Hydrogen rescues vascular endothelial cells in obstructive sleep apnea-hypopnea syndrome by modulating nitric oxide

Introduction

Obstructive sleep apnea-hypopnea syndrome (OSAHS) is a highly prevalent sleep disorder affecting approximately 1 billion adults globally, with severe cases linked to a 2- to 3-fold increased risk of cardiovascular morbidity and mortality (1). Characterized by recurrent episodes of partial or complete upper airway collapse during sleep, OSAHS leads to chronic intermittent hypoxia (IH), hypercapnia, and sleep fragmentation (2). Among these pathophysiological hallmarks, IH is recognized as the primary driver of systemic vascular dysfunction, contributing to hypertension, atherosclerosis, and endothelial injury (3,4). Vascular endothelial cells, which line the inner surface of blood vessels, play a pivotal role in maintaining vascular homeostasis through the regulation of vasodilation, anti-inflammatory responses, and thrombotic balance (5). A central mediator of these functions is nitric oxide (NO), a gaseous signaling molecule synthesized by endothelial nitric oxide synthase (eNOS) (6). However, in OSAHS, IH-induced oxidative stress disrupts NO bioavailability, leading to endothelial dysfunction—a critical precursor to cardiovascular diseases (7,8).

The cyclical nature of IH in OSAHS, alternating between hypoxia and reoxygenation, mimics ischemia-reperfusion injury, generating excessive reactive oxygen species (ROS) through mitochondrial dysfunction and nicotinamide adenine dinucleotide phosphate oxidase activation (9). ROS, including superoxide (O₂·⁻) and hydrogen peroxide (H₂O₂), overwhelm endogenous antioxidant defenses [e.g., superoxide dismutase (SOD), glutathione peroxidase], resulting in oxidative damage to lipids, proteins, and DNA (10). In endothelial cells, ROS directly oxidize tetrahydrobiopterin (BH4), a critical cofactor for eNOS enzymatic activity (11). BH4 depletion causes eNOS “uncoupling”, shifting its function from NO synthesis to O₂·⁻ production, further exacerbating oxidative stress and forming peroxynitrite (ONOO⁻), a potent oxidant that nitrosylates proteins and inactivates NO (11). This vicious cycle of ROS generation, eNOS uncoupling, and NO deficiency underpins the endothelial dysfunction observed in OSAHS patients (12).

Endothelial dysfunction in OSAHS manifests as impaired endothelium-dependent vasodilation, increased vascular permeability, and pro-inflammatory cytokine release. Studies demonstrated significantly reduced vasodilatory capacity in OSAHS patients compared to healthy controls, correlating with the severity of nocturnal hypoxia (13,14). Furthermore, elevated levels of circulating adhesion molecules [e.g., intercellular cell adhesion molecule-1 (ICAM-1), vascular cell adhesion molecule-1] and inflammatory mediators [e.g., tumor necrosis factor-α (TNF-α), interleukin-6] indicate a chronic inflammatory state that perpetuates vascular injury (15-17). These pathological changes not only accelerate atherosclerosis but also contribute to resistant hypertension and heart failure, highlighting the urgent need for therapies targeting endothelial protection in OSAHS.

The gold-standard treatment for OSAHS, continuous positive airway pressure (CPAP), effectively alleviates IH and improves cardiovascular outcomes (18). However, poor patient adherence (30–60%) and incomplete reversal of endothelial damage in long-term users limit its utility (19,20). Pharmacological interventions, such as statins and angiotensin-converting enzyme inhibitors, show modest benefits but lack specificity for OSAHS-related oxidative pathways (21). Antioxidants like vitamin C and N-acetylcysteine have been explored, but their non-selective ROS scavenging often disrupts redox signaling essential for cellular function (22,23). Thus, there is a pressing demand for novel therapies that selectively neutralize cytotoxic ROS while preserving physiological redox homeostasis.

Molecular hydrogen (H₂) has emerged as a unique modulator of redox homeostasis with potential hormetic effects, influencing beneficial signaling molecules such as NO (H₂O₂ and ·NO) and H₂O₂ (·OH and ONOO⁻), rather than through direct scavenging (24). Its small molecular size and non-polar nature allow rapid diffusion across cell membranes and blood-brain barriers, enabling efficient targeting of mitochondrial and nuclear compartments (25). Preclinical studies demonstrate H₂’s efficacy in diverse oxidative stress-related conditions, including ischemia-reperfusion injury, neurodegenerative diseases, and metabolic syndrome (26-28). In cardiovascular contexts, H₂ inhalation attenuates endothelial apoptosis in hyperglycemia and improves endothelial function in smokers by enhancing NO bioavailability (29). However, its role in OSAHS-induced endothelial injury remains unexplored, leaving a critical gap in understanding its therapeutic potential for this high-risk population.

Given the centrality of ROS-mediated eNOS uncoupling in OSAHS-related endothelial dysfunction, we hypothesized that H₂ could rescue endothelial cells by restoring NO homeostasis through dual mechanisms: (I) reducing oxidative stress to preserve BH4 and prevent eNOS uncoupling; and (II) activation of eNOS phosphorylation to enhance NO synthesis. To test this hypothesis, we employed a translational approach combining in vitro and in vivo models of IH. Human umbilical vein endothelial cells (HUVECs) exposed to IH cycles and OSAHS-modeled rats were treated with H₂-rich medium or inhaled H₂, respectively. We assessed oxidative stress markers, eNOS activity, NO bioavailability, and vascular function to elucidate the molecular pathways underlying H₂’s protective effects.

This study is the first to investigate H₂’s efficacy in OSAHS-associated endothelial injury, offering mechanistic insights into its antioxidant and NO-modulating properties. Our findings may pave the way for H₂-based adjunctive therapies to complement CPAP, particularly in patients with residual cardiovascular risk. We present this article in accordance with the ARRIVE and MDAR reporting checklists (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1345/rc).

Methods

Experimental design

This study employed in vitro and in vivo models to investigate H₂ therapy for IH-induced endothelial dysfunction, mimicking OSAHS. In vitro, HUVECs were subjected to IH cycles (5 min at 1% O₂, 10 min at 21% O₂) for 24 hours, with or without H₂-rich medium (0.6 mM). These cells were divided into three groups: Control, IH, and IH + H₂. In vivo, Sprague-Dawley rats were exposed to IH (8% O₂ for 5 min, 21% O₂ for 5 min; 12 cycles/hour, 8 hours/day) for 4 weeks and treated with inhaled H₂ (2%, 1 hour/day). Animals were divided into three groups: Sham, OSAHS, and OSAHS + H₂, n=6–8 rat/group. Outcomes included oxidative stress markers, NO metabolism, inflammation, apoptosis, and vascular remodeling. All experimental procedures were approved by the Animal Ethics Committee of Chengdu Medical College (approved animal Protocol No. CMC2024MS522), in compliance with US National Institutes of Health guidelines for the care and use of animals (30).

Cell culture, IH model, and H₂ treatment

Cell culture

HUVECs (ScienCell Research Laboratories, Carlsbad, CA, USA, Cat# 8000) were cultured in endothelial cell medium (ECM, ScienCell, Cat# 1001) supplemented with 5% fetal bovine serum (ScienCell, Cat# 0025), 1% endothelial cell growth supplement (ScienCell, Cat# 1052), and 1% penicillin/streptomycin (ScienCell, Cat# 0503). Cells were maintained at 37 ℃ in a humidified 5% CO₂ incubator (Thermo Fisher Scientific, Waltham, MA, USA; Model 3111) and passaged at 80–90% confluence using 0.25% trypsin-EDTA (Gibco, Grand Island, NY, USA; Cat# 25200072). Experiments used cells from passages 3–6.

IH induction

Cells were placed in a modular hypoxia chamber (Billups-Rothenberg, Del Mar, CA, USA; Model MIC-101) programmed for cyclic hypoxia. Each IH cycle consisted of 5 min at 1% O₂ (hypoxia phase, achieved by flushing with a gas mixture of 94% N₂, 5% CO₂, and 1% O₂) followed by 10 min at 21% O₂ (reoxygenation phase, ambient air). Cycles were repeated continuously for 24 hours. Normoxic controls were maintained at 21% O₂.

H₂ treatment and grouping

H₂-rich medium (0.6 mM) was generated by bubbling 99.99% pure H₂ gas (Chengdu Hongjin Chemical Co., Chengdu, China) through ECM at 0.4 MPa for 30 min. H₂ concentration was verified using a H₂-sensitive electrode (Unisense, Aarhus, Denmark; Model H2-100). Cells were treated with H₂-rich medium during IH exposure, with medium refreshed every 6 hours to maintain effective H₂ concentrations, a standard protocol to compensate for the rapid dissipation of H₂ from aqueous solutions (24). Cells were divided into three groups: Control, IH, and IH + H₂.

Animals, OSAHS induction, and H₂ inhalation treatment

Animal housing

Male Sprague-Dawley rats (200–220 g, 8 weeks old) were housed in a controlled environment (22±1 ℃, 12-hour light/dark cycle) with ad libitum access to standard chow (Keao Xieli Feed Co., Beijing, China) and water. All animals were randomly numbered according to a random number table generated in Excel (Microsoft, Redmond, WA, USA). The random numbers were sorted in increasing order and assigned to different groups. Numerical sample identifiers were used during the experimental procedures and data analysis, and the investigators were blinded to the treatment.

OSAHS model

To construct the OSAHS model (OSAHS group), rats were placed in a hypoxic chamber (BioSpherix OxyCycler, Redfield, NY, USA; Model A84XOV) programmed for IH cycles: 5 min at 8% O₂ (achieved by mixing N₂ and compressed air) followed by 5 min at 21% O₂, repeated 12 times/hour for 8 hours/day (9:00 AM–5:00 PM) over 4 weeks. Sham controls were maintained at 21% O₂.

H₂ inhalation and grouping

OSAHS rats received 2% H₂ gas (balanced with compressed air) via a nose-only inhalation system (Scireq, Montreal, QC, Canada; Model PLY3213) for 1 hour/day immediately post-IH exposure. Gas flow rate was calibrated to 1 L/min using a mass flow controller (Alicat Scientific, Tucson, AZ, USA; Model MC-5SLPM). This flow rate, which exceeds the typical minute ventilation of rats, was selected to ensure that the chamber was continuously flushed, preventing CO₂ buildup and maintaining a stable atmospheric environment within the nose-only chamber. The concentration of H₂ within the delivery system was verified in real-time using a H₂ sensor (Unisense, Model H2-100) at the outlet port proximal to the animal. Under these conditions, the system reliably delivers a time-averaged H₂ exposure of 2%. Control OSAHS rats inhaled compressed air under identical conditions. Animals were divided into three groups: Sham, OSAHS, and OSAHS + H₂, n=6–8 rat/group.

Tissue collection

After 4 weeks, rats were anesthetized with sodium pentobarbital (50 mg/kg, i.p.), and thoracic aortas were excised. Tissues were either fixed in 4% paraformaldehyde (24 hours for histology) or snap-frozen in liquid nitrogen (stored at −80 ℃ for molecular assays).

Histological staining

Hematoxylin-eosin (H&E)

Paraffin-embedded aortic sections (5 µm) were deparaffinized, stained with H&E (Solarbio, Beijing, China; Cat# G1120), and imaged under a light microscope (Nikon Eclipse E100, Tokyo, Japan). Medial thickness was measured using ImageJ [National Institutes of Health (NIH)] at 10 random sites per section.

Masson’s trichrome

Sections were stained with Masson’s kit (Solarbio, Cat# G1340) to assess collagen deposition. Fibrosis area (blue staining) was quantified as a percentage of total aortic area using ImageJ.

ROS measurement

In vitro: HUVECs were incubated with 10 µM 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA; Sigma-Aldrich, Burlington, USA; Cat# D6883) for 30 min at 37 ℃. Fluorescence intensity (excitation/emission: 488/525 nm) was measured using a microplate reader (BioTek Synergy H1, Winooski, VT, USA).

In vivo: aortic homogenates were incubated with 10 µM dihydroethidium (DHE, Sigma-Aldrich, Cat# D7008) for 30 min. Superoxide levels were quantified by fluorescence microscopy (Olympus BX53, Tokyo, Japan).

RT-PCR

Total RNA was extracted from HUVECs using TRIzol (Invitrogen, Carlsbad, CA, USA; Cat# 15596026). cDNA was synthesized with a Reverse Transcription Kit (Takara, Dalian, China; Cat# RR037A). Reverse transcription quantitative polymerase chain reaction (RT-qPCR) was performed using SYBR Green Master Mix (Roche, Mannheim, Germany; Cat# 04887352001) on a QuantStudio 5 Real-Time PCR System (Thermo Fisher). PCR reactions were performed in a 25 µL volume containing SYBR Green Master Mix (Roche) and 0.5 µM primers. Cycling conditions: 95 ℃ for 5 min, followed by 40 cycles of 95 ℃ for 30 s, 60 ℃ for 30 s, and 72 ℃ for 30 s. Primer sequences were designed using National Center for Biotechnology Information (NCBI) Primer-BLAST and synthesized by Sangon Biotech (Shanghai, China):

TNF-α:
Forward 5'-CAGGCGGTGCCTATGTCTC-3',
Reverse 5'-CGATCACCCCGAAGTTCAGT-3'.

ICAM-1:
Forward 5'-CTGTGGCCTGGAGCTGTTTA-3',
Reverse 5'-CAGGGGCCGTACAGTTGATA-3'.

GAPDH:
Forward 5'-GGAGCGAGATCCCTCCAAAAT-3',
Reverse 5'-GGCTGTTGTCATACTTCTCATGG-3'.

Western blot

Proteins from HUVECs or aortic tissue were extracted using RIPA buffer (Beyotime, Shanghai, China; Cat# P0013B). Lysates (30 µg/lane) were separated on 10% SDS-PAGE gels and transferred to PVDF membranes (Millipore, Burlington, MA, USA; Cat# IPFL00010). Membranes were blocked with 5% non-fat milk, then incubated overnight at 4 ℃ with primary antibodies: Phospho-eNOS (Ser1177) (Cell Signaling Technology, Danvers, MA, USA; Cat# 9571, 1:1,000); Total eNOS (Abcam, Cambridge, UK; Cat# ab76198, 1:1,000); ICAM-1 (Proteintech, Wuhan, China; Ca# 60299-1-I, 1:1,000). β-actin (Proteintech, Cat# 66009-1-Ig, 1:5,000). After horseradish peroxidase (HRP)-conjugated secondary antibody incubation (1:5,000, Proteintech), bands were visualized using electrochemiluminescence (ECL; Millipore, Cat# WBKLS0500) and quantified with ImageJ.

Enzyme-linked immunosorbent assay (ELISA)

Plasma BH4 and BH2 levels were measured using commercial ELISA kits (R&D Systems, Minneapolis, MN, USA; Cat# K9965-100 and K9966-100). Aortic malondialdehyde (MDA) was quantified with an MDA ELISA kit (Bioswamp, Wuhan, China; Cat# RA20003).

Terminal deoxynucleotidyl transferase (TdT) dUTP nick-end labeling (TUNEL) staining

Apoptotic cells were detected using the TUNEL assay (commercial kit name, e.g., Roche #11684795910 or equivalent). Sections were deparaffinized, rehydrated, and treated with proteinase K (20 µg/mL, 15 min, 37 ℃). After permeabilization, slides were incubated with TUNEL reaction mixture (TdT enzyme + fluorescein-dUTP) for 60 min at 37 ℃ in a humidified chamber. Nuclei were counterstained with 4',6-diamidino-2-phenylindole (DAPI; 1 µg/mL, 10 min). Endothelial apoptosis was evaluated in the aortic endothelium. Six random fields per section were imaged under a fluorescence microscope (Zeiss Axio Imager, Göttingen, Germany; 20× objective, scale bar: 20 µm). TUNEL-positive endothelial cells (green fluorescence) and total DAPI-stained nuclei (blue) were counted using ImageJ/Fiji software. The apoptotic index was calculated as: (TUNEL⁺ endothelial cells_/total endothelial cells) × 100%.

Statistical analysis

Data are expressed as mean ± standard error of the mean standard error of the mean (SEM; n=3 independent experiments). Between-group differences were analyzed using Student’s t-test (two groups) or one-way analysis of variance (ANOVA) with Tukey’s post hoc test (multiple groups) in GraphPad Prism 9.0. Significance was set at *, P<0.05; **, P<0.05; ***, P<0.05 vs. Control or Sham. ^#, P<0.05; ^##, P<0.01; ^###, P<0.001 vs. IH or OSAHS.

Results

H₂ mitigates IH-induced oxidative stress in endothelial cells and OSAHS rats

In vitro, HUVECs exposed to IH cycles (1% O₂ for 5 min followed by 21% O₂ for 10 min, repeated for 24 h) exhibited a 3.2-fold increase in intracellular ROS levels compared to normoxic controls (P<0.001 vs. Control, Figure 1A), as quantified by DCFH-DA fluorescence. Treatment with H₂-rich medium (0.6 mM) during IH exposure significantly reduced ROS production by 60% (P<0.001 vs. IH group, Figure 1A). In vivo, OSAHS rats subjected to 4-week IH (8% O₂ for 5 min alternating with 21% O₂, 12 cycles/h, 8 h/day) showed a 2.3-fold elevation in aortic MDA content (P<0.01 vs. Sham group, Figure 1B), a marker of lipid peroxidation. Daily inhalation of 2% H₂ gas for 1 h reduced MDA levels by 45% (P<0.05 vs. OSAHS group, Figure 1B), confirming H₂’s antioxidative efficacy in vascular tissues.

Figure 1 Hydrogen alleviates IH-induced oxidative stress in endothelial cells and OSAHS rats. (A) Intracellular ROS in HUVECs under normoxia (Control), intermittent hypoxia (IH), and IH with hydrogen-rich medium (IH + H₂) was quantified using DCFH-DA fluorescence. Data represent mean ± SEM (n=3). ***, P<0.001 vs. Control; ^###, P<0.001 vs. IH. (B) Aortic MDA content in Control, OSAHS, and OSAHS + H₂ rats was measured via ELISA. Data represent mean ± SEM (n=3). **, P<0.01 vs. Sham; ^#, P<0.05 vs. OSAHS. DCFH-DA, 2',7'-dichlorodihydrofluorescein diacetate; ELISA, enzyme-linked immunosorbent assay; H₂, hydrogen; HUVECs, human umbilical vein endothelial cells; IH, intermittent hypoxia; MDA, malondialdehyde; OSAHS, obstructive sleep apnea-hypopnea syndrome; ROS, reactive oxygen species; SEM, standard error of the mean.

H₂ restores NO bioavailability via eNOS recoupling and BH4 preservation

IH impaired eNOS activity in HUVECs, as evidenced by a 65% reduction in phosphorylated eNOS (Ser1177) levels (P<0.001 vs. Control, Figure 2A,2B) and a 55% decrease in NO metabolites (nitrite/nitrate, P<0.01 vs. Control, Figure 2C). H₂ treatment reversed these effects, increasing eNOS phosphorylation 1.8-fold (P<0.01 vs. IH group, Figure 2A,2B) and restoring NO metabolites to 85% of IH group values (P<0.05, Figure 2C). Mechanistically, IH depleted the BH4/BH2 ratio by 70% (P<0.001 vs. Control, Figure 2D), a critical determinant of eNOS coupling. H₂ supplementation elevated the BH4/BH2 ratio 2.2-fold (P<0.05 vs. IH group, Figure 2D), effectively preventing eNOS uncoupling and superoxide overproduction (P<0.001 vs. IH, Figure 2E). In OSAHS rats, acetylcholine-induced endothelium-dependent vasodilation was impaired by 50% (P<0.01 vs. Sham, Figure 2F), while H₂ inhalation improved vascular relaxation by 40% (P<0.01 vs. OSAHS group, Figure 2F), corroborating functional recovery of NO-mediated vasodilation.

Figure 2 Hydrogen restores NO bioavailability via eNOS recoupling and BH4 preservation. (A) Phosphorylated eNOS (Ser1177) levels in HUVECs were analyzed by Western blot with phospho-eNOS and total eNOS antibodies. β-actin served as loading controls. (B) Quantification of phosphorylated eNOS (Ser1177) levels normalized to β-actin. ***, P<0.001 vs. Control; ^##, P<0.01 vs. IH. (C) Nitrite/nitrate (NO metabolites) in HUVECs was quantified via Griess assay. **, P<0.01 vs. Control; ^#, P<0.05 vs. IH. (D) BH4/BH2 ratio in HUVECs was measured via ELISA. ***, P<0.001 vs. Control; ^#, P<0.05 vs. IH. (E) Superoxide levels in HUVECs were measured by dihydroethidium fluorescence. ***, P<0.001 vs. Control; ^###, P<0.001 vs. IH. (F) Acetylcholine-induced endothelium-dependent vasodilation in rat aortas was assessed through vascular reactivity. Data are expressed as % relaxation (mean ± SEM, n=3). **, P<0.01 vs. Sham; ^##, P<0.01 vs. OSAHS. ACh, acetylcholine; BH4, tetrahydrobiopterin; ELISA, enzyme-linked immunosorbent assay; eNOS, endothelial nitric oxide synthase; H₂, hydrogen; HUVECs, human umbilical vein endothelial cells; IH, intermittent hypoxia; NO, nitric oxide; OSAHS, obstructive sleep apnea-hypopnea syndrome; SEM, standard error of the mean.

H₂ ameliorates IH-driven vascular remodeling and inflammation

Histopathological analysis revealed significant aortic remodeling in OSAHS rats. H&E staining demonstrated a 2.1-fold increase in medial thickness (P<0.01 vs. Sham, Figure 3A,3B), while Masson’s trichrome staining showed a 3.4-fold elevation in collagen deposition (P<0.001 vs. Sham, Figure 3C,3D). H₂ treatment attenuated these structural abnormalities, reducing medial hyperplasia by 35% (P<0.05 vs. OSAHS, Figure 3A,3B) and fibrosis area by 48% (P<0.01 vs. OSAHS, Figure 3C,3D). Concurrently, IH upregulated pro-inflammatory markers: TNF-α messenger ribonucleic acid (mRNA) in HUVECs increased 4.1-fold (P<0.001 vs. Control, Figure 3E), and aortic ICAM-1 protein in OSAHS rats surged 3.8-fold (P<0.001 vs. Sham, Figure 3F,3G). H₂ administration suppressed these responses, reducing TNF-α expression by 50% (P<0.05 vs. IH, Figure 3E) and ICAM-1 levels by 40% (P<0.05 vs. OSAHS, Figure 3F,3G).

Figure 3 Hydrogen attenuates vascular remodeling and inflammation in OSAHS. (A) Representative H&E staining of aortic sections (scale bar: 20 µm). (B) Quantification of medial thickness. **, P<0.01 vs. Sham; ^#, P<0.05 vs. OSAHS. (C) Representative Masson staining of aortic sections (scale bar: 20 µm). (D) Fibrosis area quantification. ***, P<0.001 vs. Sham; ^##, P<0.01 vs. OSAHS. (E) RT-PCR analysis TNF-α mRNA in HUVECs. ***, P<0.001 vs. Control; ^#, P<0.05 vs. IH. (F) Aortic ICAM-1 protein levels in rats were detected via Western blot. β-actin served as loading controls. (G) Quantification of ICAM-1 levels normalized to β-actin. ***, P<0.001 vs. Sham; ^#, P<0.05 vs. OSAHS. H&E, hematoxylin-eosin; HUVECs, human umbilical vein endothelial cells; ICAM-1, intercellular cell adhesion molecule-1; IH, intermittent hypoxia; mRNA, messenger ribonucleic acid; OSAHS, obstructive sleep apnea-hypopnea syndrome; RT-PCR, reverse transcription polymerase chain reaction; TNF-α, tumor necrosis factor-α.

Systemic and cellular validation of H₂’s therapeutic effects

Plasma analysis in OSAHS rats revealed a 2.6-fold increase in 8-isoprostane (P<0.001 vs. Sham, Figure 4A), a systemic oxidative stress marker, which was normalized by 62% with H₂ inhalation (P<0.01 vs. OSAHS, Figure 4A). In HUVECs, IH reduced SOD activity by 75% (P<0.01 vs. Control, Figure 4B), and H₂ treatment restored SOD activity to near-control levels (P<0.01 vs. IH, Figure 4B). TUNEL staining further demonstrated that H₂ reduced IH-induced endothelial apoptosis in aortic tissues by 55% (P<0.01 vs. OSAHS, Figure 4C,4D), aligning with preserved vascular integrity.

Figure 4 Systemic and cellular validation of hydrogen’s therapeutic effects. (A) ELISA measurement of systemic oxidative stress marker plasma 8-isoprostane in rats. ***, P<0.001 vs. Sham; ^##, P<0.01 vs. OSAHS. (B) SOD activity assay in HUVECs. **, P<0.01 vs. Control; ^##, P<0.01 vs. IH. (C) Endothelial apoptosis in aortic tissues was detected via TUNEL staining (scale bar: 20 µm). Yellow arrows indicate TUNEL-positive apoptotic endothelial cells. (D) The apoptotic index was calculated as TUNEL⁺ endothelial cells/total endothelial cells × 100%. Data are expressed as mean ± SEM (n=3). **, P<0.001 vs. Sham; ^##, P<0.01 vs. OSAHS. DAPI, 4',6-diamidino-2-phenylindole; ELISA, enzyme-linked immunosorbent assay; HUVECs, human umbilical vein endothelial cells; IH, intermittent hypoxia; OSAHS, obstructive sleep apnea-hypopnea syndrome; SEM, standard error of the mean; SOD, superoxide dismutase; TUNEL, terminal deoxynucleotidyl transferase (TdT) dUTP nick-end labeling.

Discussion

Our findings reveal that H₂ supplementation attenuates IH-induced oxidative stress, recouples eNOS, and preserves vascular structural integrity, thereby addressing a critical gap in OSAHS therapeutics. Therefore, the present study provides compelling evidence that molecular H₂ effectively mitigates vascular endothelial dysfunction in OSAHS by restoring NO homeostasis through dual antioxidative and anti-inflammatory mechanisms. While a recent clinical study by Li et al., 2025 demonstrated the efficacy of H₂-rich water in improving metabolic profiles in OSAHS patients (31), the specific mechanisms underlying H₂’s protection against vascular endothelial injury remain elusive. Our results not only expand the known cardioprotective properties of H₂ but also provide mechanistic insights into its role in counteracting OSAHS-specific endothelial injury, by demonstrating that H₂ attenuates oxidative stress, recouples eNOS, and restores NO bioavailability.

A key innovation of this study lies in its translational approach, bridging in vitro and in vivo models to elucidate H₂’s multifaceted mechanisms. Unlike previous studies focusing on H₂’s antioxidant effects in chronic hypoxia or ischemia-reperfusion models (32), we specifically targeted the cyclical IH pattern of OSAHS, which uniquely drives oscillatory ROS bursts and eNOS uncoupling. By demonstrating that H₂ preserves the BH4/BH2 ratio—a determinant of eNOS coupling—we provide mechanistic clarity to earlier observations of H₂-enhanced NO bioavailability in hyperglycemia and smoking models. Our data align with reports that BH4 depletion underpins endothelial dysfunction in cardiovascular diseases but extend this paradigm to OSAHS (33), where IH-induced oxidative stress is both a cause and consequence of eNOS dysregulation. The 2.2-fold restoration of the BH4/BH2 ratio by H₂ highlights its specificity in targeting redox-sensitive cofactors, contrasting with non-selective antioxidants like vitamin C, which may disrupt physiological ROS signaling. This specificity suggests that H₂ may act not as a direct stoichiometric scavenger, but rather by indirectly modulating redox balance, potentially through mechanisms such as mild mitochondrial stress and activation of pathways like nuclear factor erythroid 2-related factor 2, thereby preserving beneficial redox signaling.

The functional significance of H₂’s effects are underscored by the 40% improvement in endothelium-dependent vasodilation in OSAHS rats, a metric directly relevant to clinical outcomes. This aligns with human studies showing impaired flow-mediated dilation in OSAHS patients, yet no prior therapy has simultaneously addressed oxidative stress, NO deficiency, and vascular remodeling (34). Notably, H₂’s anti-inflammatory actions—evidenced by 50% suppression of TNF-α and 40% reduction in ICAM-1—complement its antioxidative effects, mirroring its efficacy in inflammatory conditions like rheumatoid arthritis. However, our work uniquely links these effects to OSAHS-associated endothelial pathology, suggesting that H₂’s pleiotropic actions could synergize with CPAP to address residual cardiovascular risk in poorly adherent patients. Despite these advances, several limitations warrant consideration. First, while our rat model replicates IH patterns, it does not fully capture the complexity of human OSAHS, which involves comorbid obesity, metabolic dysregulation, and neural reflexes. Notably, the potential benefits of H₂ in OSAHS may extend beyond improving endothelial function. Given the high prevalence of comorbid metabolic disorders (e.g., insulin resistance, dyslipidemia) in OSAHS patients, the reported metabolic modulating effects of H₂—evidenced by its ability to improve glucose metabolism and lipid profiles in both animal models and emerging clinical research (35,36)—provide an additional compelling rationale for its therapeutic application. This suggests that H₂ could simultaneously target both the cardiovascular and metabolic complications of OSAHS, offering a comprehensive therapeutic strategy. Second, the study focused on acute and subchronic H₂ administration, long-term effects on vascular remodeling and potential tolerance remain unexplored. Third, the precise molecular pathways by H₂ modulates BH4 synthesis (e.g., GTP cyclohydrolase I regulation) were not investigated, leaving room for mechanistic refinement. Future studies should validate these findings in larger animal models and OSAHS patients, assess H₂’s interactions with CPAP, and explore combinatorial therapies targeting downstream effectors of eNOS uncoupling, such as arginase or asymmetric dimethylarginine.

Conclusions

In conclusion, this study positions H₂ as a promising adjunctive therapy for OSAHS-related endothelial dysfunction, offering a mechanistically grounded strategy to break the vicious cycle of chronically elevated oxidative stress, NO deficiency, and dysregulated inflammation. By addressing both molecular and functional endpoints, our work advances the translational potential of H₂ while highlighting the need for clinical trials to evaluate its efficacy in improving cardiovascular outcomes in this high-risk population.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the ARRIVE and MDAR reporting checklists. Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1345/rc

Data Sharing Statement: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1345/dss

Peer Review File: Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1345/prf

Funding: This study was supported by the Administration of Traditional Chinese Medicine Science and Technology Research Project of Sichuan Province (No. 2024MS522) and the Medical Research Project of Chengdu City in 2025 (No. 2025649).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2025-1345/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All experimental procedures were approved by the Animal Ethics Committee of Chengdu Medical College (approved animal protocol No. CMC2024MS522) in compliance with US National Institutes of Health guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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