Skip to main content
NCBI home page
Physiological Reports logoLink to Physiological Reports
. 2026 Mar 8;14(5):e70810. doi: 10.14814/phy2.70810

Intervention effect of small extracellular vesicles derived from dental pulp stem cells on a high‐altitude pulmonary edema model in male rats

Xue Li 1,2, Zhuang Mao 2, Changyao Wang 2, Yang Liu 2, Youwei Jiang 2, Jiawei Liu 3, XinLong Yan 1,✉, Hua Wang 2,✉
PMCID: PMC12967463  PMID: 41795851

Abstract

High‐altitude pulmonary edema (HAPE) is a life‐threatening disorder caused by hypobaric hypoxia and characterized by pulmonary injury, oxidative stress, and inflammation. We investigated the effects of small extracellular vesicles derived from dental pulp stem cells (DPSCs‐sEVs) in a rat model of HAPE as well as hypoxia‐injured pulmonary microvascular endothelial cells (PMVECs). Rats were exposed to hypobaric hypoxia for 96 h. Lung injury was assessed by histology and immunofluorescence (VEGF, TNF‐α, Occludin). Pulmonary permeability was evaluated by total protein in bronchoalveolar lavage fluid and lung homogenates and by Na+/K+‐ATPase activity. Oxidative stress, inflammatory mediators, and vasoactive factors (NO, PGI₂) were measured. DPSCs‐sEVs attenuated hypoxia‐induced lung injury, increased VEGF and Occludin, reduced TNF‐α, decreased protein leakage, and enhanced Na+/K+‐ATPase activity. DPSCs‐sEVs alleviated oxidative stress and upregulated Nrf2, HO‐1, and GPX1. In vivo, dexamethasone served as a reference treatment; DPSCs‐sEVs produced greater improvements in most endpoints, with comparable effects in selected measures. In PMVECs, DPSCs‐sEVs dose‐dependently mitigated hypoxia‐induced dysfunction. These findings suggest DPSCs‐sEVs protect against hypoxia‐induced pulmonary injury by preserving barrier integrity and improving redox and inflammatory homeostasis.

Keywords: dental pulp stem cells, high‐altitude pulmonary edema, small extracellular vesicles

1. INTRODUCTION

High‐altitude pulmonary edema (HAPE) is a severe form of non‐cardiogenic pulmonary edema induced by hypoxic stress at high altitude, which carries significant mortality risks (Ware & Matthay, 2005). The primary pathophysiological mechanisms involve a pronounced increase in pulmonary arterial pressure, elevated pulmonary vascular permeability, impaired alveolar fluid clearance, and dysregulated fluid transport (Scherrer et al., 2010; Swenson & Bärtsch, 2012). Additionally, hypoxic pulmonary vasoconstriction and elevated circulatory resistance further compromise the lung's ability to clear excess fluid, thereby exacerbating tissue damage (Bärtsch & Swenson, 2013).

Mesenchymal stem cells (MSCs) are adult stem cells with self‐renewal and immune‐regulatory properties. They have been demonstrated to repair damaged lung tissues through multiple mechanisms, such as reducing the permeability of alveolar epithelial and endothelial cells, enhancing alveolar fluid clearance, and improving the phagocytic activity of macrophages (Kim et al., 2019).

Dental pulp stem cells (hDPSCs), a class of MSCs residing in dental pulp tissue, exhibit robust self‐renewal and multi‐lineage differentiation capabilities (Yamada et al., 2019). Although initially employed primarily in oral and maxillofacial surgery (Ledesma‐Martínez et al., 2016), subsequent research has revealed their potent immunomodulatory, neuroregenerative, anti‐inflammatory, and antioxidant properties (Li et al., 2021). This expanded understanding has broadened their therapeutic relevance to a spectrum of conditions, such as spinal cord injuries, Parkinson's disease, Alzheimer's disease, myocardial infarction, diabetes, liver disorders, immune diseases, and oral pathologies (Andrukhov et al., 2019). Our previous study demonstrated that DPSCs can prevent and treat HAPE, primarily through antioxidant effects (including scavenging reactive oxygen species (ROS), activating the Nrf2/HO‐1 pathway, and protecting alveolar epithelial cells), anti‐inflammatory actions, and improving pulmonary vascular permeability. Notably, high‐dose DPSCs were associated with superior therapeutic outcomes (Mao et al., 2024).

Accumulating evidence indicates that the therapeutic benefits of MSCs are primarily mediated through paracrine signaling, involving the secretion of cytokines and extracellular vesicles (EVs) that play pivotal roles in tissue repair and immune modulation (Hu et al., 2020; Qin & Zhao, 2020; Zhao et al., 2021). Among these, small extracellular vesicles (sEVs), typically defined as EVs with a diameter less than 200 nm, represent the most extensively studied EV subtype. They have been implicated in a wide array of physiological and pathological processes, including adipose metabolism, angiogenesis, inflammation, tissue regeneration, neural regeneration, and immune regulation (Jia et al., 2022; Shan et al., 2019). Notably, our recent study demonstrated that DPSCs‐sEVs overexpressing miR‐486 can alleviate HAPE by modulating the PTEN/PI3K/AKT/eNOS pathway, thereby reducing pulmonary vascular permeability and oxidative stress (Wang et al., 2025).

In this study, we established a high‐altitude environmental simulation chamber to establish a rat model of HAPE and further evaluate the preventive and therapeutic potential of DPSCs‐sEVs. Furthermore, to elucidate the underlying endothelial protective mechanisms, we conducted in vitro experiments under hypoxic conditions. Our results demonstrate that DPSCs‐sEVs effectively mitigate HAPE, exhibiting superior efficacy compared to dexamethasone. These findings suggest that DPSCs‐sEVs are a promising class of nanotherapeutic agents for treating HAPE.

2. MATERIALS AND METHODS

2.1. Cell culture

Dental pulp stem cells (DPSCs) of human origin were generously provided by Beijing SH Biotechnology (Beijing, China) and cultured in AM‐V serum‐free medium formulated for mesenchymal stem cells (TBD Biotechnology, Tianjin, China). Donor sex was not specified by the provider.

Pulmonary microvascular endothelial cells (PMVECs) derived from rat lung tissue were purchased from BLUEFBIO (Shanghai, China; BFN60808794) and cultured in Dulbecco's Modified Eagle's Medium (DMEM; Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco, USA). Donor sex was not specified by the supplier. All cells were maintained at 37°C in a humidified atmosphere with 5% CO2.

2.2. Isolation and purification of sEVs

Conditioned medium from DPSCs cultured for 48 h was collected, and sEVs were isolated and purified by differential ultracentrifugation. The medium was sequentially centrifuged at 300×g for 10 min, 2000×g for 20 min, and 10,000×g for 30 min to remove dead cells, apoptotic bodies, and cellular debris. The supernatant was then ultracentrifuged twice at 100,000×g for 70 min each using an ultracentrifuge (XPN‐80, Beckman Coulter, USA). The sEVs pellets were resuspended in phosphate‐buffered saline (PBS) and either stored at −80°C or used immediately for subsequent experiments.

2.3. Identification of sEVs

The concentration, size distribution, polydispersity index (PDI) and zeta potential of sEVs were determined by nanoparticle tracking analysis (NTA) using the ZetaView PMX 110 system (Particle Metrix, Meerbusch, Germany). Morphology was observed under a transmission electron microscope (TEM) (HT7800, Hitachi, Japan). Additionally, Western blotting was performed to detect representative sEVs protein markers.

2.4. Rat HAPE model and treatments

Six‐week‐old male Sprague–Dawley (SD) rats (180–200 g) were obtained from Beijing Vital River Laboratory Animal Technology Co., Ltd. All animals were housed under specific pathogen‐free conditions at 22 ± 2°C with 50%–60% relative humidity and a 12 h light/dark cycle. Rats were fed standard rodent chow (Beijing Vital River Laboratory Animal Technology Co., Ltd., Beijing, China; 02006) and had free access to water. After 1 week of acclimatization, all animals, except those in the control group, were subjected to acute hypobaric hypoxia in a simulated high‐altitude chamber set at an equivalent altitude of 6000 meters. Chamber pressure was gradually reduced from ambient pressure to 46 kPa over approximately 30 min to simulate ascent. Animals were maintained at this simulated altitude for 96 h under normoxic air conditions (19% oxygen). The experimental protocol was reviewed and approved by the Ethics Committee of the Laboratory Animal Center, Academy of Military Medical Sciences (IACUC‐DWZX‐2022‐841). Animals were identified using cage‐based labeling and experimental records without additional invasive marking procedures.

To evaluate the therapeutic effects of DPSCs‐sEVs on HAPE, rats were randomly assigned to six groups (n = 6 per group): (1) The control group (Control), with 100 μL of PBS injected via the tail vein on the 1st and 3rd days; (2) The HAPE group (HAPE), with 100 μL of PBS injected via the tail vein on the 1st and 3rd days of hypoxia treatment; (3) The dexamethasone group (DXMS), with dexamethasone injected intraperitoneally at a dose of 4 mg/kg for four consecutive days, from 1 day before hypoxia treatment to the third day of hypoxia treatment; (4) DPSCs‐sEVs prevention group (sEV‐pre), DPSCs‐sEVs were administered via tail vein injection at a dose of 1 × 1010 particles in 100 μL PBS, on the first and third days prior to hypoxia treatment. (5) Low‐dose DPSCs‐sEVs group (sEV‐L) and (6) High‐dose DPSCs‐sEVs group (sEV‐H), DPSCs‐sEVs were administered via tail vein injection at a dose of 1 × 1010 particles and 2 × 1010 particles respectively in 100 μL PBS, on the 1st and 3rd days of hypoxia treatment.

2.5. Sample collection

At the end of the experiment, rats were anesthetized with pentobarbital sodium, and blood was collected from the inferior vena cava to obtain serum and plasma for subsequent analysis. After thoracotomy, the right lung of three rats was lavaged three times with sterile physiological saline via endotracheal intubation, and bronchoalveolar lavage fluid (BALF) was collected. The left lung was fixed in 4% paraformaldehyde (PFA). The left lungs of the remaining three rats were homogenized to prepare tissue extracts for the determination of total protein content, Na+/K+‐ATPase activity, malondialdehyde (MDA), superoxide dismutase (SOD), and the expression of IL‐1β and IL‐6. The right lungs were used for RNA extraction and subsequent analysis of TNF‐α, Nrf2, GPX1, and HO‐1 expression.

2.6. H&E staining and immunofluorescence

The PFA‐fixed left lungs were paraffin‐embedded and sectioned. For histopathological evaluation, sections were deparaffinized, rehydrated, and stained with hematoxylin and eosin (H&E). Alveolar area and inflammatory cell counts were quantified from the H&E‐stained sections using ImageJ software.

For immunofluorescence, sections were stained with primary antibodies against VEGF (Santa Cruz Biotechnology, TX, USA; SC‐7269), TNF‐α (Proteintech, Wuhan, China; 17590‐1‐AP), and Occludin (Proteintech, Wuhan, China; 27260‐1‐AP) to assess their protein expression and localization. Nuclei were counterstained with DAPI (BDHM, Beijing, China; BD‐DAPI‐1). Positive staining was quantified using ImageJ, and results are expressed as the percentage of positive cells.

For histological quantification, five randomly selected fields per section and three sections per lung were analyzed. The mean value for each animal was calculated and used for statistical analysis, with the animal serving as the biological unit (n = 3 per group).

2.7. Assessment of pulmonary permeability in HAPE rats

Total protein content in BALF and lung tissue homogenates was measured using the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, Illinois, USA; 23225). The activity of Na+/K+‐ATPase in lung tissue homogenates was determined using a commercial kit (Elabscience, Wuhan, China; E‐BC‐K539‐M), following the manufacturer's instructions.

2.8. Quantification of oxidative stress level

MDA levels and SOD activity in lung tissue homogenates were determined with commercial colorimetric kits (Elabscience, Wuhan, China; MDA kit: E‐BC‐K025‐M, SOD kit: E‐BC‐K020‐M). The expression of Nrf2, glutathione peroxidase 1 (GPX1), and HO‐1 in lung tissues was analyzed by quantitative real‐time PCR (qPCR), serving as indices of oxidative stress.

2.9. Expression levels of inflammatory cytokines

The levels of inflammatory cytokines IL‐1β and IL‐6 (ABclonal, Wuhan, China; IL‐1β kit: RK00009, IL‐6 kit: RK00020) in lung tissue homogenates, and interferon‐γ (IFN‐γ), IL‐1β and TNF‐⍺ (MEIKE, Jiangsu, China; IFN‐γ kit: MK1739A, IL‐1β kit: MK1588A, TNF‐⍺ kit: MK1721A) in serum were measured using ELISA kits according to the manufacturer's instructions. In addition, TNF‐⍺ expression in lung tissues was analyzed by qPCR.

2.10. Evaluation of vascular activity

Plasma levels of vasoactive substances nitric oxide (NO) and prostaglandin I2 (PGI2) were determined. NO was assessed using a colorimetric assay kit (Elabscience, Wuhan, China; E‐BC‐K035‐M), and PGI2 was quantified by ELISA (Elabscience, Wuhan, China; E‐EL‐0022c).

All biochemical and enzymatic activity assays were performed in triplicate for each lung sample, and the mean value was used for statistical analysis.

2.11. In vitro cell hypoxia model and treatment

To evaluate the effects of DPSCs‐sEVs on endothelial cells hypoxic injury, PMVECs were exposed to 200 μM cobalt chloride (CoCl2, Sigma, USA; C8661) to establish hypoxic injury model. CoCl2 chemically induces hypoxic responses by stabilizing hypoxia‐inducible factor‐1α (HIF‐1α) through inhibition of prolyl hydroxylases, thereby mimicking cellular hypoxia in vitro. The cells were divided into five experimental groups: (1) The control group (Control), consisted of untreated cells; (2) The Hypoxia group (Hypoxia), PMVECs was treated with CoCl2 alone for 24 h; (3) sEV‐prevention group (sEV‐Pre), PMVECs was treated with CoCl2 and DPSCs‐sEVs (5 × 109 particles) simultaneously for 24 h; (4) The Low‐dose sEV group (sEV‐L), PMVECs was exposed to CoCl2 for 12 h, then treated with 5 × 109 particles of DPSCs‐sEVs; (5) The High‐dose sEV group (sEV‐H), PMVECs was received CoCl2 for 12 h before treated with 1 × 1010 particles of DPSCs‐sEVs. Then PMVECs were collected to evaluate the effects of DPSCs‐sEVs on endothelial function. Cell permeability was assessed by quantifying the expression of aquaporins AQP‐1 and AQP‐5 using qPCR. Inflammatory responses were evaluated by detecting IL‐1β and IL‐6 expression. Vascular function was assessed by quantifying endothelial nitric oxide synthase (eNOS) and endothelin‐1 (ET‐1) expression. Oxidative stress was analyzed through the expression of nuclear factor erythroid 2‐related factor 2 (Nrf2) and heme oxygenase‐1 (HO‐1), measured using both qPCR and Western blotting.

2.12. RNA isolation, reverse transcription, and qPCR

Total RNA was extracted from lung tissues and PMVECs using the RNA‐Quick Purification Kit (ES Science, Beijing, China; RN001). cDNA was synthesized using the Evo M‐MLV RT Mix Kit (Accurate, Hunan, China; AG11728). Quantitative PCR (qPCR) was performed using the SYBR Green I Master kit (Yeasen, Shanghai, China; 11201ES08) on a CFX Connect Real‐Time System (Bio‐Rad, CA, USA). Primers sequences were as follows: β‐actin, sense 5′‐cacccgcgagtacaaccttc‐3′ and antisense 5′‐cccatacccaccatcacacc‐3′; TNF‐α, sense 5′‐gttggaccaattcataggcgc‐3′ and antisense 5′‐caatgtcgatcacatgcacca‐3′; Nrf2, sense 5′‐gatgatgccagccagctgaa‐3′ and antisense 5′‐gcgactgactaatggcagcaga‐3′; GPX1, sense 5′‐catcacgagaatggcaagaa‐3′ and antisense 5′‐tcacctcgcacttctcaaac‐3′; HO‐1, sense 5′‐ggcctgctagcctggttcaa‐3′ and antisense 5′‐ggtgtctgggatgaactagtgctg‐3′; IL‐1β, sense 5′‐gaagtcaagaccaaagtgg‐3′ and antisense 5′‐tgaagtcaactatgtcccg‐3′; AQP5, sense 5′‐ggccacatcaatccagccatt‐3′ and antisense 5′‐aaagatcgggctgggttgat‐3′; AQP1, sense 5′‐ctggcctttggtttgagcat‐3′ and antisense 5′‐ccacacactgggcgatgat‐3′; eNOS, sense 5′‐tactccaggctcccgatg‐3′ and antisense 5′‐aagggcagcaaaccactc‐3′; ET1, sense 5′‐cgttgctcctgctcctcc‐3′ and antisense 5′‐ggtctgtggtctttgtggg‐3′. Relative mRNA expression levels were calculated using β‐actin as the reference gene.

2.13. Western blot

Total protein was extracted using RIPA lysis buffer (Beyotime, Shanghai, China; P0013B) supplemented with protease (Epizyme, Shanghai, China, GRF101) and phosphatase inhibitors (Epizyme, Shanghai, China, GRF102). Proteins were separated by 10% SDS–PAGE and transferred onto PVDF membranes. After blocking with 5% non‐fat milk for 1 h at room temperature, the membranes were incubated with the appropriate primary antibodies overnight at 4°C, followed by incubation with HRP‐conjugated secondary antibodies for 1 h at room temperature. Protein bands were detected using an enhanced chemiluminescence reagent (ECL; Zomanbio, Beijing, China; ZD301). The antibodies and their dilution ratios are summarized in Table 1.

TABLE 1.

Primary and secondary antibodies used in this study.

Product Cat. NO. Dilution ratio Company
Anti‐beta III Tubulin ab18207 1:10000 Abcam, Cambridge, UK
Anti‐TSG101 ab125011 1:1000 Abcam, Cambridge, UK
Anti‐CD9 ab236630 1:1000 Abcam, Cambridge, UK
Thy1/CD90 Rabbit mAb 13801S 1:1000 CST, MA, USA
Anti‐Heme Oxygenase 1 Ab68477 1:1000 Abcam, Cambridge, UK
NRF2 (E3J1V) Rabbit mAb 33649 1:1000 CST, MA, USA
Beta Actin Rabbit pAb 380624 1:10000 ZENBIO, Chengdu, China
Goat Anti‐rabbit IgG (H + L) BST18D06A8E54 1:5000 Boster, Wuhan China
Goat anti‐mouse IgG (H + L), Alexa Fluor 488 ab150113 1:500 Abcam, Cambridge, UK
Goat anti‐rabbit IgG (H + L), Alexa Fluor 488 ab150077 1:500 Abcam, Cambridge, UK
Open in a new tab

2.14. Statistical analysis

Data are presented as mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism 9. Differences among multiple groups were assessed by one‐way ANOVA followed by Tukey's multiple comparisons test. p‐value <0.05 was considered statistically significant.

3. RESULTS

3.1. Identification of DPSCs‐sEVs

DPSCs‐sEVs were isolated and purified via differential centrifugation and characterized by nanoparticle tracking analysis (NTA), transmission electron microscopy (TEM), and Western blot. NTA revealed that DPSCs‐sEVs had an average diameter of 125.8 nm (Figure 1a), a zeta potential of −30.73 ± 0.46 mV at room temperature (Figure 1b), a concentration of 1.8 × 1011 particles/mL, and a polydispersity index (PDI) of 0.231. TEM confirmed the typical bilayer membrane structures with diameters of approximately 110 nm (Figure 1c), consistent with NTA measurements. Western blot analysis demonstrated expression of exosomal markers TSG101 and CD9, as well as the stem cell marker CD90 (Figure 1d).

FIGURE 1.

FIGURE 1

Open in a new tab

Characterization of DPSCs‐sEVs. Size distribution profile (a) and Zeta Potential (b) of DPSCs‐sEVs were detected by NTA. (c) TEM was used to examine the morphology of DPSCs‐sEVs (Scale bar = 100 nm). (d) Western blot analysis for sEVs marker proteins TSG101 and CD9, and DPSCs marker CD90.

3.2. DPSCs‐sEVs attenuate lung injury in a rat HAPE model

To evaluate the therapeutic potential of DPSCs‐sEVs against HAPE, we established a rat model in a simulated hypobaric hypoxic environment equivalent to 6000 meters above sea level. DPSCs‐sEVs were administered intravenously either prior to hypobaric exposure (−72 and −24 h before altitude simulation) or during hypobaric exposure (24 and 72 h after initiation of altitude simulation), using dexamethasone (DXMS) as a positive control. The experimental grouping and protocol are illustrated in Figure 2a. Histopathological analysis (H&E staining) of lung tissue from HAPE rats revealed characteristic injuries, including thickened alveolar walls, inflammatory cell infiltration, and alveolar edema accompanied by erythrocyte exudation. These pathological alterations were substantially ameliorated by treatment with either DXMS or DPSCs‐sEVs (Figure 2b). Quantitative analysis of alveolar area and inflammatory cells infiltration verified these improvements (Figure 2c). Notably, the high‐dose DPSCs‐sEVs group (sEV‐H) exhibited more pronounced histological improvement than the low‐dose group (sEV‐L), indicating a clear dose‐dependent protective effect.

FIGURE 2.

FIGURE 2

Open in a new tab

Histological analysis of the lung tissue in rat HAPE model after DPSCs‐sEVs treatment. (a) Schematic description of the establishment and treatment of the HAPE model (by Figdraw). (b) Representative images of H&E staining (Scale bar = 50 μm). The red arrow represents the alveolar septa, yellow arrow marks the alveolar walls, and black arrow points to inflammatory cells. (c) Quantitative analysis of the percentage of alveolar area and inflammatory cell infiltration. Data are presented as mean ± SD, n = 3. #, vs. control group; *, vs. HAPE group; @, vs. DXMS group; $, vs. sEV‐pre group; ¥, vs. sEV‐L group.

3.3. DPSCs‐sEVs restore pulmonary barrier function

To investigate the mechanisms underlying DPSCs‐sEVs mediated protection, we first assessed pulmonary barrier integrity. In HAPE rats, total protein concentrations in bronchoalveolar lavage fluid (BALF) and lung homogenates were significantly elevated (Figure 3a,b), indicative of alveolar‐capillary barrier disruption. This leakage was markedly attenuated by treatment with either DXMS or DPSCs‐sEVs. DPSCs‐sEVs administration also significantly enhanced Na+/K+‐ATPase activity in lung tissues (Figure 3c), suggesting a promotion of active alveolar fluid clearance. Immunofluorescence analysis corroborated these findings, showing that the hypoxia‐induced reduction in Occludin‐positive cells was partially reversed by DPSCs‐sEVs in a dose‐dependent manner, with the high‐dose group (sEV‐H) exhibiting the most pronounced effect (Figure 3d). In vitro, hypoxia downregulated the expression of the water channel proteins aquaporin‐1 (AQP1) and aquaporin‐5 (AQP5) in pulmonary microvascular endothelial cells (PMVECs), and this effect was dose‐dependently rescued by DPSCs‐sEVs treatment (Figure 3e,f). Collectively, these data indicate that DPSCs‐sEVs preserve alveolar‐capillary barrier integrity and promote alveolar fluid clearance under hypobaric hypoxia.

FIGURE 3.

FIGURE 3

Open in a new tab

DPSCs‐sEVs restore pulmonary permeability in vivo and in vitro. Total protein content in BALF (a) and lung tissue homogenates (b). (c) Specific activity of Na+/K+‐ATPase in lung tissues. (d) Representative immunofluorescence staining of Occludin in lung tissues (scale bar = 25 μm) and quantification of Occludin‐positive cells (percentage of total cells). The mRNA expression of AQP1 (e) and AQP5 (f) in PMVECs was determined by qPCR. Data are presented as mean ± SD, n = 3. #, vs. control group; *, vs. HAPE group; @, vs. DXMS group; $, vs. sEV‐pre group; ¥, vs. sEV‐L group.

3.4. DPSCs‐sEVs reduce oxidative stress

Hypobaric hypoxia induced profound oxidative stress in lung tissues, as evidenced by a significant increase in malondialdehyde (MDA) and a concomitant decrease in superoxide dismutase (SOD) activity (Figure 4a,b). Both DXMS and DPSCs‐sEVs treatments mitigated lipid peroxidation (reduced MDA) and restored antioxidant capacity (increased SOD). At the molecular level, the expression of key antioxidant regulators—Nrf2 and its downstream effectors HO‐1 and GPX1—was significantly downregulated in HAPE rats. DPSCs‐sEVs treatment dose‐dependently restored their expression (Figure 4c–e). Consistently, in hypoxic PMVECs, the suppression of Nrf2 and HO‐1 expression was effectively reversed by DPSCs‐sEVs in a dose‐dependent manner, as confirmed by both qPCR and western blot analyses (Figure 4f–h). These findings suggest activation of the Nrf2 axis as a key contributor to the antioxidant effects of DPSCs‐sEVs.

FIGURE 4.

FIGURE 4

Open in a new tab

DPSCs‐sEVs mitigate oxidative stress in vivo and in vitro. The concentration of MDA (a) and the activity of SOD (b) in lung tissue homogenate. Expression levels of Nrf2 (c), HO‐1 (d), and GPX1 (e) in lung tissues. The mRNA expression levels of Nrf2 (f) and HO‐1 (g) in PMVECs determined by qPCR. (h) Representative Western blot showing protein expression levels of Nrf2 and HO‐1 in PMVECs. (i) Gray values analysis of Figure H. Data are presented as mean ± SD, n = 3. #, vs. control group; *, vs. HAPE group; @, vs. DXMS group; $, vs. sEV‐pre group; ¥, vs. sEV‐L group. Full‐length blot images with labeled molecular weight markers are provided in the Supplementary Materials.

3.5. DPSCs‐sEVs attenuate inflammatory responses

We next evaluated the impact of DPSCs‐sEVs on the hypobaric hypoxia‐triggered inflammatory cascade. HAPE rats exhibited elevated levels of pro‐inflammatory cytokines, including IL‐1β and IL‐6 in lung homogenates, IL‐1β, IFN‐γ, and TNF‐α in serum, and TNF‐α in lung tissues (Figure 5a–c). Treatment with DXMS or DPSCs‐sEVs significantly reduced these cytokine elevations. Notably, high‐dose DPSCs‐sEVs (sEV‐H) demonstrated a superior effect in reducing lung IL‐6 and tissue TNF‐α levels compared to DXMS (Figure 5a,c). This was consistent with immunofluorescence staining, which revealed an increased percentage of TNF‐α–positive cells in HAPE lungs that was most evidently reduced in the sEV‐H group (Figure 5d). In vitro, the hypoxia‐induced upregulation of IL‐1β and TNF‐α in PMVECs was dose‐dependently suppressed by DPSCs‐sEVs (Figure 5e,f).

FIGURE 5.

FIGURE 5

Open in a new tab

Effects of DPSCs‐sEVs on inflammatory cytokine expression in vivo and in vitro. (a) Concentrations of IL‐1β and IL‐6 in lung tissue homogenates. (b) Concentrations of IL‐1β, IFN‐γ and TNF‐⍺ in serum. (c) The mRNA expression level of TNF‐α in lung tissues. (d) Representative immunofluorescence staining of TNF‐α in lung tissues (scale bar =25 μm) and quantification of TNF‐α–positive cells (percentage of total cells). The mRNA expression levels of IL‐1β (e) and TNF‐⍺ (f) in PMVECs. Data are presented as mean ± SD, n = 3. #, vs. control group; *, vs. HAPE group; @, vs. DXMS group; $, vs. sEV‐pre group; ¥, vs. sEV‐L group.

3.6. DPSCs‐sEVs improve vascular reactivity

Acute hypobaric hypoxia in the HAPE model significantly impaired vascular function, as evidenced by marked reductions in plasma levels of the vasodilators nitric oxide (NO) and prostacyclin (PGI2) compared with normoxic controls. Treatment with either DXMS or DPSCs‐sEVs effectively reversed these reductions, with high‐dose DPSCs‐sEVs (sEV‐H) restoring NO and PGI2 levels to an extent comparable to, or greater than, DXMS (Figure 6a,b). To explore the underlying cellular mechanisms, we examined key regulators of vascular tone in PMVECs. Hypoxia downregulated endothelial nitric oxide synthase (eNOS) expression while upregulating endothelin‐1 (ET‐1), and DPSCs‐sEVs dose‐dependently reversed these alterations by increasing eNOS and decreasing ET‐1 (Figure 6c,d). In parallel, DPSCs‐sEVs increased VEGF‐positive signals in lung microvessels, consistent with enhanced endothelial repair and angiogenic responses under hypobaric hypoxia (Figure 6e).

FIGURE 6.

FIGURE 6

Open in a new tab

Effects of DPSCs‐sEVs on vasoactive substances in vivo and in vitro. The expression levels of NO (a) and PGI2 (b) in plasma. eNOS (c) and ET‐1 (d) mRNA expression in PMVECs determined by qPCR. (e) Representative immunofluorescence staining of VEGF in lung tissues (scale bar = 25 μm) and quantification of VEGF–positive cells (percentage of total cells). Data are presented as mean ± SD, n = 3. #, vs. control group; *, vs. HAPE group; @, vs. DXMS group; $, vs. sEV‐pre group; ¥, vs. sEV‐L group.

4. DISCUSSION

High‐altitude pulmonary edema (HAPE) is a life‐threatening, non‐cardiogenic pulmonary edema that is primarily driven by exaggerated hypoxic pulmonary vasoconstriction and elevated pulmonary vascular pressures, leading to alveolar–capillary stress failure and fluid leakage (Hultgren, 1996; Tian et al., 2024). Although descent and supplemental oxygen remain the most effective interventions, pharmacological prophylaxis and treatment are often required during rapid ascent (Pennardt, 2013). Dexamethasone (DXMS) is commonly used in clinical practice and has been reported to improve alveolar fluid clearance, reduce protein leakage, and modulate oxidative stress and nitric oxide (NO) bioavailability (Joyce et al., 2018; Kosutova et al., 2021; Maggiorini et al., 2006; Mehta et al., 2008; O'Hara et al., 2014; Suhail, 2010); therefore, it was employed as a positive control in the present study.

In this study, we established a rat HAPE model using simulated hypobaric hypoxia equivalent to 6000 m and observed typical pathological features, including alveolar septal thickening, inflammatory cell infiltration, and edema. We demonstrate that small extracellular vesicles derived from dental pulp stem cells (DPSCs‐sEVs) confer substantial protection against hypobaric hypoxia‐induced lung injury. Importantly, DPSCs‐sEVs improved lung histopathology in a dose‐dependent manner and, in several endpoints, high‐dose sEVs achieved effects comparable to or exceeding DXMS, suggesting that DPSCs‐sEVs may represent a promising cell‐free therapeutic strategy for HAPE.

Disruption of the alveolar–capillary barrier and impaired alveolar fluid clearance are central to HAPE pathogenesis (Paralikar, 2012). Protein leakage into bronchoalveolar lavage fluid (BALF) is a sensitive indicator of increased microvascular permeability, while epithelial sodium transport and Na+/K+‐ATPase activity are critical determinants of active fluid reabsorption (Baloglu et al., 2011; Vivona et al., 2001; Yue & Guidry, 2019). We found that hypobaric hypoxia markedly increased total protein levels in BALF and lung homogenates and reduced Na+/K+‐ATPase activity, consistent with severe barrier compromise. DPSCs‐sEVs significantly reduced protein leakage and restored Na+/K+‐ATPase activity, indicating improved barrier integrity and enhanced fluid clearance capacity. Concordantly, DPSCs‐sEVs partially restored Occludin‐positive signals in lung tissues and rescued the hypoxia‐induced downregulation of aquaporin‐1 (AQP1) and aquaporin‐5 (AQP5) in pulmonary microvascular endothelial cells (PMVECs), further supporting a protective role in preserving alveolar–capillary homeostasis under hypobaric hypoxia.

Oxidative stress is increasingly recognized as a key driver of hypoxia‐induced endothelial dysfunction and pulmonary edema (Pena et al., 2022). In this study, hypobaric hypoxia induced pronounced lipid peroxidation, evidenced by elevated malondialdehyde (MDA), together with diminished antioxidant capacity, reflected by reduced superoxide dismutase (SOD) activity. DPSCs‐sEVs mitigated these changes and restored the expression of Nrf2 and its downstream antioxidant effectors (HO‐1 and GPX1) in vivo and in hypoxic PMVECs. These results suggest that activation of the Nrf2 axis may contribute to the antioxidant effects of DPSCs‐sEVs and provide a mechanistic link between redox homeostasis and improved endothelial barrier function.

Inflammation has been variably implicated in HAPE, with evidence supporting both an early contribution and a secondary amplification phase following endothelial injury (El Alam et al., 2022; Hartmann et al., 2000; Ottolenghi et al., 2020; Sarada et al., 2008; Zhou et al., 2017). In the present study, hypobaric hypoxia increased multiple pro‐inflammatory cytokines in lung tissues and serum, and DPSCs‐sEVs significantly suppressed these inflammatory mediators. Notably, high‐dose DPSCs‐sEVs showed a strong inhibitory effect on IL‐6 and TNF‐α, which aligns with the observed improvement in lung pathology and barrier integrity. However, clinical studies have suggested minimal inflammation in the earliest phase of human HAPE, implying that inflammatory activation may occur downstream of mechanical and redox‐mediated endothelial injury (Swenson et al., 2002). The apparent discrepancy between human observations and our rat model may reflect differences in species‐specific immune responses, exposure paradigms, and sampling timing. Future work incorporating time‐course analyses and additional models will be important to clarify whether DPSCs‐sEVs primarily prevent inflammatory initiation or predominantly attenuate secondary inflammatory amplification.

In addition to barrier disruption, dysregulated pulmonary vascular tone is fundamental to HAPE progression (Berger et al., 2009; Ingram et al., 2010; Scherrer et al., 1996). Hypobaric hypoxia reduced circulating vasodilators, including NO and prostacyclin (PGI2), and disrupted endothelial vasoactive balance, characterized by decreased eNOS and increased endothelin‐1 (ET‐1) in PMVECs. DPSCs‐sEVs dose‐dependently restored NO/PGI2 levels and normalized the eNOS/ET‐1 axis, indicating improved endothelial function and vascular reactivity. Furthermore, DPSCs‐sEVs increased VEGF‐positive signals in lung microvessels. Here, VEGF is interpreted as an indicator of endothelial repair/angiogenic signaling rather than a direct surrogate of barrier tightness, and its restoration may reflect enhanced endothelial resilience and microvascular repair capacity under hypoxic stress.

Collectively, our in vivo and in vitro data support a multifaceted protective profile of DPSCs‐sEVs in hypobaric hypoxia‐induced lung injury, integrating preservation of alveolar–capillary integrity, enhancement of fluid clearance, suppression of oxidative stress through Nrf2‐associated pathways, attenuation of inflammatory responses, and restoration of endothelial vasoactive homeostasis. These convergent mechanisms likely act in concert to limit edema formation and improve pulmonary function in HAPE.

This study has several limitations. First, while the association between DPSCs‐sEVs and activation of Nrf2 signaling is robust, causality remains to be established; pharmacological inhibition or genetic modulation of Nrf2/HO‐1 would strengthen mechanistic inference. Second, the bioactive cargo within DPSCs‐sEVs (e.g., miRNAs or proteins) responsible for these effects was not identified and warrants systematic profiling and functional validation. Third, the current work primarily focuses on acute injury; longer‐term outcomes, optimal dosing schedules, and biodistribution of DPSCs‐sEVs should be examined to support translational development. Finally, although PMVECs provide a relevant endothelial model, the alveolar epithelium and immune compartments likely contribute to HAPE pathophysiology. Future studies incorporating specific immune cell phenotyping (e.g., neutrophils, macrophages, and mast cells) using immunohistochemical or flow cytometric approaches, as well as multi‐cellular or organoid models, will be important to further delineate the cellular immune mechanisms underlying the protective effects of DPSCs‐sEVs.

Importantly, accumulating evidence from preclinical and early‐phase clinical studies suggests that mesenchymal stem cell‐derived extracellular vesicles exhibit favorable safety characteristics, including low immunogenicity and minimal risk of tumorigenicity compared with cell‐based therapies (Wu et al., 2025; Zhang et al., 2023). These characteristics may lower the translational barrier relative to cell‐based therapies and support the feasibility of advancing EV‐based strategies toward clinical evaluation.

Overall, DPSCs‐sEVs markedly attenuated hypobaric hypoxia‐induced lung injury in a rat HAPE model and improved vascular reactivity, with dose‐dependent efficacy and favorable performance relative to DXMS in selected endpoints. These findings highlight DPSCs‐sEVs as a potential cell‐free therapeutic candidate for HAPE and provide a mechanistic basis for further translational investigation.

5. CONCLUSIONS

In summary, our findings suggest that DPSCs‐sEVs possess both prophylactic and therapeutic potential against HAPE, providing a foundation for future translational research in high‐altitude medicine.

AUTHOR CONTRIBUTIONS

X.L.: Investigation; methodology; writing—original draft. Z.M.: Investigation; MEthodology. C.W.: Investigation; methodology. Y.L., Y.J., J.L.: Investigation. X.Y.: Conceptualization; supervision; writing—review and editing. H.W.: Conceptualization; investigation; supervision; validation; writing—review and editing. All authors have read and agreed to the published version of the article.

FUNDING INFORMATION

The laboratory and animal experiments reported in this manuscript were financially supported by the National Natural Science Foundation of China (Grant No. 81573086).

CONFLICT OF INTEREST STATEMENT

The authors declare no conflicts of interest.

ETHICS STATEMENT

All animal procedures were approved by the Ethics Committee of the Laboratory Animal Center, Academy of Military Medical Sciences (Approval No. IACUC‐DWZX‐2022‐841) and were conducted in accordance with relevant institutional guidelines for the care and use of laboratory animals.

Supporting information

Data S1.

PHY2-14-e70810-s001.xlsx (18.5KB, xlsx)

Data S2.

PHY2-14-e70810-s002.pdf (1.2MB, pdf)

ACKNOWLEDGMENTS

The authors thank Professor Rili Ge and Feng Tang for their assistance in establishing the HAPE model.

Contributor Information

XinLong Yan, Email: yxlong2000@bjut.edu.cn.

Hua Wang, Email: 18511712135@163.com, Email: wanghua@bmi.ac.cn.

DATA AVAILABILITY STATEMENT

All data generated or analyzed during this study are included in this published article. However, the data that support the findings of this study are available from the corresponding author upon reasonable request.

REFERENCES

  1. Andrukhov, O. , Behm, C. , Blufstein, A. , & Rausch‐Fan, X. (2019). Immunomodulatory properties of dental tissue‐derived mesenchymal stem cells: Implication in disease and tissue regeneration. World Journal of Stem Cells, 11(9), 604–617. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Baloglu, E. , Reingruber, T. , Bärtsch, P. , & Mairbäurl, H. (2011). β2‐Adrenergics in hypoxia desensitize receptors but blunt inhibition of reabsorption in rat lungs. American Journal of Respiratory Cell and Molecular Biology, 45(5), 1059–1068. [DOI] [PubMed] [Google Scholar]
  3. Bärtsch, P. , & Swenson, E. R. (2013). Clinical practice: Acute high‐altitude illnesses. The New England Journal of Medicine, 368(24), 2294–2302. [DOI] [PubMed] [Google Scholar]
  4. Berger, M. M. , Dehnert, C. , Bailey, D. M. , Luks, A. M. , Menold, E. , Castell, C. , Schendler, G. , Faoro, V. , Mairbäurl, H. , Bärtsch, P. , & Swenson, E. R. (2009). Transpulmonary plasma ET‐1 and nitrite differences in high altitude pulmonary hypertension. High Altitude Medicine & Biology, 10(1), 17–24. [DOI] [PubMed] [Google Scholar]
  5. El Alam, S. , Pena, E. , Aguilera, D. , Siques, P. , & Brito, J. (2022). Inflammation in pulmonary hypertension and edema induced by hypobaric hypoxia exposure. International Journal of Molecular Sciences, 23(20), 12656. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Hartmann, G. , Tschöp, M. , Fischer, R. , Bidlingmaier, C. , Riepl, R. , Tschöp, K. , Hautmann, H. , Endres, S. , & Toepfer, M. (2000). High altitude increases circulating interleukin‐6, interleukin‐1 receptor antagonist and C‐reactive protein. Cytokine, 12(3), 246–252. [DOI] [PubMed] [Google Scholar]
  7. Hu, Q. , Su, H. , Li, J. , Lyon, C. , Tang, W. , Wan, M. , & Hu, T. Y. (2020). Clinical applications of exosome membrane proteins. Precision Clinical Medicine, 3(1), 54–66. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Hultgren, H. N. (1996). High‐altitude pulmonary edema: Current concepts. Annual Review of Medicine, 47, 267–284. [DOI] [PubMed] [Google Scholar]
  9. Ingram, T. E. , Pinder, A. G. , Bailey, D. M. , Fraser, A. G. , & James, P. E. (2010). Low‐dose sodium nitrite vasodilates hypoxic human pulmonary vasculature by a means that is not dependent on a simultaneous elevation in plasma nitrite. American Journal of Physiology. Heart and Circulatory Physiology, 298(2), H331–H339. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Jia, Y. , Yu, L. , Ma, T. , Xu, W. , Qian, H. , Sun, Y. , & Shi, H. (2022). Small extracellular vesicles isolation and separation: Current techniques, pending questions and clinical applications. Theranostics, 12(15), 6548–6575. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Joyce, K. E. , Lucas, S. J. E. , Imray, C. H. E. , Balanos, G. M. , & Wright, A. D. (2018). Advances in the available non‐biological pharmacotherapy prevention and treatment of acute mountain sickness and high altitude cerebral and pulmonary oedema. Expert Opinion on Pharmacotherapy, 19(17), 1891–1902. [DOI] [PubMed] [Google Scholar]
  12. Kim, S. Y. , Joglekar, M. V. , Hardikar, A. A. , Phan, T. H. , Khanal, D. , Tharkar, P. , Limantoro, C. , Johnson, J. , Kalionis, B. , & Chrzanowski, W. (2019). Placenta stem/stromal cell‐derived extracellular vesicles for potential use in lung repair. Proteomics, 19(17), e1800166. [DOI] [PubMed] [Google Scholar]
  13. Kosutova, P. , Kolomaznik, M. , Calkovska, A. , Mokra, D. , & Mikolka, P. (2021). Nitric‐oxide‐releasing dexamethasone derivative NCX‐1005 improves lung function and attenuates inflammation in experimental lavage‐induced ARDS. Pharmaceutics, 13(12), 2092. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Ledesma‐Martínez, E. , Mendoza‐Núñez, V. M. , & Santiago‐Osorio, E. (2016). Mesenchymal stem cells derived from dental pulp: A review. Stem Cells International, 2016, 4709572. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Li, B. , Ouchi, T. , Cao, Y. , Zhao, Z. , & Men, Y. (2021). Dental‐derived mesenchymal stem cells: State of the art. Frontiers in Cell and Development Biology, 9, 654559. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Maggiorini, M. , Brunner‐La Rocca, H. P. , Peth, S. , Fischler, M. , Böhm, T. , Bernheim, A. , Kiencke, S. , Bloch, K. E. , Dehnert, C. , Naeije, R. , Lehmann, T. , Bärtsch, P. , & Mairbäurl, H. (2006). Both tadalafil and dexamethasone may reduce the incidence of high‐altitude pulmonary edema: A randomized trial. Annals of Internal Medicine, 145(7), 497–506. [DOI] [PubMed] [Google Scholar]
  17. Mao, Z. , Wang, C. , Liu, J. , Li, X. , Duan, H. , Ye, Y. , Liu, H. , Lv, L. , Xue, G. , He, Z. , Wuren, T. , & Wang, H. (2024). Superoxide dismutase 1‐modified dental pulp stem cells alleviate high‐altitude pulmonary edema by inhibiting oxidative stress through the Nrf2/HO‐1 pathway. Gene Therapy, 31(7–8), 422–433. [DOI] [PubMed] [Google Scholar]
  18. Mehta, S. R. , Chawla, A. , & Kashyap, A. S. (2008). Acute Mountain sickness, high altitude cerebral Oedema, high altitude pulmonary Oedema: The current concepts. Medical Journal, Armed Forces India, 64(2), 149–153. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. O'Hara, R. , Serres, J. , Dodson, W. , Bruce, W. , Ordway, J. , Powell, E. , & Wade, M. (2014). The use of dexamethasone in support of high‐altitude ground operations and physical performance: Review of the literature. Journal of Special Operations Medicine, 14(4), 53–58. [DOI] [PubMed] [Google Scholar]
  20. Ottolenghi, S. , Zulueta, A. , & Caretti, A. (2020). Iron and sphingolipids as common players of (mal)adaptation to hypoxia in pulmonary diseases. International Journal of Molecular Sciences, 21(1), 307. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Paralikar, S. J. (2012). High altitude pulmonary edema‐clinical features, pathophysiology, prevention and treatment. Indian Journal of Occupational and Environmental Medicine, 16(2), 59–62. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Pena, E. , El Alam, S. , Siques, P. , & Brito, J. (2022). Oxidative stress and diseases associated with high‐altitude exposure. Antioxidants, 11(2), 267. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Pennardt, A. (2013). High‐altitude pulmonary edema: Diagnosis, prevention, and treatment. Current Sports Medicine Reports, 12(2), 115–119. [DOI] [PubMed] [Google Scholar]
  24. Qin, H. , & Zhao, A. (2020). Mesenchymal stem cell therapy for acute respiratory distress syndrome: From basic to clinics. Protein & Cell, 11(10), 707–722. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Sarada, S. , Himadri, P. , Mishra, C. , Geetali, P. , Ram, M. S. , & Ilavazhagan, G. (2008). Role of oxidative stress and NFkB in hypoxia‐induced pulmonary edema. Experimental Biology and Medicine (Maywood, N.J.), 233(9), 1088–1098. [DOI] [PubMed] [Google Scholar]
  26. Scherrer, U. , Rexhaj, E. , Jayet, P. Y. , Allemann, Y. , & Sartori, C. (2010). New insights in the pathogenesis of high‐altitude pulmonary edema. Progress in Cardiovascular Diseases, 52(6), 485–492. [DOI] [PubMed] [Google Scholar]
  27. Scherrer, U. , Vollenweider, L. , Delabays, A. , Savcic, M. , Eichenberger, U. , Kleger, G. R. , Fikrle, A. , Ballmer, P. E. , Nicod, P. , & Bärtsch, P. (1996). Inhaled nitric oxide for high‐altitude pulmonary edema. The New England Journal of Medicine, 334(10), 624–629. [DOI] [PubMed] [Google Scholar]
  28. Shan, S. K. , Lin, X. , Li, F. , Xu, F. , Zhong, J. Y. , Guo, B. , Wang, Y. , Zheng, M. H. , Wu, F. , & Yuan, L. Q. (2019). Exosomes and bone disease. Current Pharmaceutical Design, 25(42), 4536–4549. [DOI] [PubMed] [Google Scholar]
  29. Suhail, M. (2010). Na, K‐ATPase: Ubiquitous multifunctional transmembrane protein and its relevance to various pathophysiological conditions. Journal of Clinical Medicine Research, 2(1), 1–17. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Swenson, E. R. , & Bärtsch, P. (2012). High‐altitude pulmonary edema. Compr Physiol, 2(4), 2753–2773. [DOI] [PubMed] [Google Scholar]
  31. Swenson, E. R. , Maggiorini, M. , Mongovin, S. , Gibbs, J. S. , Greve, I. , Mairbäurl, H. , & Bärtsch, P. (2002). Pathogenesis of high‐altitude pulmonary edema: Inflammation is not an etiologic factor. JAMA, 287(17), 2228–2235. [DOI] [PubMed] [Google Scholar]
  32. Tian, L. , Zhao, C. , Yan, Y. , Jia, Q. , Cui, S. , Chen, H. , Li, X. , Jiang, H. , Yao, Y. , He, K. , & Zhao, X. (2024). Ceramide‐1‐phosphate alleviates high‐altitude pulmonary edema by stabilizing circadian ARNTL‐mediated mitochondrial dynamics. Journal of Advanced Research, 60, 75–92. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Vivona, M. L. , Matthay, M. , Chabaud, M. B. , Friedlander, G. , & Clerici, C. (2001). Hypoxia reduces alveolar epithelial sodium and fluid transport in rats: Reversal by beta‐adrenergic agonist treatment. American Journal of Respiratory Cell and Molecular Biology, 25(5), 554–561. [DOI] [PubMed] [Google Scholar]
  34. Wang, C. , Mao, Z. , Gomchok, D. , Li, X. , Liu, H. , Shao, J. , Cao, H. , Xue, G. , Lv, L. , Duan, J. , Wuren, T. , & Wang, H. (2025). Small extracellular vesicles derived from miRNA‐486 overexpressed dental pulp stem cells mitigate high altitude pulmonary edema through PTEN/PI3K/AKT/eNOS pathway. Heliyon, 11(2), e41960. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Ware, L. B. , & Matthay, M. A. (2005). Clinical practice. Acute pulmonary edema. The New England Journal of Medicine, 353(26), 2788–2796. [DOI] [PubMed] [Google Scholar]
  36. Wu, T. , Liu, Y. , Wang, S. , & Shi, C. (2025). MSC‐derived extracellular vesicles: Roles and molecular mechanisms for tissue repair. International Journal of Nanomedicine, 20, 7953–7974. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Yamada, Y. , Nakamura‐Yamada, S. , Kusano, K. , & Baba, S. (2019). Clinical potential and current Progress of dental pulp stem cells for various systemic diseases in regenerative medicine: A concise review. International Journal of Molecular Sciences, 20(5), 1132. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Yue, X. , & Guidry, J. J. (2019). Differential protein expression profiles of Bronchoalveolar lavage fluid following lipopolysaccharide‐induced direct and indirect lung injury in mice. International Journal of Molecular Sciences, 20(14), 3401. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Zhang, W. , Wang, T. , Xue, Y. , Zhan, B. , Lai, Z. , Huang, W. , Peng, X. , & Zhou, Y. (2023). Research progress of extracellular vesicles and exosomes derived from mesenchymal stem cells in the treatment of oxidative stress‐related diseases. Frontiers in Immunology, 14, 1238789. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Zhao, M. , Liu, S. , Wang, C. , Wang, Y. , Wan, M. , Liu, F. , Gong, M. , Yuan, Y. , Chen, Y. , Cheng, J. , Lu, Y. , & Liu, J. (2021). Mesenchymal stem cell‐derived extracellular vesicles attenuate mitochondrial damage and inflammation by stabilizing mitochondrial DNA. ACS Nano, 15(1), 1519–1538. [DOI] [PubMed] [Google Scholar]
  41. Zhou, Q. , Wang, D. , Liu, Y. , Yang, X. , Lucas, R. , & Fischer, B. (2017). Solnatide demonstrates profound therapeutic activity in a rat model of pulmonary edema induced by acute hypobaric hypoxia and exercise. Chest, 151(3), 658–667. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Data S1.

PHY2-14-e70810-s001.xlsx (18.5KB, xlsx)

Data S2.

PHY2-14-e70810-s002.pdf (1.2MB, pdf)

Data Availability Statement

All data generated or analyzed during this study are included in this published article. However, the data that support the findings of this study are available from the corresponding author upon reasonable request.

Articles from Physiological Reports are provided here courtesy of Wiley

RESOURCES