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Exerc Sci > Volume 33(2); 2024 > Article
Hong and Shin: Prolonged Hypoxic Exposure Impairs Endothelial Functions: Possible Mechanism of HIF-1α Signaling



Hypoxic training enhances oxygen availability by elevating hemoglobin, red blood cells, and capillaries. hypoxia-inducible factor (HIF) stabilization under hypoxic conditions triggers glycolysis, erythropoiesis, and angiogenesis. However, prolonged hypoxic exposure causes endothelial cell dysfunction, contributing to cardiovascular diseases. Furthermore, endothelial mitochondria play a crucial role in maintaining endothelial cell homeostasis through the processes of biogenesis, signaling cellular response, mitochondrial dynamics, and calcium homeostasis. However, the molecular response of endothelial mitochondria has not been fully elucidated. Therefore, this study aimed to investigate the endothelial mitochondria functions and morphological change in response to hypoxia.


Cobalt chloride and 2% hypoxic condition were applied to HUVECs. The endothelial cell functions were assessed using BrdU-based proliferation assay and scratch wound healing assays. Mitochondrial functions were evaluated using Mitosox and Mito-Tracker staining for mitochondrial reactive oxygen species (mtROS) and mitochondrial morphology analysis, respectively.


The hypoxic condition (2% of O2) significantly decreased endothelial proliferation/migration and mitochondrial function. Furthermore, CoCl2 treatment increased the mtROS levels and mitochondrial fragmentation in HUVECs. Additionally, hypoxic conditions were found to regulate the phosphorylation of eNOS, mitochondrial biogenesis, mitophagy, and cell cycle-related protein expression.


These results suggest that the HIF-α pathway leads to endothelial dysfunction via mitochondrial dysfunction. Further mechanistic studies will be needed to elucidate the cellular and molecular mechanisms by which hypoxic-response pathways influence the health and homeostasis of endothelial cells. The findings reported herein underscore the importance of strategies for hypoxic training to prevent cardiovascular diseases.


The vascular endothelium is a single layer of endothelial cells lining blood vessels, which is in direct contact with blood, exchanging oxygen and nutrients to working tissues of the body for homeostasis. Furthermore, endothelial cells play a crucial role in regulating vascular tone, vascular permeability, angiogenesis, and maintaining immune response for cardiovascular homeostasis. Maintaining optimal oxygen concentration in endothelial cells is critical for sustaining endothelial homeostasis, otherwise, it accompanies many vascular complications, such as stroke and atherosclerosis.
Hypoxic exercise training, commonly referred to as altitude training, involves physical exercise performed under reduced oxygen conditions. Under low oxygen tension, hypoxia-inducible factor (HIF) subunits are stabilized by inactivated prolyl hydroxylase domains (PHDs), leading to ubiquitination of HIF-1α and HIF-2α via von Hippel-Lindau protein (VHL) E3 ubiquitin ligase [1]. Subsequently, these stabilized HIF subunits translocate into the nucleus to activate various target genes associated with hypoxic adaptation, including upregulation of glycolysis, erythropoiesis, and angiogenesis [2-5]. These adaptive responses to hypoxic training confer beneficial effects such as increased levels of hemoglobin, red blood cells, and capillaries, ultimately enhancing endurance capacity and cardiorespiratory fitness [6]. Our previous work demonstrated that endurance exercise training increases muscle fiber transition toward slow-twitch phenotype and skeletal muscle microvasculature with a stabilization of HIF-α in hypoxia-exposed PHD2-deficient mice compared to wild-type mice under normoxic condition [7,8]. However, acute or chronic hypoxic exposure shows a controversial response such as causing cellular stress. Indeed, hypoxia impairs skeletal muscle regen-eration and induces muscle fiber atrophy [9]. Additionally, our previous work demonstrated that mice subjected to 10% oxygen levels for 4 weeks showed reductions in body weight and skeletal muscle mass [8]. Furthermore, HIF-1α-pathway in endothelial cells plays a pivotal role in the angiogenesis process, while HIF-1α activation has been known to induce endothelial cell dysfunction linked to cardiovascular diseases, including hypertension, atherosclerosis, and heart failure [4,10-12]. Some studies revealed that chronic hypoxic exposure is associated with endothelial cell dysfunction; however, the precise molecular mechanisms of endothelial cell function and homeostasis upon hypoxic-responsive pathway remain incompletely understood. Specifically, endothelial mitochondrial phenotypic change is critical for maintaining endothelial cell function. However, it has not fully elucidated how hypoxia influences endothelial mitochondria phenotype. Consequently, the present study aims to elucidate the possibility that hypoxic training potentially occurs in cardiovascular diseases through the phenotypic change of endothelial cell mitochondria in the hypoxic response pathway. We assessed the endothelial and mitochondrial function and underlying hypoxic response pathway using cobalt chloride (CoCl2), widely employed chemical HIF-α activa-tor, and 2% actual hypoxic condition.


1. Cell culture

Human umbilical vein endothelial cells (HUVECs) were cultured with M199 basal media supplemented with 20% fetal bovine serum and endothelial cell growth supplement (Sigma-Aldrich, #E2759), and incubated at 37ºC under a humidified atmosphere containing 5% CO2. For hypoxic conditions, HUVECs were incubated in a hypoxia chamber maintained at 93% N, 2% O2, and 5% CO2. For experiments with cobalt chloride (CoCl2) treatment, HUVECs were treated with DMSO-vehicle control, or CoCl2 (0.6 mM, Sigma-Aldrich, #232696) for 24 hours. The HUVECs were cultured under normoxic and hypoxic conditions for 24 hours. HUVECs were used for experiments between passages 5 to 10 [13-16].

2. Mitosox staining

Mitochondrial ROS production was measured using the fluorescent probe Mitosox Red mitochondrial superoxide indicator for live-cell imaging (Invitrogen, #M36008). Cells were incubated with Mitosox working solution (5 μM) for 15 minutes at 37ºC. The cells were gently washed three times with pre-warmed PBS after the removal of the working solution. The images were collected using a fluorescence microscope (Axiom-ager, Zeiss or SP8 confocal microscope, Leica) with a 63X oil objective lens.

3. Mito-tacker red staining and mitochondrial morphology analysis

HUVECs were incubated with Mito-Tracker solution (200 nM, Molecular Probes, M22425) with growth media at 37°C for 30 minutes. After removal of the incubation solution, cells were washed three times with pre-warmed PBS, and then the PBS was replaced with pre-warmed growth media. Stained HUVECs were observed using a fluorescence microscope with a 63× oil objective lens (Zeiss). Mitochondrial morphology was examined by mitochondrial fission count (MFC) as previously de-scribed [3,17,18]. Briefly, acquired images of mitochondria were imported into Image J and subjected to background subtraction and kernel convolution [19] to identify mitochondria segments. The individual mitochondrial particles, including circularity and major/minor axes, were assessed using ImageJ software. Mitochondrial Fission Count (MFC) was calculated by the number of individual mitochondria per cell divided by the total mitochondria area of a given cell (MFC=mitochondrial number/total mitochondrial area). An elevation of the MFC value represents more mitochondria per given area, indicating increased fission. Form Factor (FF: the reciprocal of circularity value/(perimeter2/4π*area)) and aspect ratio (AR: major axis/minor axis of an ellipse equivalent to the object) were calculated.

4. Immunoblot

The cultured HUVECs were harvested with RIPA buffer (40 mM Tris (pH 7.5), 300 mM KCl, 1% Triton X-100, 0.5 M EDTA, Protease inhibitor (ThermoFisher, #87785)) and phosphatase inhibitor (Sigma-Aldrich, #04906845001), to extract total protein. The cell lysates were incubated on ice for 30 minutes and centrifuged at 16,000 g for 20 minutes at 4°C, and the supernatants were isolated. Protein concentrations were determined using the BCA protein assay kit (Thermo Fisher Scientific, Rock-ford, IL) with bovine serum albumin (BSA) as the standard. Total protein was separated via 8-12% SDS-PAGE and transferred to a PVDF membrane (Invitrogen). The membrane was blocked using Tris-buffered saline with 0.05 % Tween 20 (TBST) containing 3% BSA for 1 hour and incubated overnight with appropriately diluted (1:500-1,000) primary antibody in TBST at 4°C. After incubation, the membranes were rinsed three times in TBST for 5 minutes and incubated with secondary antibody in 5% skim milk for 1 hour at room temperature. Protein bands were visualized by SuperSignal West Femto or Dura Chemiluminescent Substrates.

5. Cell proliferation

The 5-bromo-2’-deoxyuridine (BrdU) assay was performed to evaluate proliferating cells. For the assay, culture media was removed and replaced with 10 µM of BrdU labeling solution, for incubation with the cells at 37°C for 2 hours. The labeled cells were washed with PBS and then fixed with 3.7% PFA for 15 minutes. The fixed cells were permeabilized and blocked with 5% normal goat-serum (NGS) in PBS. After blocking, the cells were incubated with BrdU primary antibody (Novus, # MAB7225) in blocking buffer (5% NGS in PBS) overnight at room temperature. The next day, samples were washed with PBS and incubated with a fluorescently labeled secondary antibody (Alexa fluor 488 goat Antimouse secondary antibody, Invitrogen A11001) for 1 hour at room temperature. After secondary antibody incubation, samples were mounted with a mounting solution containing DAPI. Cell proliferation was determined by the number of BrdU positive cells/total number of cells.

6. Scratch wound healing assay

HUVECs were seeded in 24-well plate for wound healing assay. The cell monolayer was scraped in a straight line to create a scratch using p200 pipet tips. To remove the cell debris, the wells were washed with warmed PBS, and then the media was replaced. The plate was then placed in 37°C incubator for 12 hours.

7. Statistics

Data are presented as mean±SEM. All probability values were calculated using a two-tailed distribution Student's t-test using Instant v3.06 software (Graph pad 9.0) and considered significant at p <.05.


1. Prolonged hypoxic condition aggravates endothelial proliferative and migratory capacities

To investigate endothelial functions under physiological hypoxic conditions, we performed the functional assay using endothelial cells exposed to a 2% hypoxic condition. Our findings revealed a suppression of endothelial cell proliferation in response to hypoxic exposure (Fig. 1A). Furthermore, analysis using scratch wound-healing assay demonstrated a marked inhibition of endothelial cell migratory ability under hypoxic conditions compared to normoxic conditions (Fig. 1B).
Fig. 1.
Fig. 1.
Hypoxia suppresses endothelial cell proliferation and migration. (A) Representative images of BrdU (bromodeoxyuridine) proliferation of HUVECs under normoxia (Nx) and hypoxia (Hx). (B) Quantification of BrdU positive cells/total cells (n=7-8). Scale bar=20 μm. (C) Representative images of scratch assay. (D) Quantification of percentage of wound closure at 12 hours-time point (n=7). Data are shown as mean±SEM. *** p<.001, **** p<.0001.
Next, we examined the protein expression related to nitric oxide synthase, cell cycle regulation, and mitochondrial biogenesis under hypoxic conditions. Hypoxia induced a significant increase in HIF-2α, PHD2, and p16 contributing to the deceleration of cell cycle progression. Conversely, hypoxic conditions downregulated the expression of mitochondrial biogenesis markers including TFAM and SIRT1, as well as the phosphorylation of eNOS (Fig. 2A and B). Taken together, our findings suggest that hypoxic exposure induces endothelial cell dysfunction mediated by HIF-α signaling pathways.
Fig. 2.
Fig. 2.
Hypoxic condition aggravates endothelial functions. (A) Representative western blot images showing protein levels from HUVECs under normoxia (Nx) or hypoxia (Hx). (B) Densitometry quantification of western blots (n=7). Data are shown as mean±SEM. * p<0.05, ** p<.01, *** p<.001, **** p<.0001.

2. Prolonged cobalt chloride induces endothelial mitochondrial dysfunctions

To elucidate the involvement of HIF-1α in endothelial cell functions, we utilized cobalt chloride (CoCl2), a chemical agent known to stabilize HIF-1α, thereby inducing a mimetic hypoxic condition [20]. Our investigation revealed a notable increase in mitochondrial superoxide production in endothelial cells following CoCl2 treatment (Fig. 3A and B). We confirmed that CoCl2 treatment led to significant upregulation in the protein expression of HIF-1α and PHD2, which is an oxygen sensor and one of the target genes of HIF-1α, and AMPKα, a key regulator of metabolic enzyme activity. Additionally, we assessed the protein expression related to autophagy, antioxidative defense, and apoptosis. CoCl2 treatment elevated the levels of the antioxidant manganese superoxide dismutase (MnSOD), autophagic marker p62, apoptotic regulators Caspase-3, and BNIP3 and resulted in a significant reduction in the phosphorylation of endothelial nitric oxide synthesis (p-eNOS) (Fig. 3C and D).
Fig. 3.
Fig. 3.
Cobalt chloride treatment increases mtROS production and mitochondrial fragmentation in HUVECs. (A) Representative micrographs of MitoSOX staining. mtROS production determined by Mitosox™ staining under vehicle control or CoCl2 treatment (24 hours) (n=3-4). Scale bar=20 μm. 1:630 magnification. (B) Quantification of fluorescence intensity (n=3-4). (C) Representative western blot images showing protein levels from HUVECs treated CoCl2 and vehicle control. (D) Densitometry quantification of western blots (n=3). Data are shown as mean±SEM. * p<.05, ** p<.01, *** p<.001.
Furthermore, we analyzed mitochondrial morphology to corroborate the impact of CoCl2-induced HIF-1α stabilization on endothelial cells. Quantitative morphometrics parameters such as aspect ratio (AR), form factor (FF), and mitochondrial fragmentation count (MFC), demonstrated that CoCl2 induced a shift towards fragmented mitochondrial morphology compared to the vehicle control (Fig. 4A and B). Prolonged cobalt chloride treatment adversely affects mitochondrial morphological changes. Additionally, CoCl2 induced the phosphorylation of Drp1, a mitochondrial fission protein, and decrease in mitochondrial fusion protein 2 (MFN2). Conversely, CoCl2 treatment as well as decreased expression of mitochondrial biogenesis markers (TFAM, CHCHD4) (Fig. 4C and D). Collectively, HIF-1α-induced by CoCl2 treatment aggravated endothelial mitochondrial functions and activated cell death mechanisms such as autophagy and apoptosis.
Fig. 4.
Fig. 4.
Cobalt chloride upregulates autophagy and apoptosis regulatory proteins and downregulates mitochondrial biogenesis markers. (A) Micrographs of endothelial mitochondria under vehicle control or CoCl2 treatment (24 hours) (n=4). Scale bar=20 μm. 1:630 magnification. (B) Quantification plots of mitochondrial morphology analyses (AR and FF) under vehicle control or CoCl2 treatment. MFC was calculated as number of particles/total area of mitochondria (n=4). (C) Representative western blot images showing protein levels from HUVECs treated CoCl2 and vehicle control. (D) Densitometry quantification of western blots (n=3). Data are shown as mean±SEM. * p<.05, ** p<.01, *** p<.001.


Our work elucidates the impact of HIF-α signaling under both cobalt chloride-induced chemical hypoxic and actual hypoxic conditions on endothelial mitochondrial function in vitro. We have demonstrated that the HIF-α stabilization causes increased mitochondrial ROS production and fragmentation, accompanied by a decline in endothelial function. These findings enhance our understanding of the molecular mechanisms that unfold in endothelial cells when exposed to hypoxic stimuli.
Chronic hypoxic exposure is implicated in the development of cardiovascular diseases, including hypertension and atherosclerosis [10-12]. Specifically, endothelial-specific depletion of PHD2 led to vascular fibrosis, elevated blood pressure, increased arterial thickness, and leading pulmonary arterial remodeling in the HIF-α-dependent pathway [21,22]. Notably, our current study revealed that while the requirement of oxygen for mitochondrial ROS generation at complex I and III of the electron transport chain during oxidative phosphorylation, the inhibition of mitochondrial biogenesis by HIF-1α activation paradoxically increases mitochondrial ROS production [23]. Many studies have shown that hypoxia or HIF-1α is involved in mitochondrial ROS generation; however, further studies are required to elucidate the precise molecular mechanisms by which HIF-1α induces mitochondrial ROS production.
Furthermore, we have demonstrated that HIF-1α disrupts the equilibrium of mitochondrial dynamics in endothelial cells. Mitochondria are dynamic organelles, continuously undergoing the cycle of fusion and fission processes to maintain mitochondrial homeostasis and quality [24]. The imbalance of mitochondrial dynamics is strongly associated with endothelial cell dysfunctions [24]. Some studies have shown that hypoxic stimuli upregulate mitochondrial fission-related proteins such as DRP1 and FIS1, through the action of a-kinase anchoring protein 121 (AKAP121), a scaffold protein located on the mitochondrial membrane. The AKAP121 promotes DRP1 phosphorylation and DRP1, in turn, fa-cilitates interaction between DRP1 and FIS1 [25,26]. Additionally, Chen group has demonstrated that HIF-1α directly modulates DRP1 expression via binding to a hypoxia-responsive element within the promoter of the DRP1 gene [27,28].
Interestingly, our observation revealed that CoCl2-treated HUVECs showed elevated levels of mtROS, despite an increase in manganese-dependent superoxide dismutase (MnSOD), an enzyme responsible for mitigating mitochondrial ROS. Peter group reported that the upregulation of MnSOD inhibits mitochondrial oxidative metabolism and contributes to the exacerbation of mitochondrial biogenesis to activate the glycolysis process via mitochondrial hydrogen peroxide (mtH2 O2) production and AMPK activation [29]. Thus, it is plausible that HIF-1α-mediated MnSOD may contribute to the suppression of mitochondrial oxidative metabolism rather than the reduction of mtROS levels.
Furthermore, we observed that ECs have shown the reduction of proliferation and migration under actual hypoxic condition, despite glycolysis being a primary source for EC proliferation and migration, rather than oxidative phosphorylation via mitochondria [24]. However, mitochondrial function, specifically mitochondrial respiratory chain activation at complex I and maintenance of NAD+/NADH ratio at the complex III, are critical as the metabolic intermediates utilized as building blocks for biosynthetic macromolecules through the mitochondrial TCA cycle are necessary for EC proliferation [30].
While our observation did not provide direct evidence that hypoxic condition impairs mitochondrial respiration and TCA cycle, we confirmed that hypoxia inhibited the expression of TFAM and SIRT1, pivotal factors for mitochondrial biogenesis, respiratory capacity and TCA cycle [31,32].
Furthermore, hypoxic exposure suppressed EC migratory capacity. One possible explanation for this finding is that hypoxia suppressed eNOS phosphorylation and increased p16, negatively regulating the cell cycle. A previous study demonstrated that the inhibition of eNOS impairs endothelial migration through the cell adhesion molecules such as integrin αvβ3, VCAM-1, ICAM-1, and gap junction protein connexin 43 which are associated with endothelial cell angiogenesis [33].


The current study provides evidence that HIF-α pathway leads to endothelial dysfunction via mitochondrial dysfunctions. Further mechanistic studies are needed to elucidate the cellular and molecular mechanisms by which hypoxic-response pathway influence vascular cells health and homeostasis. Consequently, our finding underscores the importance of strategies for hypoxic training to prevent cardiovascular diseases.

Conflict of Interest

The authors declare that they have no competing interests.


Conceptualization: J Shin; Data curation: J Shin; Formal analysis: J Hong, J Shin; Funding acquisition: J Shin; Project administration: J Hong; Writing - original draft: J Hong, J Shin; Writing - review & editing: J Hong, J Shin.


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