The experimental protocols and animal care procedures employed in the present study were approved by the Medical Ethics Committee of Qinghai Provincial People's Hospital (approval no. 2023-95; Xinning, China). Male Sprague-Dawley rats (weight, 200–250 g; age, 7–8 weeks) were procured from Jiangsu Huachuang Xinnuo Pharmaceutical Science and Technology Co., Ltd.

The rat model of hypoxia was created by exposing the animals to a hypobaric hypoxia chamber (cat. no. DYC-300; Guizhou Fenglei Oxygen Chamber Co., Ltd.). Previous studies have shown that acute mountain sickness usually occurs within a few hours to 24 h upon entering high-altitude environments, with symptoms peaking within 24–72 h and then gradually subsiding (17–20). Thus, the present study centered on the first 3 days of acute hypoxia. Meanwhile, for an improved comparison of the physiological changes under acute hypoxia, it additionally included the observation results of hypoxic treatments at 7 days and 4 weeks. The initial animal models were exposed to hypoxia or normoxia for 0, 1, 2, 3 and 7 days and for 4 weeks (n=60; 5/group; each group comprised different animals). The iron intervention animal models were exposed to hypoxia for 3 days (n=40; 10/group). The air pressure and oxygen content corresponded to an altitude of 6,000 meters above sea level. The pressure was 52.9 kPa, the oxygen concentration was 10% (hypoxia), the temperature was 22.0°C, the relative humidity was 50% and there was a 12-h light/dark cycle (21–23).

The pharmacological intervention rat model was generated as follows: Male Sprague-Dawley rats were administered deferoxamine (DFO; 200 mg/kg; cat. no. HY-B0988; MedChemExpress) in the hypoxia + DFO (HD) group or iron sucrose (20 mg/kg; cat. no. HY-B2068; MedChemExpress) in the hypoxia + iron sucrose (HI) group via daily intraperitoneal injection (24). The rats were then placed in a hypoxic (10%) environment for 3 days, as previously described (19–21). The control groups received an equivalent volume of solvent via intraperitoneal injection under either hypoxic (10% O₂; HC group) or normoxic (21% O₂; NC group) conditions.

The rats (n=3/group) were anesthetized with isoflurane gas (induction, 2%; maintenance, 1.5–2.0%; inhalation) and the mean pulmonary artery pressure (mPAP) was determined using a right heart catheter. The catheter was positioned in the right external jugular vein and advanced into the pulmonary artery via the right ventricle. The position of the insertion was determined by analyzing the waveforms produced by a biosignal acquisition system (MP150; Biopac Systems, Inc.). After right cardiac catheterization, the rats were sacrificed via cervical dislocation under isoflurane anesthesia, and their death was confirmed by observing the arrest of cardiac and respiratory functions and the disappearance of the pain response. The lung tissue was then excised and its wet weight was precisely measured. Subsequently, the excised lung tissue underwent a 72-h drying process at 65°C; after the completion of this period, the tissue was re-weighed to obtain the dry weight. The lung wet/dry weight ratio was calculated to reflect the water content of the lung tissue.

Blood samples (1 ml) (n=3/group in the hypoxia study at different time points; n=5/group in the pharmacological intervention rat model) were collected from rats via cardiac puncture under 1.5% isoflurane inhalation anesthesia, followed by euthanasia through cervical dislocation while maintaining anesthesia. Aliquots were transferred to EDTA-coated anticoagulant tubes for complete blood count analysis following gentle inversion (5–10 cycles), or were transferred to clot activator tubes for centrifugation (4°C, 1,006 × g, 10 min) to obtain serum. Complete blood count was performed using a YAN-305A hematology analyzer (Shanghai Yuyan Scientific Instrument Co., Ltd.) with species-specific calibration. Serum iron levels were quantified using a commercial kit (cat. no. A039-1-1; Nanjing Jiancheng Bioengineering Institute) according to the manufacturer's guidelines. Raw data were processed for statistical analysis.

Following anesthesia and euthanasia of the rats, the heart and lungs were simultaneously removed and placed in 1X PBS. The main pulmonary artery and its branches were meticulously isolated from the right ventricle. Subsequently, the artery was stabilized with 3% glutaraldehyde for 2 h at 4°C, followed by post-fixation with 1% osmium tetroxide for 1 h at 4°C. A graded series of acetone treatments was employed for dehydration, followed by infiltration with Epon 812 and embedding. The semithin sections (50 nm) were subjected to methylene blue staining at room temperature for 2 min, whereas the ultrathin slides were meticulously sliced using a single diamond knife and were dual-stained with uranyl acetate at room temperature for 30 min and lead citrate at room temperature for 15 min. Subsequently, the slides were subjected to observation under a JEM-1400-FLASH transmission electron microscope (JEOL Ltd.).

The lung tissue and main pulmonary artery from 3 rats/group were fixed in 4% paraformaldehyde at room temperature for 12 h, dehydrated in a graded series of ethanol, cleared with xylene (2×10 min) and embedded in paraffin via triple wax immersion (56°C, 1 h/cycle). Sections (5 µm) were then stained with hematoxylin at room temperature for 5 min and eosin at room temperature for 3 min following standard hematoxylin and eosin (H&E) staining protocols. An image was acquired using a BA400 digital light microscope with a digital interface (Motic Incorporation, Ltd.). The rat lung tissue was evaluated for evidence of lung injury in accordance with the established criteria for pathological assessment of lung injury, as outlined by the American Thoracic Society (25). The scoring indexes were based on the following criteria: Neutrophil infiltration of lung tissue, hyaline membrane formation, fibrin deposition in alveoli and thickening of alveolar septa. Each index was scored on a 5-point scale, with the severity of injury corresponding to the following categories: No injury, 0 points; mild injury, 1 point; moderate injury, 2 points; severe injury, 3 points; and very severe injury, 4 points (22,23). A total of five randomly selected fields of H&E-stained lung tissue from each animal were scored and two experts scored the sections in a double-blind manner.

The protocols for fixation, embedding and sectioning of the main pulmonary artery and alveolar tissue were the same as those performed for H&E staining. Sections were deparaffinized in xylene (2×10 min), rehydrated through a graded ethanol series (5 min/step), and subjected to microwave-mediated antigen retrieval in citrate buffer (pH 6.0) at sub-boiling temperature for 10 min. After cooling, endogenous peroxidase was blocked with 3% H₂O₂ for 10 min at room temperature, followed by blocking with 5% BSA (cat. no. SW3015; Beijing Solarbio Science & Technology Co., Ltd.) for 30 min at room temperature. Primary anti-HIF1α antibody (1:100; cat. no. ab179483; Abcam) was applied overnight at 4°C, followed by an incubation with a HRP-conjugated goat anti-rabbit secondary antibody (1:100; cat. no. GB23303; Wuhan Servicebio Technology Co., Ltd.) for 1 h at room temperature; Subsequently, DAB was employed to develop the color (cat. no. GB1212; Wuhan Servicebio Technology Co., Ltd.), hematoxylin was used to counterstain the nuclei at room temperature for 10 min and neutral balsam was used to seal the mount. Observations were conducted using a BA400 digital light microscope (Motic Incorporation, Ltd.).

Human-derived primary pulmonary artery endothelial cells (cat. no. 337714) were obtained from BeNa Culture Collection; Beijing Beina Chunglian Institute of Biotechnology. The Research Ethics Committee of Qinghai Provincial People's Hospital confirmed that commercially procured human primary cells fall outside the scope of required ethics approval. At the time of purchase, the cells were in the first passage. The cells were cultured in endothelial cell medium (cat. no. ECM 1001; ScienCell Research Laboratories, Inc.) containing 1% vascular endothelial growth factor at 37°C with 5% CO₂. The cells were passaged, and those between passages 3 and 5 were used in the present study. The cells in the hypoxia group were cultured at 37°C under hypoxia (1% O₂) for 2 h.

Human-derived pulmonary artery endothelial cells were treated with DFO (100 µmol/l; cat. no. HY-B0988; MedChemExpress) in the HD group or with iron sucrose (50 µmol/l; cat. no. HY-B2068; MedChemExpress) in the HI group as previously described (26). The control groups received an equivalent volume of solvent under either hypoxic (1% O₂, HC group) or normoxic (21% O₂, NC group) conditions.

For immunofluorescence experiments conducted in vivo (n=3/group), lung vascular tissue samples underwent fixation, dehydration, embedding and sectioning following the aforementioned standard H&E protocols. Subsequently, the sections were dewaxed, rehydrated and subjected to microwave-assisted antigen retrieval in citrate buffer (pH 6.0), as per the immunohistochemistry methods. After blocking with immunostaining buffer (cat. no. P0102; Beyotime Institute of Biotechnology) for 1 h at room temperature, the following primary antibodies were used to incubate the sections overnight at 4°C: PTEN-induced putative kinase 1 (PINK1 1:100; cat. no. GB114934-100; Wuhan Servicebio Technology Co., Ltd.), smooth muscle actin (SMA; 1:100; cat. no. A2547; MilliporeSigma) and HIF1α (1:100; cat. no. ab179483; Abcam). Samples were washed three times with 1X PBS (10 min each), followed by incubation with a fluorescent secondary antibody [Goat Anti-Rabbit Alexa Fluor^® 488 for HIF1α and Caspase 3 (cat. no. ab150077), Alexa Fluor 647 for PINK1 (cat. no. ab150079) or Goat Anti-Mouse Alexa Fluor 647 for SMA (cat. no. ab150115); 1:1,000; Abcam] for 1 h at room temperature. Nuclei were counterstained with DAPI, and images were acquired using a confocal laser microscope (FV1000; Olympus Corporation).

Pulmonary arterial endothelial cells were routinely cultured to an appropriate density and were treated with hypoxia and DFO or iron sucrose for 2 h. Cells were fixed with 4% paraformaldehyde for 15 min at room temperature, permeabilized with 0.1% Triton X-100 for 10 min at room temperature and blocked with immunostaining buffer (cat. no. P0102; Beyotime Institute of Biotechnology) for 1 h at room temperature. Subsequently, the sections were incubated with the primary antibodies against Caspase 3 (1:100; cat. no. 19677-1-AP; Proteintech Group, Inc.) and HIF1α (1:100; cat. no. ab179483; Abcam) overnight at 4°C. The secondary antibody incubation, nuclear counterstaining with DAPI and confocal imaging (FV1000; Olympus Corporation) were performed according to the aforementioned protocols.

To assess mitochondrial membrane potential (MMP) and apoptosis of pulmonary arterial endothelial cells, mitochondrial superoxide and Annexin V-FITC fluorescence assays were conducted in accordance with the instructions provided in the MitoSOX and Annexin V-FITC kits (cat. nos. S0061M and C1071M; Beyotime Institute of Biotechnology). The images were examined using a fluorescence microscope (THUNDER Imager; Leica Microsystems, Inc.) and the immunofluorescence intensity of protein signals was subsequently semi-quantified using ImageJ software (version 1.54f; National Institutes of Health).

Proteins were extracted from lung tissues and pulmonary arterial endothelial cells using RIPA lysis buffer (cat. no. P0013B; Beyotime Institute of Biotechnology) and were quantified using a BCA kit (Thermo Fisher Scientific, Inc.). A 20-µg protein sample from each group was then subjected to SDS-PAGE on a 10% gel, followed by transfer to a polyvinylidene fluoride membrane (PVDF; 0.22 µm; Merck KGaA). Following the transfer of the proteins, 5% skimmed milk (BD Biosciences) was used to block the PVDF membranes for 1 h at room temperature. The membranes were then incubated overnight at 4°C with the following primary antibodies: LC3 (1:1,000; cat. no. 14600-1-AP) and β-actin (1:5,000; cat. no. 66009-1-Ig) (both from Proteintech Group, Inc.). Subsequently, the membranes were incubated with HRP-conjugated secondary antibodies (1:5,000; cat. nos. E-AB-1003 and E-AB-1001; Wuhan Elabscience Biotechnology Co., Ltd.) for 1 h at room temperature and chemiluminescence signals were detected using an ECL kit (Abbkine Scientific Co., Ltd.). Finally, ImageJ software (version 1.54f) was employed to semi-quantify the intensity of the bands (27–29).

The Cell Counting Kit 8 (CCK8) assay (cat. no. E-CK-A362; Wuhan Elabscience Biotechnology Co., Ltd.) was used to investigate cell proliferation in accordance with the manufacturer's instructions. The absorbance at 450 nm was measured using a microplate reader (Biotek; Agilent Technologies, Inc.). In addition, cell proliferation was detected using the Edu Cell Proliferation Kit (cat. no. C0071L; Beyotime Institute of Biotechnology) according to the manufacturer's instructions. Images were captured using a fluorescence microscope (ECHO/RVL-100; Discover Echo, Inc.) and ImageJ software (version 1.54f) was employed for data analysis.

All data were processed using the statistical software package SPSS 28.0 (IBM Corp.) and the graphical software package GraphPad 10.0 (Dotmatics). Data with a normal distribution are presented as the mean ± standard deviation. Statistical comparisons of parameters (including lung wet/dry weight ratio, serum iron and erythroid parameters) across different durations of hypoxic treatment were performed using two-way ANOVA, followed by Tukey's honestly significant difference post hoc test for multiple comparisons. One-way ANOVA followed by the Tukey's post hoc test was performed for multiple group comparisons when the data met the assumptions of normality and homogeneity of variance. Non-normally distributed ordinal data (lung injury scores) are presented as the median (IQR), and were analyzed using Kruskal-Wallis test with Dunn's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

The lung dry/wet weight ratio was significantly increased on days 2 and 3 of hypoxia compared with day 0 (Fig. 1C), indicating the presence of pulmonary edema. Lung morphology results indicated that there were no abnormal changes in lung tissue morphology during days 1–7 under normoxia. However, under hypoxia, erythrocyte leakage from alveolar spaces accompanied by inflammatory cell infiltration was observed. This phenomenon was most pronounced on day 3 of acute hypoxia and then gradually resolved, with a marked infiltration of inflammatory cells reappearing after 4 weeks of hypoxia (Fig. 1A). Semi-quantitative analysis further confirmed that hypoxia-induced lung tissue injury was most pronounced on day 3 and peaked at 4 weeks compared with on day 0 under hypoxia (Fig. 1B).

To elucidate the involvement of iron in acute mountain sickness, serum iron and erythropoiesis levels were quantified under normoxia/hypoxia at multiple timepoints. The results demonstrated that, under normoxia, there were no statistically significant differences in serum iron, red blood cell (RBC), hemoglobin and hematocrit (HCT) levels among the groups. By contrast, under hypoxia, as the duration of hypoxia increased, serum iron concentration exhibited a gradually decreasing trend, the serum iron concentration in the 4-week hypoxia group exhibited statistically significant differences compared with that in the day 0 group. By contrast, RBC, HCT and hemoglobin levels were increased with prolonged hypoxia exposure. RBC parameters in hypoxia groups at day 3 and 7 and week 4 were significantly different compared with those in the day 0 group under hypoxia, whereas hemoglobin and HCT levels in hypoxia groups day 1, 3 and 7, and week 4 exhibited significant changes relative to the day 0 control under hypoxia (Fig. 1D and E). These findings indicated that the oxygen content in the rats was diminished and compensatory erythrocyte hyperplasia was exhibited. Given the pivotal role of iron chelators in the initiation of mitophagy, western blot analysis was performed to assess the expression levels of the mitophagy protein LC3B. The results demonstrated that LC3B-II/LC3B-I expression was significantly upregulated in response to acute hypoxia, particularly on day 3 of hypoxia exposure compared to day 0 under hypoxia, and peaked after 4 weeks of hypoxic treatment (Fig. 1F).

Given the reduced serum iron concentration observed in rats during acute hypoxia, the present study aimed to intervene in iron metabolism in a rat model using an iron chelator and iron sucrose to elucidate the role of iron in acute high-altitude sickness (Fig. 2A). The results of right heart catheterization for mPAP measurements indicated that the iron chelator DFO was associated with an elevation in mPAP under acute hypoxia. By contrast, iron sucrose was associated with a reduction in mPAP elevation under acute hypoxia (Fig. 2B and D). Morphological analysis indicated the presence of alveolar tissue destruction and inflammatory cell infiltration of the alveolar tissue under acute hypoxia, which was accompanied by fracturing and disorganization of elastic fiber septa in the main pulmonary artery (Fig. 2C). Quantitative data corroborated the findings of H&E staining (Fig. 2E). Furthermore, the wet/dry weight ratio of the lungs was increased under acute hypoxia (Fig. 2F). The administration of an iron chelator resulted in more pronounced acute hypoxia-induced lung injury, whereas iron sucrose demonstrated a protective effect against this injury. The serum iron concentration analysis provided additional evidence supporting the efficacy of the iron chelator and iron sucrose modeling approaches (Fig. 2G).

Given the pivotal role of iron as a cofactor for HIF1α activation, the present study assessed HIF1α expression. Immunohistochemistry and immunofluorescence staining of the main pulmonary artery and alveolar tissues in rats demonstrated that acute hypoxia resulted in an increase in HIF1α expression. In addition, the iron chelator promoted an increase in HIF1α expression, whereas iron sucrose inhibited the increase in HIF1α expression under acute hypoxic conditions (Fig. 3A-E). Furthermore, alterations in the expression of SMA in the pulmonary vasculature and lung tissue in response to acute hypoxia were in accordance with the expression of HIF1α (Fig. 3B-E).

Given the pivotal role of iron chelators and hypoxia in triggering mitophagy, the current study conducted a detailed examination of this process. The results of transmission electron microscopy of the main pulmonary artery demonstrated that acute hypoxia resulted in mitochondrial swelling, accompanied by increased mitophagy (Fig. 4A). The results of immunofluorescence analysis of the pulmonary vasculature demonstrated that acute hypoxia resulted in elevated expression of PINK1, a pivotal protein for mitophagy (Fig. 4B and D); this was accompanied by an increase in the expression of the mitophagy protein LC3B-II. Furthermore, the results indicated that DFO promoted hypoxia-driven PINK1 and LC3B-II upregulation, whereas iron sucrose inhibited this response (Fig. 4B-E).

To further validate the role of iron in acute HPV, in vitro experiments were conducted to examine the effects of iron on the proliferation of human pulmonary artery endothelial cells. The Edu and CCK8 assay results demonstrated that the proliferation of human pulmonary artery endothelial cells was diminished under acute hypoxia (Fig. 5A-C). Furthermore, the administration of an iron chelator led to a further reduction in endothelial cell proliferation under hypoxic conditions, whereas the use of iron sucrose inhibited this reduction (Fig. 5A-C).

The present study also employed immunofluorescence to elucidate the effect of iron on the apoptosis of pulmonary artery endothelial cells under acute hypoxia. The findings demonstrated that acute hypoxia resulted in the upregulation of the apoptotic protein Caspase 3 and Annexin V-FITC (Fig. 5D-G), accompanied by a reduction in MMP (Fig. 5G). The administration of an iron chelator led to a further exacerbation of these hypoxia-induced abnormalities, whereas iron sucrose was observed to exert a protective effect, ameliorating the aforementioned acute hypoxia-induced abnormalities in pulmonary artery endothelial cells (Fig. 5D-G).

The expression of HIF1α was observed to be elevated in pulmonary artery endothelial cells subjected to acute hypoxia (Fig. 6A and B). The administration of an iron chelator was revealed to facilitate HIF1α expression, whereas the introduction of iron sucrose was demonstrated to impede HIF1α expression in the context of acute hypoxia. The results of western blot analysis of endothelial cells demonstrated that acute hypoxia induced an increase in LC3B-II expression, DFO promoted LC3B-II expression under acute hypoxia, whereas iron sucrose suppressed its upregulation (Fig. 6C and D).