R213G‐Mediated Redistribution of EC‐SOD Protects Against Sugen‐Hypoxia Pulmonary Hypertension in Mice

ABSTRACT

Pulmonary hypertension (PH) is a progressive and life‐threatening disease characterized by pulmonary vascular remodeling that leads to elevated pulmonary artery pressures, and subsequent right ventricular dysfunction. Despite advances in understanding PH pathogenesis, treatment options remain limited, underscoring the need to define novel mechanisms that contribute to disease progression. Inflammation and oxidative stress are recognized drivers of pulmonary vascular injury, with extracellular superoxide dismutase (EC‐SOD or SOD3), a matrix‐bound antioxidant enzyme, playing a role in multiple lung and vascular pathologies. A common human single nucleotide polymorphism (rs1799895) in the SOD3 gene leads to an arginine to glycine amino acid substitution (R213G) and alters EC‐SOD localization by reducing its affinity for the extracellular matrix, resulting in increased circulating but decreased lung EC‐SOD content. Using a murine knock‐in model of the R213G variant, we have previously demonstrated exacerbated chronic hypoxia‐induced PH. In this study, we examined the impact of EC‐SOD redistribution due to the R213G SOD3 variant on the development of PH in the Sugen‐hypoxia (SuHx) model, a more severe model of PH. We hypothesized that R213G mice would also have exaggerated hemodynamic changes, vascular remodeling, and inflammatory changes in this model. However, while SuHx increased pulmonary artery pressure and vascular remodeling in wild‐type mice, the R213G variant unexpectedly attenuated SuHx‐induced PH. Following SuHx, the increase in pulmonary artery pressures was attenuated in R213G mice. Early immune profiling revealed that SuHx triggered significant neutrophil and interstitial macrophage infiltration in the lungs of wild‐type mice, which was markedly blunted in R213G mice. These findings suggest that the redistribution of EC‐SOD into the extracellular fluid, though it lowed lung EC‐SOD levels, attenuated early inflammatory responses and protected against the development of PH in SuHx. This work highlights a novel, compartment‐specific role for EC‐SOD in modulating immune‐driven mechanisms of PH and may inform future therapeutic strategies targeting oxidative stress and inflammation in vascular disease.

1. Introduction

Pulmonary hypertension (PH) is a progressive and life‐threatening condition characterized by pulmonary vascular remodeling and elevated pulmonary artery pressures that ultimately result in right ventricular failure and premature death [1, 2]. Despite scientific advances that elucidate key disease mechanisms, current treatments offer limited efficacy, primarily targeting vasoconstriction rather than halting or reversing disease progression [1, 2, 3]. This highlights the critical need to unravel the molecular and cellular mechanisms driving PH pathogenesis to identify novel therapeutic targets. Dysregulated redox signaling, which can be augmented by a disruption in antioxidant enzyme levels, plays a crucial role in promoting vascular dysfunction, inflammation, and remodeling, all hallmark features of PH [4, 5, 6].

Extracellular superoxide dismutase (EC‐SOD or SOD3) is a key antioxidant enzyme that catalyzes the dismutation of superoxide into hydrogen peroxide and oxygen, thereby mitigating vascular oxidative stress [7, 8, 9, 10]. EC‐SOD is predominantly secreted by vascular smooth muscle cells and binds to the extracellular matrix (ECM) via its heparin‐binding domain (HBD), providing site‐specific protection against reactive oxygen species (ROS)‐mediated vascular injury [7, 8, 11]. In the context of PH, in both murine and human studies, EC‐SOD activity is reduced, and lower levels of lung EC‐SOD have been associated with worse PH [10, 12, 13, 14]. Our lab has become particularly interested in a naturally occurring single‐nucleotide polymorphism (SNP) in the SOD3 gene (R213G) that results in reduced ECM binding, leading to increased EC‐SOD in the bronchoalveolar (BAL) fluid and circulation and reduction of EC‐SOD in the lung and vessel wall, without changes in its enzymatic activity [15, 16, 17]. We have demonstrated that mice expressing a knock‐in of the R213G variant of SOD3 (R213G mice) develop exacerbated chronic hypoxic PH, suggesting that the localization of EC‐SOD is important in disease pathogenesis [18].

In this study, we hypothesized that the loss of vascular EC‐SOD in R213G mice would exacerbate PH in a well‐described model of PH induced by concurrent treatment with the VEGF receptor antagonist, Sugen, and hypoxia (SuHx). This model has been utilized in both mice and rats by numerous groups because it closely mimics the severe pulmonary vascular remodeling observed in human PAH, including the presence of angio‐obliterative vascular lesions and right ventricular hypertrophy, providing a robust platform for studying disease mechanisms and potential therapeutic interventions [19, 20]. We thus investigated the impact of the human variant of SOD3 characterized by the redistribution from the ECM to the circulation on PH development in the SuHx model.

2. Methods

2.1. Mouse Model

Animal studies were conducted in accordance with ethical standards and were approved by the University of Colorado Institutional Animal Care and Use Committee (IACUC). Wild ‐type (C67BL/6) mice (WT) and homozygous mice expressing the R213G variant of EC‐SOD (rs1799895) on the C57BL/6 background (R213G) were used. The R213G knock‐in mice used in this study harbor the human rs1799895 single nucleotide polymorphism in the endogenous murine Sod3 gene, resulting in an arginine‐to‐glycine substitution at position 213 (R213G) within the heparin‐binding domain of EC‐SOD. This global knock‐in model has been previously described and validated. Mice were genotyped by PCR amplification of genomic DNA using allele‐specific primers (Forward: 5′‐AGGCTCAAGTCTGTGCCAGAAGG‐3′, Reverse: 5′‐TTTCCAATTCATTCACACACATGGG‐3′) that distinguish wild‐type alleles (357 bp amplicon) from R213G knock‐in alleles (419 bp amplicon) as previously described [18]. The R213G mutation results in ubiquitous expression of the variant EC‐SOD protein in all tissues, as the modification was introduced into the endogenous Sod3 locus [18]. Wild‐type littermates served as controls and were genotyped using the same strategy. Mice were assigned to either receiving Sugen‐hypoxia (SuHx) or DMSO‐normoxia. For the SuHx treatment, male and female WT and R213G mice age 8–12 weeks received intraperitoneal injections of Sugen5416 (20 mg/kg in dimethylsulfoxide (DMSO), Cayman Chemical) once weekly for 3 weeks while concurrently being maintained in hypobaric hypoxia chambers (395 Torr; to simulate 10% FiO₂). The study was not powered to detect sex‐specific differences. Therefore, data were pooled across sexes. At the end of the 3‐week period, the mice were removed from the hypobaric hypoxic chambers and maintained at Denver ambient air (1609 m, normoxia) for an additional week. The control group received once‐weekly intraperitoneal DMSO injections for 3 weeks and was maintained in normoxia throughout the exposure period. For experiments designed to investigate peak inflammation, which occurs early in the chronic hypoxia model, mice were analyzed at 4 days post Sugen‐hypoxia (early time point) [21].

2.2. Hemodynamics

At the culmination of the 4‐week exposure period, anesthetized mice underwent right heart catheterization (RHC). Anesthesia was induced using 2%–5% isoflurane to ensure a controlled state of unconsciousness. Employing an open‐chest technique, a catheter was sequentially introduced into the right ventricle, pulmonary artery, and left ventricle, and data collected to calculate pressure‐volume loops and cardiac function, including pulmonary artery pressures, cardiac output, ejection fraction, pulmonary vascular resistance, systemic vascular resistance, afterload, end‐diastolic pressure‐volume relationship, end‐systolic pressure‐volume relationship (specifically, Ees as a metric of contractility), and ventricular–arterial coupling (quantified as Ea/Ees). Right ventricular hypertrophy was assessed by Fulton index‐ (left ventricle plus septum (RV/(LV + S)).

2.3. Blood/Plasma Collection and Tissue Harvesting

In anesthetized mice, prior to tissue harvesting (early timepoint) or following hemodynamics (late timepoint), blood was obtained from the right ventricle using a 21‐gauge needle and heparin coated syringes. An aliquot of blood was used for complete blood counts (CBC) using the hematologic analyzer Heska HT5 (Loveland, CO). Plasma was isolated from heparinized whole blood by centrifugation at 2000g x 5 min. Plasma samples were stored at −80°C until use. Mice were then euthanized for tissue harvesting. Mouse lungs were flushed via the right ventricle with 10 mL of cold PBS and flash frozen or prepared for paraffin‐embedding and immunohistochemistry. For lung slides, lungs were inflation‐fixed at 20 cm H₂O for 30 min with 4% paraformaldehyde then placed in 4% paraformaldehyde for 24 h before transfer to 70% ethanol and tissue sectioning. Hearts were stored in 70% alcohol prior to dissection for RV and LV septal weights to calculate Fulton's index.

2.4. Protein Isolation

Lung homogenates were prepared from 30 mg of flash frozen lung tissue using lysis buffer containing T‐PER (Thermo Fisher Scientific, Waltham, MA, USA), along with protease inhibitor cocktail (Sigma‐Aldrich) and phosphatase inhibitor cocktails 2 and 3 (Sigma‐Aldrich, Burlington, VT, USA). Tissue was disrupted using the Bead Ruptor 12 (Omni International, Kennesaw, GA, USA) at high speed for 45 s, followed by cooling on ice for three cycles. After homogenization, the samples were incubated on ice for 30 min, then centrifuged at 10,000 × g for 5 min. Protein samples were stored at −80°C until further use.

2.5. Western Blot

The protein concentration was assessed utilizing the Pierce Rapid Gold BCA protein assay kit (Thermo Fisher Scientific). For lung homogenate samples, 30 µg of protein were loaded per well and then separated by gel electrophoresis using Criterion XT 4–12% Bis‐Tris Precast Gel (BioRad) with XT MES running buffer (Bio‐Rad). For serum samples, 4 µL were loaded per well. Using a Trans‐Blot Turbo rapid transfer system (Bio‐Rad), proteins were transferred from the gel to 0.2 µm polyvinylidene fluoride membranes (Bio‐Rad) which were activated using methanol. Membranes were then blocked with 5% nonfat dry milk in Tris‐buffered saline containing 0.05% Tween 20 (TBST) for 1 h prior to incubation with primary antibodies: EC‐SOD goat IgG anti‐mouse (R&D System) at 1:1000 in 5% milk in TBST at 4°C overnight and β‐actin mouse monoclonal (Sigma‐Aldrich) at 1:10,000 in 5% milk in TBST at room temperature for 1 h. Subsequently, the appropriate horseradish peroxidase‐conjugated anti‐goat or anti‐mouse secondary antibody (R&D System) was applied at 1:10,000 in TBST for 1 h at room temperature. Band signal intensity was determined using the SuperSignal Femto Chemiluminescent Substrates (Thermo Fisher Scientific) with quantified of band intensity via densitometry with Image Laboratory Software (Bio‐Rad). The signals were normalized to the WT DMSO‐normoxia.

2.6. Immunohistochemistry Analysis of Small Vessel Muscularization

Five‐micron lung sections were evaluated for muscularization of small vessels by immunohistochemical staining for alpha‐smooth muscle actin (SMA) using polyclonal antibody anti mouse αSMA (Sigma A2547) at 1:1200 dilution and Dako EnVision + Dual Link detection system per the manufacturer's instructions. Small vessel muscularization was determined by counting the number of small vessels (< 30 µm) with greater than 70% staining for αSMA in 6–10 different randomly selected high power field sections. Number of cells with positive staining was counted in 6–10 different randomly selected high power field sections. Two blinded investigators analyzed the slides and the data were averaged for each mouse.

2.7. ELISA

IL‐6 concentrations were measured in 50 µL of undiluted plasma using mouse‐specific ELISA per instructions (R&D Systems, Minneapolis, MN). Samples were run in duplicate and averaged and fit to a standard curve.

2.8. Flow Cytometry

Intra‐vascular leukocytes were labeled by a retro‐orbital (RO) injection of 1 μg CD45‐PE antibody (BD Biosciences, Clone No. 30‐F11) in 100 μL PBS 5 min before collecting blood and lungs. Following blood collection via open chest cardiac puncture, lungs were flushed with 10 mL PBS (without calcium or magnesium; Sigma) via the right ventricle to dislodge the majority of circulating leukocytes, then isolated. Lungs were processed for flow cytometric analysis and sorting as previously described [21, 22]. In brief, collected whole lungs were digested in 1 mL of HBSS (without calcium or magnesium; Sigma) with Liberase TM (Sigma, 0.4 mg/mL), and DNAse I (Sigma, 100 U/mL) utilizing mechanical digestion with the GentleMACS system (preinstalled program m_lung_01_02 followed by 15 min 37°C incubation with agitation and dissociation using the m_lung_02_01 program for 8 s). After filtering the cell digest through a 70 μm cell strainer, red blood cells were lysed (eBioscience 1X RBC Lysis Buffer, 3 min, room temperature), and cell suspensions were centrifuged (10 min, 300 g), washed, and incubated in FA3 buffer (PBS, 1 mM EDTA, 10 mM HEPES, 1% FBS, pH 7.4) at 4°C. For staining, FCγR blocking was first performed with anti‐CD16 and anti‐CD32 antibody (BD Biosciences) for at least 20 min. Cell suspensions were then stained with LIVE DEAD Fixable Blue (Thermofisher) for 30 min according to the manufacturer's instructions, followed by 30 min incubation with the antibody panel. Antibody details: anti‐mouse CD45‐AF700 (BioLegend, Clone No. 3—F11, 1:100), CD64‐AF647 (BD Pharminogen, Clone No. X54‐5/7.1, 1:100), CD11b‐FITC (BioLegend, Clone No. M1/70; 1:100), CD11c‐BV421 (BioLegend, Clone No. N418, 1:100), MHCII‐APC‐Cy7 (Clone No. M5/114.15.2, 1:100), Ly6G‐BV510 (BioLegend, Clone No. 1A8, 1:100), and Lyve1‐PE‐Cy7 (Invitrogen, Clone No. ALY7, 1:40). In a separate panel, CD3‐BV421 (BioLegend, Clone No. 17A2, 1:100), B220‐FITC (BD Biosciences, Clone No. RA3‐6B2, 1:100) were used for identification of T and B cell populations from the IV‐CD45+ population. Cell suspensions were then washed and fixed (10% PFA, 30 min 4°C), filtered through flow tubes with 40 µm filter caps (Falcon) and run on the NovoCyte Penteon Analyzer (Agilent) within the University of Colorado AMC Cancer Center Flow Cytometry Shared Resource Core Facility. All cell populations were gated using fluorescence minus one (FMO) controls and absolute counts were calculated using 123 counting beads (Invitrogen, Waltham, MA). Data were analyzed using NovoExpress analysis software (Agilent).

2.9. Statistics

Data analysis was conducted using Prism 10 software (GraphPad; La Jolla, CA, USA). For all experiments, both male and female mice were included with 6–10 mice per group to achieve 80% power for detecting any significant effect. For comparisons between two groups, a Student's t‐test was used. For comparisons involving three or more groups, two‐way ANOVA with Tukey's post hoc test was applied.

2.10. Results

2.10.1. In R213G Mice, SuHx Did Not Alter Baseline Lung EC‐SOD Levels but Further Increased Serum EC‐SOD

The EC‐SOD R213G variant reduces matrix binding affinity, resulting in low lung content and elevated circulating EC‐SOD levels [15, 16, 17]. Furthermore, the R213G variant worsens cardiovascular disease outcomes in humans and chronic hypoxic PH in mouse models [18]. To evaluate how the R213G variant impacts EC‐SOD content following SuHx, we measured lung and serum EC‐SOD protein expression in WT and R213G mice at baseline and at 35 days following SuHx. In the lung, EC‐SOD content is significantly reduced in R213G mice compared to WT under both normoxic control (DMSO) and SuHx conditions (Figure 1A, p < 0.05). Lung EC‐SOD levels did not change from baseline in response to SuHx in either genotype. Serum EC‐SOD content is significantly elevated in R213G mice compared to WT under both normoxic and SuHx conditions (Figure 1B, p < 0.05). In contrast to the lung, serum EC‐SOD content increased from baseline following the SuHx exposure in the R213G mice, but not in the WT mice (Figure 1B, p < 0.001). Together, these findings confirm that the R213G polymorphism alters the compartmentalization of EC‐SOD at baseline, with reduced lung content and increased serum levels compared to WT EC‐SOD, and serum levels increase SuHx only in the R213G strain.

FIGURE 1.

SuHx enhances serum EC‐SOD in the R213G mice though lung levels remain low. WT and R13G mice were treated with weekly doses of IP Sugen 5416 (10 mg/kg) concurrent with hypobaric hypoxia (395 torr, equivalent to 10% oxygen) for 3 weeks followed by normoxia for 1 additional week. Normoxic mice received an equivalent volume of vehicle, DMSO. (A) Relative densitometry for lung EC‐SOD protein relative to beta‐actin normalized to WT normoxia with representative immunoblot images. (B) Relative densitometry for serum EC‐SOD protein in 4 µL of serum normalized to WT normoxia with representative immunoblot images. N = 9, *p < 0.05, **p < −0.01, *** p < 0.005, ****p < 0.001 by two‐way ANOVA. b‐actin, beta actin; EC‐SOD, extracellular superoxide dismutase; Nx, normoxia and DMSO; SuHx, Sugen‐hypoxia; WT, wild‐type mice; R213G, mice with the R213G polymorphism.

2.11. SuHx Induced Pulmonary Hypertension Was Attenuated in R213G Mice

We previously reported that mice expressing the EC‐SOD R213G variant develop worsened chronic hypoxic PH and RV hypertrophy [18]. To examine whether this strain worsened PH in a second robust model of PH compared to WT mice, we evaluated established endpoints of PH in WT and R213G mice in response to SuHx. In WT mice, SuHx significantly increases mPAP compared to normoxic (DMSO) controls (Figure 2A, p < 0.001). However, contrary to our hypothesis, the SuHx‐induced increase in mPAP was attenuated in EC‐SOD R213G variant mice compared to WT (Figure 2A, p < 0.005). SuHx induced right ventricular hypertrophy (RV/LV + S) in both groups, and was not worse in the EC‐SOD R213G variant mice (Figure 2B, p < 0.05). To assess vascular remodeling, we quantified the number of small muscularized vessels (< 30 µm, αSMA+ after completion of the model) in the lung by immunohistochemistry. SuHx significantly increased muscularized vessel counts in both WT and R213G mice with no differences between strains (Figure 3A–E).

FIGURE 2.

The R213G polymorphism attenuated SuHx‐induced PH. WT and R13G mice were treated with weekly doses of IP Sugen 5416 (20 m/kg) concurrent with hypobaric hypoxia (395 torr, equivalent to 10% oxygen) for 3 weeks followed by normoxia for 1 additional week. Normoxic mice received an equivalent dose of vehicle, DMSO. (A) Mean pulmonary artery pressure was measured via right heart catheterization in Normoxic and SuHx WT and R213G mice. N = 7–8, *p < 0.05, two‐way ANOVA. (B) Right ventricular hypertrophy (RVH) was assessed by the ratio of RV weight:LV + Septum weight (Fulton index) in both mouse strains in normoxia and SuHx. N = 8–9, **p < −0.01, *** p < 0.005, ****p < 0.001 by two‐way ANOVA. mPAP, mean pulmonary artery pressure; RV/(LV + S), right ventricle/(left ventricle/septum); SuHx, Sugen‐hypoxia; WT, wild‐type mice; R213G, mice with the R213G polymorphism.

FIGURE 3.

The R213G polymorphism did not attenuate SuHx‐induced muscularization. WT and R13G mice were treated with weekly doses of IP Sugen 5416 (20 m/kg) concurrent with hypobaric hypoxia (395 torr, equivalent to 10% oxygen) for 3 weeks followed by normoxia for 1 additional week. Normoxic mice received an equivalent dose of vehicle, DMSO. (A) Small vessel muscularization was assessed by α‐SMA staining in vessels < 30 µm. Representative image of staining of α‐SMA (brown) for muscularized vessels in WT (B) and R213G (C) mice treated in normoxia and WT (D) and R213G (E) mice treated with SuHx. Black arrows are non‐muscularized vessels < 30 μm, scale bar is 100 μm, ×20, N = 5–13, ***p < 0.005, ****p < 0.001 by two‐way ANOVA. WT, wild‐type mice; R213G, mice with the R213G polymorphism; SuHx, Sugen‐hypoxia; #Musc vessel/hpf, number of muscularized vessels per high power field.

2.12. Hematologic Indices in WT and R213G Mice Following SuHx Exposure

It is well established that hemoglobin levels increase in response to chronic hypoxia [23, 24]. To examine how the EC‐SOD R213G variant impacts hemoglobin and circulating cells in response to SuHx, we measured hematologic indices, including hemoglobin (Hb) levels, as well as white blood cell (WBC) counts and differential, and platelets at 4 days after initiation of SuHx. Hb increased in WT mice following SuHx (Figure 4A, p < 0.001) similar to previously published studies in chronic hypoxia [25]. Interestingly, Hb levels were significantly lower in R213G mice compared to WT following SuHx (Figure 4A, p < 0.05). WBC counts and lymphocyte counts tended to increase in SuHx‐exposed groups, though the difference was not statistically significant. Neutrophils, monocytes and platelets did not differ among strains or treatments (Figure 4C,D,F).

FIGURE 4.

The R213G EC‐SOD variant attenuates the increase in hemoglobin under SuHx while but did not significantly impact white blood cell and platelet counts. Blood was obtained from WT and R213G mice under normoxia and a 4‐day SuHx conditions. Using the Heska HT5 hematologic analyzer we report complete blood counts, including hemoglobin (A), white blood cell and differential (B–E), as well as platelet (F) counts. N = 4–6, *p < 0.05, **p < −0.01, ****p < 0.001, two‐way ANOVA. WT, wild‐type mice; R213G, mice with the R213G polymorphism; (4D) SuHx, 4‐day Sugen‐hypoxia; WBC, white blood cell.

2.13. SuHx Induced Accumulation of Lung Neutrophil and Interstitial Macrophage Accumulation and Increase in Circulating IL‐6 Is Attenuated in R213G Mice

Exposure of mice to hypoxia results in an early and transient increase in accumulation of immune cells, in particular interstitial macrophages, in the lung [26], therefore, we evaluated the impact of the R213G SNP on immune cell infiltration in the SuHx at an early 4‐day timepoint. Total neutrophil counts significantly increased in WT mice under SuHx but are attenuated in R213G mice (Figure 5A, p < 0.01). Total and resident alveolar macrophage counts were not significantly different at baseline in the normoxic R213G mice compared to WT (Figure 5B,C). The total macrophage counts were unaffected by SuHx in WT and R213G mice. Similar to hypoxia alone, total interstitial macrophages (IMs) significantly increased in both WT and R213G mice at 4 days SuHx (Figure 5D, p < 0.05). The increase in IMs was blunted in R213G mice (Figure 5D, p < 0.05). We then examined IM subsets. We observed that the IM1 significantly increased in response to SuHx in WT mice but not in R213G mice and was significantly lower in SuHx than WT mice (Figure 5E, p < 0.05). IM2 macrophage subsets significantly increased in WT mice but not in R213G in response to SuHx (Figure 5F, p < 0.06). IM3 did not increase in either WT or R213G mice with SuHx (Figure 5F). Finally, plasma IL‐6 levels were assessed under Sugen‐hypoxia (SuHx) conditions as a key inflammatory cytokine. Plasma IL‐6 levels increased in response to SuHx in WT mice (p < 0.05). In contrast, IL‐6 levels did not increase in R213G mice following SuHx (Figure 5H).

FIGURE 5.

The R213G polymorphism attenuates SuHx induced interstitial macrophage infiltration in the lung and reduces circulating IL‐6 levels. Flow cytometry analysis was performed on lung tissue from WT and R213G mice under normoxia and 4‐day Sugen‐hypoxia (4D SuHx) conditions to assess immune cell infiltration, including: total neutrophil (A), total macrophage (B), along with resident alveolar macrophages (C), total interstitial macrophages (D). Subpopulations of interstitial macrophages (E–G) were also analyzed by flow cytometry by evaluating the expression of CD11c and MHCII, for example, IM1 = CD11c^lowMHCII^low, IM2 = CD11c^lowMHCII^hi, and IM3 = CD11c^intMHCII^hi (E–G). (H) ELISA was performed to measure circulating IL‐6 levels in WT and R213G mice. N = 7–9, *p < 0.05, **p < −0.01, *** p < 0.005, ****p < 0.001 by two‐way ANOVA. AM, alveolar macrophages; IM1, interstitial macrophage subset 1; IM2, interstitial macrophage subset 2; IM3, interstitial macrophages subset 3; IL‐6, interleukin‐6; IM, interstitial macrophages; R213G, mice with the R213G polymorphism; (4D) SuHx, 4‐day Sugen‐hypoxia; WT, wild‐type mice.

3. Discussion

We and others have previously demonstrated that loss of EC‐SOD exacerbates hemodynamic changes in chronic hypoxic pulmonary hypertension (CHPH), while overexpression of lung EC‐SOD is protective in this model [10, 14, 27, 28]. Notably, the EC‐SOD R213G variant has been studied in chronic hypoxia, where it is shown to exacerbate PH severity [18, 29]. Building on these findings, we hypothesized that this variant, which decreases vascular EC‐SOD by reducing its matrix binding, would similarly worsen PH outcomes and inflammatory infiltration in a more severe model induced by concurrent treatment with Sugen and hypoxia (SuHx). Contrary to this hypothesis, our study shows that increased circulating EC‐SOD due to the R213G variant attenuates SuHx‐induced hemodynamic changes despite reduced EC‐SOD content in the lung and vessel wall. This protection is accompanied by a reduction in lung interstitial macrophage accumulation, particularly in the IM1 and IM2 subsets, as well as decreased circulating IL‐6 levels. These findings provide important insights into the compartment‐specific role of EC‐SOD in modulating early inflammatory responses in PH and underscore key differences between chronic hypoxia and SuHx models of hypoxia‐induced PH.

Several striking findings emerge from this study. First, despite reduced lung and vascular EC‐SOD content, the R213G variant attenuates SuHx‐induced PH, directly contrasting with prior reports showing worsened disease in chronic hypoxia and in human cardiovascular pathology associated with this variant [15, 29]. These findings challenge the premise that low vascular EC‐SOD alone drives PH severity. The discrepant response to chronic hypoxia vs. SuHx supports the concept that different pathways are responsible for driving PH and vascular remodeling in these two models, and implicates a benefit of the elevated circulating EC‐SOD in the R213G mice in SuHx‐induced PH. Since Sugen, as a VEGF receptor antagonist, targets pulmonary endothelial cells, we speculate that the endothelial cells are protected in the R213G mice either due to low vascular EC‐SOD or high circulating EC‐SOD. Prior studies have shown that loss of EC‐SOD reduces baseline lung VEGFR2 expression, so it is possible that reduced VEGF receptor expression at baseline may limit Sugen efficacy, although this remains to be directly tested [14]. Interestingly, we also demonstrated that the R213G mouse was protected against bleomycin‐induced lung fibrosis and subsequent PH [16]. In this model, the increase in alveolar EC‐SOD promoted apoptosis of the recruited alveolar macrophage however SuHx did not significantly alter alveolar macrophage abundance [16]. An intriguing finding of this study is the dissociation between hemodynamic improvement and persistent vascular muscularization in R213G mice following SuHx. While R213G mice demonstrated significantly lower mPAP compared to WT, vascular muscularization did not differ between genotypes. This observation suggests that vasoconstriction and structural remodeling represent distinct, differentially regulated processes in PH. Together, these observations underscore the model‐specific consequences of EC‐SOD redistribution and highlight that distinct inflammatory and vascular mechanisms drive PH development in chronic hypoxia vs. Sugen plus hypoxia, with important implications for understanding disease heterogeneity and for tailoring therapeutic strategies.

A second major finding of this study is the striking attenuation in interstitial macrophage (IM1 and IM2) and neutrophil accumulation in the lungs of R213G mice exposed to SuHx. In WT mice, SuHx produced a robust inflammatory response characterized by expansion of IM1 and IM2 populations, consistent with prior reports describing the early IM response to SuHx [30, 31, 32]. This pattern differs from chronic hypoxia alone, where there has been previously observed prominent IM1/IM2 expansion with minimal neutrophil involvement [26]. The enhanced neutrophil recruitment induced by SuHx aligns with studies demonstrating a role for neutrophils in vascular injury and remodeling in this model [33, 34, 35, 36, 37]. That both IM subsets and neutrophils are significantly reduced in R213G mice highlights a modulatory effect of EC‐SOD redistribution on the early inflammatory landscape.

While macrophage phenotype was not assessed fully in this study, prior work in CH demonstrated that R213G promotes a more proinflammatory macrophage transcriptome despite only modest numerical changes [29]. In SuHx, by contrast, the numerical reduction in IMs is more substantial. IM1 macrophages have been implicated in hypoxia‐induced vasoconstriction through thrombospondin‐1 (TSP‐1) expression, and bone marrow–derived TSP‐1 deficiency protects against hypoxia‐induced PH without altering vascular remodeling [38, 39, 40]. Given the marked reduction in IM1 cells in R213G mice, it is plausible that decreased IM1 abundance, and potentially reduced production of redox‐regulated mediators such as TSP‐1, contribute to the observed hemodynamic protection. The SuHx‐induced rise in IL‐6, a key cytokine implicated in PH pathogenesis, was also blunted in R213G mice [38, 41, 42, 43, 44, 45]. This finding is consistent with the reduced inflammatory cell recruitment and supports a broader anti‐inflammatory effect of increased circulating EC‐SOD. We also observed that SuHx rapidly increased hemoglobin levels in WT mice, whereas this response was blunted in R213G mice. Although the mechanism is unclear, the early timing suggests potential effects on HIF‐dependent erythropoiesis [46, 47, 48, 49]. Hematologic parameters were assessed only at the early 4‐day time point, which was chosen to coincide with our comprehensive inflammatory profiling and to capture early systemic responses to SuHx. While assessment at later time points (21–28 days) could provide information about chronic hematologic adaptations, we do not believe this would substantially alter the interpretation of our findings. Future studies could examine whether late hematologic differences correlate with the degree of hemodynamic impairment at 4 weeks.

Taken together, our findings demonstrate that EC‐SOD redistribution in R213G mice significantly alters the early inflammatory and hemodynamic response to SuHx, resulting in attenuated PH despite low lung EC‐SOD. These results emphasize that the local redox environment, both in the lung extracellular matrix and the circulation, shapes the response to pulmonary vascular injury, and that this response is strongly influenced by the nature of the injurious stimulus [50, 51, 52, 53, 54]. Chronic hypoxia and SuHx represent distinct forms of hypoxic stress that engage overlapping but non‐identical inflammatory and vascular pathways, and our results underscore the importance of considering model‐specific mechanisms when evaluating PH biology or therapeutic strategies. The early reduction in macrophage and neutrophil recruitment, along with blunted IL‐6 induction highlights the critical role of immune–vascular interactions in initiating PH. Future studies leveraging transcriptional profiling and longer‐duration exposures will be essential to determine how EC‐SOD localization shapes the functional programming of immune cells and whether these early protective effects can ultimately modify vascular remodeling and right ventricular adaptation.

Author Contributions

Daniel Colon Hidalgo and Eva S. Nozik conceived and designed the study. Daniel Colon Hidalgo, Caitlin V. Lewis, Thi‐Tina N. Nguyen, Janelle N. Posey, Samuel D. Burciaga, Nathan Dee, Christina Sul and Julie Harral performed experiments. Daniel Colon Hidalgo, David Irwin, Cassidy Delaney and Eva S. Nozik analyzed and interpreted the data. Daniel Colon Hidalgo drafted the manuscript. All authors critically revised the manuscript and approved the final version of this manuscript.

Ethics Statement

All animal experiments were approved by the Institutional Review Board of the University of Colorado (protocol 0355 and protocol 0520) and conducted in accordance with institutional guidelines.

Conflicts of Interest

The authors declare no conflicts of interest.