermore, the addition of C75 to the co-culture system significantly reduced the levels of VEGF and HGF in the cell culture supernatant (Fig. 5K and 5L), suggesting that C75 inhibits the secretion of angiogenic factors. Overall, these findings substantiate the role of MSCs in mediating vascular regeneration in ARDS through the regulation of fatty acid synthesis via mitochondrial transfer.

Stimulation of the TCA Cycle by MSC-Transferred Mitochondria Activates Citrate-Dependent Fatty Acid Synthesis in MPMECs

While it's known that MSCs can facilitate vascular regeneration in ARDS through mitochondrial transfer, the specific pathways through which this occurs, particularly in relation to fatty acid synthesis, are not fully understood. Given that mitochondria are central to the TCA cycle, an upstream pathway of fatty acid synthesis, we first investigated whether MSC-mediated mitochondrial transfer stimulates the TCA cycle in MPMECs. We co-cultured MPMECs with both functionally intact and mitochondrial damaged MSCs for 24 hours and measured the expression of key TCA cycle enzymes CS, IDH, and OGDH using PCR. The results showed a significant decrease in the expression of these enzymes in the group with damaged MSCs (Fig. 6A, 6B, and 6C), indicating that mitochondrial transfer from MSCs activates the TCA cycle in MPMECs. We then assessed the impact of TCA cycle inhibition on fatty acid synthesis in MPMECs. Upon adding Devimistat, a TCA cycle inhibitor, to the co-culture system, we observed a notable decrease in CS mRNA expression in MPMECs (Fig. 6D), confirming the inhibitory effect of Devimistat on the TCA cycle. Further PCR analysis of key fatty acid synthesis enzymes FAS, ACC, and ACLY revealed a significant increase in their expression following MSC co-culture. However, this increase was substantially reduced upon the addition of Devimistat (Fig. 6E, 6F, and 6G), suggesting that mitochondrial transfer from MSCs to MPMECs promotes fatty acid synthesis by stimulating the TCA cycle.

Citrate, a critical link between cellular TCA cycle and fatty acid synthesis, was also studied. After 24 hours of co-culture of MSCs with damaged MPMECs, the addition of Devimistat alone or in combination with citrate was tested. Immunofluorescence analysis of FAS indicated that inhibiting the TCA cycle with Devimistat reduced FAS expression and cell proliferation, whereas the addition of citrate increased FAS expression (Fig. 6H). Further PCR analysis of FAS, ACC, and ACLY mRNA expression revealed that citrate partially restored the expression of these key enzymes in fatty acid synthesis (Fig. 6I, 6J, and 6K). These findings collectively confirm that mitochondrial transfer from MSCs to MPMECs stimulates the TCA cycle, thereby activating citrate-dependent fatty acid synthesis.

Citrate Restores the Angiogenic Potential of Mitochondrially Impaired MSCs

To explore the role of citrate in restoring the angiogenic potential of mitochondrially impaired MSCs in ARDS, we first conducted cell experiments to observe the effects of citrate on fatty acid synthesis in MPMECs co-cultured with damaged MSCs. Upon adding citrate to the culture system, it was observed that mitochondrially impaired MSCs alone did not increase the expression of key fatty acid synthesis enzymes FAS, ACC, and ACLY mRNA in MPMECs. However, the addition of citrate significantly enhanced the expression of these enzymes (Fig. 7A, 7B, and 7C), suggesting that citrate can restore the fatty acid synthesis-promoting function of damaged MSCs. Further, we examined whether the addition of citrate in the co-culture system could restore the angiogenic function of damaged MSCs. PCR analysis of MPMECs revealed that mitochondrially impaired MSCs did not increase the expression of angiogenic factors VEGF and HGF mRNA, whereas their expression significantly increased after adding citrate to the culture system (Fig. 7D and 7E). Additionally, ELISA was used to measure the concentrations of VEGF and HGF in the cell culture supernatant, assessing the secretion of angiogenic factors. The results showed a significant increase in the secretion of VEGF and HGF by MPMECs upon the addition of citrate (Fig. 7F and 7G), indicating that citrate can restore the ability of damaged MSCs to promote MPMEC-mediated angiogenic factor release.

Further animal studies were conducted to observe the effect of citrate in restoring the angiogenic potential of damaged MSCs and in repairing lung injury in ARDS. HE staining was used to examine lung injury in ARDS mice, and immunohistochemical staining of vascular endothelial cells was performed to assess the vascular morphology in lung tissues. The results showed that intravenous injection of MSCs reduced lung injury and improved the integrity of lung tissue vasculature in ARDS mice. This reparative effect was diminished when the mitochondria of MSCs were damaged with rotenone. However, treating mitochondrially impaired MSCs with citrate enhanced their protective effect on lung injury and vascular integrity in ARDS mice (Fig. 7H). These findings collectively confirm that citrate can restore the angiogenic potential of mitochondrially impaired MSCs.

MSCs Alleviate LPS-Induced Lung Injury

Finally, we evaluated the reparative effects of MSCs on LPS-induced lung injury. An ARDS mouse model was induced by intratracheal instillation of LPS, followed by the intravenous injection of MSCs four hours post-model establishment. Pathological examination revealed that LPS-induced lung injury in ARDS mice was significantly more severe compared to the control and sham groups. Intravenous administration of MSCs notably reduced the extent of lung pathology in ARDS mice (Fig. 8A and 8B). Additionally, immunohistochemical staining for the cell adhesion protein Occludin showed that MSC treatment significantly increased Occludin expression (Fig. 8C). Quantitative analysis of Occludin immunostaining using HALO software indicated a significant increase in Occludin-positive cells following MSC treatment (Fig. 8D). Further, we evaluated the expression of inducible and endothelial nitric oxide synthase (iNOS and eNOS, respectively) in mouse lung tissues using ELISA. We observed that iNOS expression was significantly elevated, and eNOS expression was reduced in ARDS mouse lungs. MSC treatment decreased iNOS expression and increased eNOS expression, thereby exerting a protective effect on the lungs (Fig. 8E and 8F). The lung wet-to-dry weight ratio was used to assess pulmonary edema, revealing a significant increase in ARDS mice compared to healthy controls. This ratio was reduced in ARDS mice treated with MSCs, indicating an improvement in pulmonary edema. Furthermore, the Evans Blue assay was used to evaluate the permeability of pulmonary capillaries. The results showed a significant increase in Evans Blue content in the lungs of ARDS mice, which was markedly reduced following MSC treatment, indicating a decrease in pulmonary capillary permeability. In summary, our findings demonstrate that MSCs can effectively alleviate lung injury in ARDS.

ARDS is a severe pulmonary inflammatory response characterized primarily by extensive alveolar damage and capillary leakage. Mitochondrial injury is one of the key factors in the pathophysiology of ARDS, as mitochondria play a central role in maintaining cellular function and energy metabolism. In ARDS, mitochondrial function in lung tissue cells is severely compromised due to inflammatory responses and oxidative stress[27]. Currently, there is a lack of effective treatments specifically targeting mitochondrial damage in ARDS. MSCs, a type of pluripotent stem cells capable of self-renewal and differentiation into various cell types, have gained attention in recent years for their potential in tissue repair, immunomodulation, and anti-inflammatory actions[28]. Our study demonstrates that MSCs can repair mitochondrial damage in ARDS endothelial cells through mitochondrial transfer. This process activates the TCA cycle and fatty acid synthesis in endothelial cells, leading to enhanced cell proliferation and the release of pro-angiogenic factors, which subsequently promote vascular regeneration. These findings provide a novel perspective on ARDS treatment, underscoring the therapeutic potential of MSCs in mitigating mitochondrial damage in pulmonary endothelial cells and enhancing vascular repair.

MSCs can transfer mitochondria to the pulmonary endothelial cells in ARDS, a critical process for alleviating pulmonary endothelial injury. Specifically, the transfer of healthy mitochondria from MSCs to ARDS endothelial cells results in a significant increase in mitochondrial complex I expression, decrease in ROS production and endothelial cell apoptosis. These actions are vital for maintaining the integrity and function of pulmonary vasculature in ARDS. Previous studies support the role of MSC mitochondrial transfer in cell protection, metabolic regulation, and oxidative stress mitigation[29]. For instance, earlier research has demonstrated that MSCs can promote repair and regeneration of damaged lung tissue[30, 31]. Additionally, studies have highlighted the importance of MSC mitochondrial transfer in alleviating metabolic disturbances and reducing oxidative stress[32]. Our study underscore the immense potential of MSC-based mitochondrial transfer therapies in improving the prognosis of ARDS patients.

MSCs can significantly enhance the proliferation of MPMECs in ARDS through mitochondrial transfer and stimulate the release of angiogenic factors, thereby promoting vascular regeneration. Mitochondria, as critical organelles in cellular energy metabolism, play an essential role in promoting cell proliferation and functional recovery[33]. In our study, MSCs provided healthy mitochondria to MPMECs via mitochondrial transfer, which not only likely improved the energy metabolism of the damaged cells but also may have stimulated cell proliferation by activating relevant signaling pathways. This is consistent with earlier studies demonstrating that MSCs can promote cell proliferation and functional recovery through various mechanisms, including secretory factors and cell-to-cell contact[34, 35]. Additionally, the release of angiogenic factors by MPMECs is crucial for promoting angiogenesis. Angiogenesis is a key process for restoring normal vascular function and improving tissue oxygenation, especially in ARDS, where lung vascular damage and inflammation are significant components of the pathology. Therefore, by promoting MPMEC proliferation and the release of angiogenic factors, MSCs provide potential therapy for the repair of lung endothelial injury in ARDS.

MSCs can transfer mitochondria to MPMECs in ARDS via TNTs. TNTs represent a specialized form of intercellular communication, consisting of long, thin cellular protrusions that allow direct transfer of organelles and signaling molecules between cells[36]. In our research, TNTs formed a 'bridge' between MSCs and MPMECs, enabling MSCs to directly transfer mitochondria to mitochondrially impaired endothelial cells. Previous studies have indicated the presence of TNTs in various cell types[37]. For instance, one study demonstrated that cardiomyocytes transfer mitochondria via TNTs in a heart disease model, aiding in the repair and functional recovery of damaged cells[38]. Another study found that neuronal cells transfer mitochondria through TNTs in models of neurological diseases, playing a crucial role in cell survival and the stability of neural networks[39, 40]. Given that ARDS involves extensive pulmonary inflammation and oxidative stress in endothelial cells, maintaining and restoring endothelial cell function is key to treatment[41]. The mitochondria transferred via TNTs could provide the necessary energy support and metabolic substances to the damaged endothelial cells, helping to maintain cellular activity, reduce cell death, and thus promote lung repair and functional recovery.

Mitochondrially impaired MPMECs can internalize healthy mitochondria from MSCs through dynamin-dependent clathrin-mediated endocytosis. The identification of this mechanism offers a new perspective in understanding intercellular mitochondrial transfer. Dynamin, a GTPase assisting in membrane invagination and vesicle formation at the cell membrane, and clathrin, a protein involved in vesicle transport and intracellular material translocation, play crucial roles in this process[42]. In our research, we used Dynasore, an inhibitor of dynamin-dependent clathrin-mediated endocytosis, and found that the ability of MPMECs treated with Dynasore to accept mitochondria was significantly reduced. Although previous studies reported mitochondrial transfer between cells via extracellular vesicles[43], they did not fully elucidate how mitochondria are specifically accepted and internalized by certain cells. The mechanism of dynamin-dependent clathrin-mediated endocytosis provides a potential method for the reception of mitochondria by recipient cells, adding a new dimension to the mechanisms of mitochondrial transcellular transfer.

MSCs promote fatty acid synthesis in MPMECs via mitochondrial transfer, subsequently enhancing the release of angiogenic factors by MPMECs. Intriguingly, when fatty acid synthesis in MPMECs is inhibited, their capacity to release angiogenic factors is also significantly reduced. This finding underscores the critical role of fatty acid synthesis in the process of angiogenesis. Fatty acid synthesis, a key aspect of cellular metabolism, plays an important role not only in energy storage and provision for cells but also in numerous cellular functions, including cell signaling, membrane construction, and molecular regulation[44]. The importance of fatty acid synthesis in vascular regeneration has been confirmed in previous studies[45]. Some research indicates that fatty acid synthesis provides essential biomolecular components for endothelial cells and directly impacts their proliferation[46], a pivotal step in angiogenesis. Furthermore, fatty acid synthesis affects the function and survival of endothelial cells, maintains vascular integrity, and promotes repair after damage[47]. In some studies, fatty acids have been found to act as signaling molecules, regulating intracellular signaling pathways and influencing cell behavior, including the promotion of angiogenic factor release[48, 49]. Therefore, fatty acid synthesis may be one of the crucial mechanisms for endothelial cells to respond to injury and stimulate vascular regeneration.

A significant finding of this study is that after MSCs transfer mitochondria to MPMECs in ARDS, the TCA cycle is stimulated. Moreover, we observed that inhibition of the TCA cycle led to a decrease in the fatty acid synthesis capability of MPMECs, which could be partially restored by the addition of citrate. These results highlight the importance of the TCA cycle and citrate in cellular metabolism, especially in fatty acid synthesis. The TCA cycle is a central pathway in cellular metabolism, playing a critical role not only in energy production but also in providing precursors for many biosynthetic processes, including key metabolic intermediates like citrate for fatty acid synthesis[50]. Previous studies have confirmed the importance of TCA cycle activity in maintaining the anabolic metabolism of cells, particularly in rapidly proliferating cells[51]. As a crucial biosynthetic process, fatty acid synthesis requires the raw materials and energy provided by the TCA cycle. Additionally, citrate, as an essential component of the TCA cycle, is not only a key intermediate in energy metabolism but also a vital precursor for fatty acid synthesis. Citrate can leave the mitochondria and enter the cytoplasm, where it is cleaved by citrate lyase into acetyl-CoA, a direct substrate for fatty acid synthesis. Therefore, when the TCA cycle is inhibited, leading to a reduced supply of citrate and other intermediates, fatty acid synthesis is impacted. This connection between the TCA cycle and fatty acid synthesis has also been supported by previous studies, demonstrating the close link between these two processes[48].

Despite significant progress made in elucidating the impact of MSC-mediated mitochondrial transfer on MPMEC function, this study has several limitations. Firstly, it primarily relies on an ARDS mouse model, which may limit the direct applicability of the findings to humans. While mouse models are valuable tools for studying lung injury and repair mechanisms, they differ from humans in terms of physiology and immune responses. Secondly, while we have observed the influence of the TCA cycle and citrate on fatty acid synthesis in MPMECs, this study does not delve into the specific molecular mechanisms of these metabolic pathways. The precise signaling pathways or molecules that mediate the interaction between the TCA cycle and fatty acid synthesis remain unidentified. Moreover, although the addition of citrate partially restored fatty acid synthesis in our experiments, the effectiveness of this remedial measure in more complex biological systems requires further validation. Thirdly, the observation of MPMECs accepting healthy mitochondria from MSCs through dynamin-dependent clathrin-mediated endocytosis mainly relies on the use of the endocytosis inhibitor Dynasore. While the inhibition of endocytosis with Dynasore allows for indirect inference of mitochondrial transfer mechanisms, it lacks direct visual evidence to confirm the endocytic process. Ideally, direct imaging techniques such as transmission electron microscopy (TEM) to observe mitochondrial translocation and internalization within cells would provide more direct and conclusive evidence. Hence, our conclusions may require further validation through subsequent studies using direct imaging technologies. Finally, the focus of this study is primarily on endothelial cells in ARDS lung tissue. How MSC-mediated mitochondrial transfer affects overall lung function, inflammatory responses, and long-term repair of lung injury still requires further research for elucidation.

Based on the findings of this study regarding mitochondrial transfer from MSCs to MPMECs in ARDS, along with its limitations, future research should focus on several key areas: Firstly, an in-depth investigation into the initiating signals of the mitochondrial transfer is needed. This includes elucidating signaling pathways, particularly those that prompt the release of mitochondria from MSCs and the acceptance of mitochondria by MPMECs. Secondly, the study of signals targeting mitochondrial transfer is equally crucial. Specifically, it is important to explore the mechanisms that dictate the targeting of mitochondria within MSCs to specific damaged cells and the recognition of molecular markers during this process. Moreover, direct observation and verification of the mitochondrial transfer process will be a focal point in future research. The use of advanced imaging techniques, such as TEM, to directly observe mitochondrial translocation within cells, along with the development of in vivo mitochondrial labeling and tracking techniques, will provide key evidence to validate existing hypotheses and understand the dynamics of mitochondrial transfer. Finally, exploring the physiological and pathological implications of mitochondrial transfer will add depth to research in this field. This includes studying the impact of mitochondrial transfer on cellular functions in different disease models and exploring its potential applications in clinical treatment, particularly in regenerative medicine and tissue repair.

In conclusion, this study has made significant advancements in revealing the role of MSC-mediated mitochondrial transfer in repairing the function of MPMECs in ARDS. We discovered that MSCs can transfer mitochondria to MPMECs in ARDS, a process that stimulates the TCA cycle and subsequently promotes fatty acid synthesis. This leads to an increased release of angiogenic factors and enhances vascular regeneration. These findings not only provide new insights into the mechanisms behind MSCs' reparative effects on lung endothelial injury but also offer new strategies for utilizing MSCs in treating mitochondrial damage disease.

ACC: Acetyl-CoA Carboxylase; ACLY: ATP Citrate Lyase; ARDS: Acute Respiratory Distress Syndrome; CS: Citrate Synthase; ELISA: Enzyme-Linked Immunosorbent Assay; FAS: Fatty Acid Synthase; HGF: Hepatocyte Growth Factor IDH: Isocitrate Dehydrogenase; LPS: Lipopolysaccharide; MPMECs: Mouse Pulmonary Microvascular Endothelial Cells; MSCs: Mesenchymal Stem Cells; OGDH: α-Ketoglutarate Dehydrogenase; PCR: Polymerase Chain Reaction; ROS: Reactive Oxygen Species; TCA: Tricarboxylic Acid Cycle; TNTs: Tunneling Nanotubes; VEGF: Vascular Endothelial Growth Factor.

Ethics approval: The Animal Experimental Ethics Committee of Southeast University approved these experiments (approval number: 20200226001).

Consent for publication: Not applicable.

Availability of data and materials: The datasets generated and analyzed during the current study are available from the corresponding author on reasonable request.

Competing interests: The authors declare that there are no conflicts of interest regarding the publication of this paper.

Funding: This study was supported by the National Key R&D Program of China (2022YFC2304600), the National Natural Science Foundation of China (82272235, 82102300, 82272211, 82302470), the Science Foundation of the Commission of Health of Jiangsu Province (ZDB2020009), the Jiangsu Province Key research and development Program (Social Development) Special Project (BE2021734), the China Postdoctoral Science Foundation (2022M710685), and the Special fund project for health science and technology development of Nanjing Municipal Health Commission (YKK21265).

Authors' contributions:

Jinlong Wang: Conception and design, Provision of study material, Collection and assembly of data, Data analysis and interpretation, Manuscript writing, Final approval of manuscript

Shanshan Meng: Conception and design, Financial support, Administrative support, Data analysis and interpretation, Manuscript writing, Final approval of manuscript

Yixuan Chen: Provision of study material, Collection and assembly of data, Data analysis and interpretation, Final approval of manuscript

Haofei Wang: Provision of study material, Collection and assembly of data, Data analysis and interpretation, Final approval of manuscript

Wenhan Hu: Provision of study material, Collection and assembly of data, Data analysis and interpretation, Final approval of manuscript

Shuai Liu: Provision of study material, Collection and assembly of data, Data analysis and interpretation, Final approval of manuscript

Lili Huang: Financial support, Administrative support, Data analysis and interpretation, Final approval of manuscript

Jingyuan Xu: Financial support, Administrative support, Data analysis and interpretation, Final approval of manuscript

Qing Li: Financial support, Administrative support, Data analysis and interpretation, Final approval of manuscript

Xiaojing Wu: Conception and design, Administrative support, Provision of study material, Final approval of manuscript

Wei Huang: Conception and design, Administrative support, Data analysis and interpretation, Manuscript writing, Final approval of manuscript

Yingzi Huang: Conception and design, Financial support, Administrative support, Provision of study material, Data analysis and interpretation, Manuscript writing, Final approval of manuscript

Acknowledgments: In expressing gratitude for the support and guidance received throughout the course of this research, I foremost extend my sincere appreciation to laboratory directors Haibo Qiu and Yi Yang, whose expertise and insightful feedback have been invaluable. I am also grateful to the staff and colleagues within Jiangsu Provincial Key Laboratory of Critical Care Medicine for providing essential resources and an encouraging research environment. Finally, I wish to thank my family and friends for their unwavering encouragement and understanding. This thesis reflects the collective support and dedication of all who were involved.

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MSC-Mediated Mitochondrial Transfer Promotes Metabolic Reprogramming in Endothelial Cells and Vascular Regeneration in ARDS

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Abstract

Background

Methods

Results

Conclusion

Figures

Background

Methods

Establishment of Animal and Cell Models

Husbandry of C57BL/6J Mice

Establishment of the ARDS Mouse Model

Intravenous Injection of MSCs in Mice

Culture, Passaging, and Cryopreservation of MSCs

Culture, Passaging, and Cryopreservation of MPMECs

Mitochondrial Damage Induction with Rotenone

Method Details

Flow Cytometry Analysis of Mitochondrial Transfer from MSCs to MPMECs

Flow Cytometry Analysis of Mitochondrial Transfer from MSCs to MPMECs in ARDS Mouse Model

Confocal Microscopy Observation of Mitochondrial Transfer and Tunneling Nanotubes Between MSCs and MPMECs

Fluorescence Microscopy Observation of Mitochondrial Transfer from MSCs to MPMECs in Isolated Culture

Preparation of Single-Cell Suspension from Mouse Lung Tissue

Western Blot Analysis of Mitochondrial Complex I Expression

Flow Cytometry Analysis of CFSE Mean Fluorescence Intensity in MPMECs

Flow Cytometry Analysis of ROS Mean Fluorescence Intensity in MPMECs

Flow Cytometry Analysis of Cell Apoptosis in MPMECs

Hematoxylin and Eosin (H&E) Staining and Lung Injury Scoring of Mouse Lung Tissue

Quantitative Real-Time PCR Analysis

Enzyme-Linked Immunosorbent Assay

Immunohistochemistry and HALO Quantitative Analysis

ATP Quantitative Analysis in MPMECs

Immunofluorescence Analysis of FAS Protein in MPMECs

Mouse Lung Wet-to-Dry Weight Ratio

Evans Blue Permeability Assay in Mouse Pulmonary Microvasculature

Statistical Analysis

Results

Alleviation of Endothelial Injury in ARDS by MSCs through Mitochondrial Transfer

MSCs Promote Endothelial Cell Proliferation and Vascular Regeneration in ARDS Lung Tissue via Mitochondrial Transfer

MSCs Transfer Mitochondria to MPMECs via TNTs

MPMECs Uptake Mitochondria from MSCs via Dynamin-Dependent Clathrin-Mediated Endocytosis

Mitochondrial Transfer from MSCs to MPMECs Enhances Fatty Acid Synthesis, Facilitating Vascular Regeneration in ARDS

Stimulation of the TCA Cycle by MSC-Transferred Mitochondria Activates Citrate-Dependent Fatty Acid Synthesis in MPMECs

Citrate Restores the Angiogenic Potential of Mitochondrially Impaired MSCs

MSCs Alleviate LPS-Induced Lung Injury

Discussion

Conclusion

Abbreviations

Declarations

References

Supplementary Files

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