Certain role of extra-domain A containing fibronectin in the development of pulmonary hypertension in the sugen/hypoxia mouse model

Introduction

Pulmonary hypertension (PH) is hemodynamically defined as an increased mean pulmonary arterial pressure (mPAP) >20 mmHg at rest. Due to chronic pressure overload, PH ultimately can lead to right heart failure (RHF), resulting in a severely unfavourable prognosis. PH presents with a spectrum of unspecific clinical symptoms and, etiologically, encompasses a heterogeneous variety of underlying disorders, on which the clinical classification into five distinct groups is based: group 1: pulmonary arterial hypertension (PAH); group 2: PH associated with left heart disease; group 3: PH associated with lung diseases and/or hypoxia; group 4: PH associated with pulmonary artery obstructions; and group 5, which comprises PH with unclear and/or multifactorial mechanisms. This classification enables individualized risk stratification, prognostic assessment, and the selection of appropriate targeted therapy (1-3). To date, disease-specific therapies are available exclusively for group 1 PH (for instance endothelin receptor antagonists, prostanoids, phosphodiesterase inhibitors) and group 4 PH (soluble guanylate cyclase antagonists). Aforementioned therapies primarily modulate vascular tone to reduce mPAP, thereby decreasing right ventricular (RV) load. However, they do not halt or reverse disease progression. Regarding groups 2 and 3 PH, which together account for over 90% of cases in Europe, treatment remains limited to symptom management and addressing the underlying condition. Due to its nonspecific symptoms, frequently resulting in delayed diagnosis, and the absence of effective causal treatment options, PH remains a disease with poor prognosis and high mortality rates (1,2,4-6). As such, novel treatment approaches, particularly for groups 2 and 3 PH, represent a critical unmet clinical need.

Pathophysiologically, PH is characterized by vasoconstriction, microthrombosis, inflammation, and extensive remodelling of the pulmonary vasculature (2,7,8). Pulmonary vascular remodelling affects all layers of the vessel wall and involves unregulated endothelial cell proliferation leading to the formation of plexiform lesions, alongside stimulation and proliferation of smooth muscle cells in the tunica media, resulting in massive vascular wall thickening. By contributing to elevated pulmonary arterial pressure, subsequently leading to RV overload and RHF, pulmonary vascular remodelling has a pivotal role in both the onset and progression of PH (2,9). Fibronectin (Fn) is a relevant structural component of the extracellular matrix (ECM). Through alternative splicing of its pre-mRNA, various fetal molecular variants are generated, among which the extra-domain A containing fibronectin (ED-A⁺ Fn) is of particular interest. While ED-A⁺ Fn is physiologically expressed during embryological heart development, it is virtually absent in healthy adult cardiac tissue. However, in context of cardiovascular and chronic inflammatory diseases (e.g., atherosclerosis, arthritis, psoriasis), ED-A⁺ Fn is abundantly re-expressed and released into circulation (10-14). Following tissue injury, ED-A⁺ Fn is secreted predominantly by macrophages and other immune cells. It activates toll-like receptor 4 (TLR-4), promoting the release of proinflammatory cytokines and the recruitment of immune cells, such as mast cells and T cells, thereby amplifying the inflammatory response (15-18). ED-A⁺ Fn further is a crucial cofactor of TGF-β1 induced phenotype transition of fibroblasts into contractile myofibroblasts (MyoFb) and stimulates vascular smooth muscle cells (VSMC). These activated cell types subsequently secrete additional ED-A⁺ Fn, establishing a positive feedback loop that perpetuates tissue remodelling. This loop plays a central role in vasculopathy and fibrosis, significantly contributing to the development, severity, and chronicity of disease (19-23).

Previous studies conducted by our group have demonstrated that the extensive re-expression of ED-A⁺ Fn is critically involved in the development and progression of PH and consecutive RHF in the mouse model of monocrotaline (MCT)-induced PH (24-26). In an ED-A⁺ Fn knockout (KO) model, the absence of the molecule led to significant attenuation of pulmonary vascular and RV myocardial remodelling. These findings support the hypothesis that ED-A⁺ Fn represents a promising biomarker and potential therapeutic target able to mitigate pathological tissue remodelling in PH, for instance through the administration of specific neutralizing antibodies (26).

We therefore aimed to further elucidate the functional contribution of ED-A⁺ Fn to the development and progression of hypoxia associated PH and consecutive RHF. Accordingly, we implemented the mouse model of Sugen5416/hypoxia (SuHx) induced PH in a comparison of ED-A⁺ Fn KO and corresponding wild-type (WT) mice. This preclinical model of PH not only depicts the features of group 1 PH but also key pathological mechanisms of group 3 PH (27,28).

Given that groups 2 and 3 PH together account for more than 90% of PH cases in Europe and that, apart from treatment of the underlying left heart or lung disease, no causal or disease-specific therapeutic options are currently available, validation in the SuHx model was essential. We hypothesized that the absence of ED-A⁺ Fn would essentially ameliorate disease-associated remodelling and RHF in this preclinical PH-model as well, thereby providing further evidence of its potential as a novel therapeutic target and extending our findings beyond PAH to clinically predominant PH subtypes. We present this article in accordance with the ARRIVE reporting checklist (available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-443/rc).

Methods

Mouse model of Sugen5416/hypoxia-induced PH

A total of 26 male mice (age: 10 to 12 weeks; body weight: 20 to 30 g) were included in this study. Prior to PH induction, all animals underwent a seven-day acclimatization period, which included controlled light/dark cycles, a constant ambient temperature, ad libitum access to food and water, and care by experienced animal caretakers.

A protocol was prepared before the study without registration. The experimental protocol received approval from the Thuringian State Office for Food Safety and Consumer Protection (TLLV, Bad Langensalza, Germany; registration number: UKJ21-010). All experimental procedures were carried out in accordance with the guidelines for the care and use of laboratory animals (National Research Council Committee, Guide for the Care and Use of Laboratory Animals, 2011) and the current version of the German Animal Welfare Act and applicable animal husbandry guidelines.

Of the 26 animals applied, 16 were C57BL/6J mice obtained from Janvier Labs (Rte du Genest, 53940 Le Genest-Saint-Isle, France) and 10 were ED-A⁺ Fn-KO mice. The latter were originally provided to the Cardiovascular Remodelling working group of University Hospital Jena (UKJ) by the International Centre for Genetic Engineering and Biotechnology in Trieste, Italy and established at UKJ via embryo transfer. These ED-A⁺ Fn-KO mice share the same C57BL/6J genetic background but lack the ability to produce ED-A⁺ Fn due to the targeted deletion of the exon encoding the ED-A domain (29).

The 26 animals were divided into four experimental groups (Figure 1): normoxic wild-type mice (WTNx, n=8), SuHx-induced PH in wild-type mice (WTSuHx, n=8), normoxic ED-A⁺ Fn-KO mice (KONx, n=5) and SuHx-induced PH in ED-A⁺ Fn-KO mice (KOSuHx, n=5). Allocation of animals to the respective experimental groups was performed in a non-randomized manner. The experiment was conducted without any exclusions. PH was induced by subjecting the animals to 28 days of chronic normobaric hypoxia (10% O₂) in a ventilated chamber. In addition, on day 1, the mice were administered a single subcutaneous (s.c.) injection of Sugen5416 (Semaxinib, MCE-HY-10374, Biozol Diagnostica Vertrieb GmbH, Hamburg, Germany, 200 mg/kg body weight), a vascular endothelial growth factor receptor antagonist, suspended in dimethyl sulfoxide (DMSO). Normoxic control animals, which did not undergo PH induction, were maintained in room air and received a s.c. injection of the vehicle DMSO only.

Figure 1 Experimental design. DMSO, dimethyl sulfoxide; ED-A⁺ Fn, extra-domain A of fibronectin; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; s.c., subcutaneous; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

Monitoring of clinical health status

Throughout the study period, all mice were assessed daily, and their clinical status was monitored twice weekly through body weight measurements and examinations using a clinical severity score (CSS) system, which was adapted for the SuHx-induced PH model. The CSS evaluates general signs of disease, spontaneous activity, response to external stimuli, posture, and body weight. Each parameter was scored individually from 0 to 3 (30). No adverse reactions were observed in the mice during the experimental period or procedures.

Echocardiography

On day 28 post-induction (p.i.) of PH, transthoracic echocardiography (TTE) was performed on all animals under volatile anaesthesia using isoflurane (2.5% isoflurane-CP, FiO₂ 1.0, oxygen inhalation flow rate) by applying the Vevo 770 Rodent Ultrasound System (VisualSonics, Canada) with a 17MHz RMV176 probe to assess both, morphological and functional parameters. The morphological parameters included RV basal and medial diameters (in mm) and RA area (in mm²). Functional parameters for assessing RV systolic function included tricuspid annular plane systolic excursion (TAPSE, in mm) and fractional area change [FAC, in %, calculation: (enddiastolic RV area − endsystolic RV area)/enddiastolic RV area].

Right heart catheterization (RHC)

As a final step, RHC was conducted on all animals through by introducing a 1.4F micro conductance pressure-volume catheter (Model 10 SPR-839, Millar Instruments Inc., connected to PowerLab system, ADInstruments Ltd., Oxford, UK) into the right internal jugular vein. For the procedure, animals were anesthetized with a combination of ketamine (100 mg/kg body weight) and xylazine (10 mg/kg body weight). Pressure measurements were obtained in the right ventricle or pulmonary artery. Following RHC, the mice were euthanized under deep anesthesia and analgesia for organ harvesting by opening the thorax and transecting the diaphragm.

Histology

PH-associated damage of lung and RV myocardial tissue was assessed using semi-quantitative scoring systems developed and validated by our group (25,26). For histological analysis, 4 µm-thick sections of lung and RV myocardial tissue were prepared and stained with hematoxylin and eosin (H&E) as well as picrosirius red (PSR) following standardized protocols. Microscopic evaluation was performed using the Zeiss AxioImager microscope equipped with the Axiocam 506 color camera (Carl Zeiss AG, Oberkochen, Germany). Image acquisition was carried out using the ZEN Software (Carl Zeiss AG, Oberkochen, Germany), with all scoring procedures performed in a double-blinded manner.

The scoring system of PH associated lung tissue damage is based on five histopathological features commonly observed in PH: the percentage of atelectasis area (0: not detectable, 1: <30%, 2: ≥30% of tissue area), the percentage of emphysematous areas (0: not detectable, 1: <30%, 2: ≥30% of tissue area), the degree of media hypertrophy of peribronchial arteries (0: not detectable, 1: mild, 2: moderate, 3: severe hypertrophy), the presence of perivascular cellular edema of peribronchial arteries (0: absent, 2: present), and the degree of media hypertrophy of small arteries not directly associated with bronchi (0: not detectable, 1: mild, 2: moderate, 3: severe hypertrophy). The maximum score for lung tissue damage was 12 points, with higher scores indicating more severe pathology.

RV myocardial tissue was evaluated assessing interstitial cellularity, particularly the infiltration of inflammatory cells, and the extent of interstitial fibrosis. Both parameters were assessed semi-quantitatively on a scale from 0 to 3 (0: not detectable, 1: mild, 2: moderate, 3: severe). The total score for RV tissue damage ranged from 0 to 6 points, with higher scores indicating more severe tissue damage.

Immunofluorescence labelling

Double labelling of α-SMA and CD31 in lung tissue

Double immunofluorescence labelling of α-smooth muscle actin (α-SMA) and CD31 in lung tissue was performed sequentially, beginning with the protocol used for single labelling of α-SMA. Sections were first incubated with a rabbit anti-α-SMA primary antibody (ab5694, Abcam, Berlin, Germany; dilution 1:200) for 60 minutes at room temperature (RT). Following washing with TBS-T, the corresponding Cy^TM3-conjugated AffiniPure donkey anti-rabbit IgG secondary antibody (Jackson ImmunoReseach Laboratories Inc., Pennsylvania, USA; dilution 1:400) was applied for 45 minutes at RT, followed by additional TBS-T washes. Subsequently, CD31 staining was done by applying the same procedure. Sections were incubated with rat anti-mouse CD31 primary antibody (MEC-13.3, BD Biosciences, Franklin Lakes, NJ, USA; dilution 1:1,000) for 60 minutes at RT, washed with TBS-T, and then incubated with FITC-conjugated AffiniPure donkey anti-rat IgG secondary antibody (Jackson ImmunoReseach Laboratories Inc., Pennsylvania, USA; dilution 1:400) for 45 minutes at RT. After final washing with TBS-T and distilled water, sections were mounted using Vectashield mounting medium containing DAPI (Vector Laboratories, Burlingame, CA, USA) for nuclear counterstaining and stored at −20 ℃ until immunofluorescence microscopy analysis. Antibody specificity controls were included by omitting the primary antibody during incubation.

Detection of ED-A⁺ Fn in lung tissue

Detection of ED-A⁺ Fn in lung tissue was performed using the Tyramide SuperBoost^TM Kit with Alexa Fluor^TM Tyramides (Thermo Fisher Scientific, Dreireich, Germany). The recombinant, biotinylated F8-small immunoprotein (SIP) antibody specifically recognizing ED-A⁺ Fn was kindly provided by Prof. Dr. D. Neri (Philochem AG, Otelfingen, Switzerland). Following the protocol for single immunohistochemical labelling, 4 µm cryosections were fixed sequentially in methanol and acetone, then washed with phosphate-buffered saline containing Tween-20 (PBS-T). To block endogenous biotin-binding sites, a biotin-blocking system was applied, which consisted of a 30-minute incubation with avidin followed by a 30-minute incubation with biotin. Endogenous peroxidase activity was quenched by incubating the sections in a 100× hydrogen peroxide (H₂O₂) solution (50 µL H₂O₂ in 1 mL distilled water) for 60 minutes. Blocking was then performed using the kit’s blocking buffer (10% goat serum) for 60 minutes at RT. The biotinylated primary antibody (F8-SIP-bio, dilution 1:100) was applied and incubated overnight at 4 ℃, followed by a 60-minute incubation with streptavidin-horseradish peroxidase to bind the biotinylated antibody. Tyramide signal amplification was performed by incubating the sections with the tyramide-Alexa488 working solution for 10 minutes, followed by a 1-minute stop solution application. Finally, sections were mounted using Vectashield mounting medium containing DAPI (Vector Laboratories, Burlingame, CA, USA) and stored at −20 ℃ until immunofluorescence microscopy analysis. Antibody specificity controls were included by omitting the primary antibody during incubation.

Collagen type III alpha 1 chain (Col3α1) in myocardial tissue, CD45-positive and CD68-positive cells in lung tissue

Immunofluorescence staining of the Col3α1, CD45-positive, and CD68-positive cells was performed on 4 µm cryostat sections. Tissue sections were fixed in methanol (−20 ℃) for 20 seconds, followed by fixation in acetone (−20 ℃) for nine minutes. For Col3α1 detection, a polyclonal rabbit anti-Col3α1 antibody (ab7778, Abcam, Berlin, Germany; dilution 1:200) was applied and incubated for 60 minutes at RT. CD45 was detected using a rat anti-mouse CD45 antibody (Clone 30-F11, BD Biosciences, Heidelberg, Germany; dilution 1:200), and CD68 was detected using a rat anti-mouse CD68 antibody (FA-11, Bio-Rad Laboratories GmbH, Munich, Germany; dilution 1:200), both incubated for 90 minutes at RT. All primary antibodies were diluted in Dako Antibody Diluent with Background Reducing Components (Dako Agilent Technologies Inc., Waldbronn, Germany). Following primary antibody incubation, slides were washed with TBS-T washing buffer, then incubated for 45 minutes at RT with the corresponding secondary antibodies. For Col3α1, Cy™3-conjugated AffiniPure donkey anti-rabbit IgG antibody was used; for CD45, Cy™3-conjugated AffiniPure donkey anti-rat IgG antibody; and for CD68, FITC-conjugated AffiniPure donkey anti-rat IgG antibody (Jackson ImmunoResearch, Pennsylvania, USA; dilution 1:400 for all secondary antibodies). After a final wash in TBS-T and once in distilled water, sections were mounted using Vectashield mounting medium containing DAPI (Vector Laboratories, Burlingame, CA, USA) for nuclear counterstaining (blue fluorescence), and stored at −20 ℃ until immunofluorescence microscopy analysis. Antibody specificity controls were included by omitting the primary antibody during incubation.

Immunohistochemical evaluation

Microscopic evaluation of α-SMA/CD31-, Col3α1-, ED-A⁺ Fn-, CD45-, and CD68-labelled tissue sections was performed using the Zeiss AxioImager microscope equipped with the Axiocam 506 mono camera (Carl Zeiss AG, Oberkochen, Germany). Image acquisition was carried out using the ZEN Software (Carl Zeiss AG, Oberkochen, Germany).

Semi-quantitative assessment of staining patterns was performed using an established semiquantitative scoring system: 0= no detectable signal in the tissue, 1= low to mild expression or infiltration, 2= moderate expression or infiltration, and 3= strong expression or infiltration throughout the tissue. To ensure reliable differentiation between specific immunoreactivity and non-specific background staining, corresponding negative controls were regularly included in the analysis for each sample.

Statistical analysis

Statistical analyses and graphical representation were performed using Microsoft Excel (2024, Version 16.85), IBM SPSS Statistic (2022, Version 29), and JMP Statistical Discovery (2024, Version 18.0.1). For multiple group comparisons, the Kruskal-Wallis test was used first. When global significance was found, group differences were assessed using non-parametric Mann-Whitney U test. A P value of <0.05 was considered statistically significant. To account for multiple comparisons, pairwise Mann-Whitney U tests were adjusted using the Bonferroni-Holm correction method. Data are presented as the mean ± standard deviation (SD), and all statistical tests were two-sided. An a priori power analysis based on hemodynamic and echocardiographic data from our preliminary studies indicated that a sample size of 5–6 animals per group yields a statistical power exceeding 95%. In addition, individual pairwise group comparisons (e.g., WTNx vs. WTSuHx) can be conducted with a statistical power of 87.5%.

Results

Health status of mice during experimental period

All animals were healthy at baseline (CSS 0 on day 1). Around day 10 p.i., SuHx-induced mice showed clinical deterioration, manifested as reduced activity, particularly in WTSuHx mice (CSS 2 on day 28) compared to KOSuHx (CSS 1 on day 28) and normoxic controls (CSS 0 on day 28 for WTNx and KONx; Figure 2A). Body weight changes from day 1 to 28 differed significantly between groups (Figure 2B and Figure S1). While normoxic WT (2.78±0.49 g) and KO animals (2.6±1.08 g) gained weight, SuHx-induced WT mice tended to lose weight (−1.08±1.5 g).

Figure 2 Assessment of health status in SuHx-induced WT and KO mice during experimental period. The boxplots (median and interquartile range) represent the CSS on day 28; P values for CSS: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P=0.02 (A) and weight difference between day 1 and day 28; P values for weight difference: WTNx vs. KONx: P=0.87; KONx vs. KOSuHx: P=0.048 (B) of each experimental group (n=26, WTNx n=8, WTSuHx n=8, KONx n=5, KOSuHx n=5). The Mann-Whitney U test was applied to screen for significant differences between the groups. For clearer illustrations statistically non-significant P values were left out. CSS, clinical severity score; ED-A⁺ Fn, extra-domain A of fibronectin; KO, knockout; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; SuHx, Sugen5416/hypoxia; WT, wild-type; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

RHC

Animals were subjected to invasive RHC on day 28 p.i. to assess systolic right ventricular pressure (RVP_sys) in all animals. RHC revealed a significant increase in RVP_sys in PH-induced WTSuHx mice (85.2±11.4 mmHg) compared to the WTNx (45.2±4.6 mmHg, P=0.006), KONx (38.9±4 mmHg, P=0.02), and KOSuHx (49.8±9.5 mmHg, P=0.02) mice (Figure 3A). No significant differences in pressure values were observed within the KO groups (P=0.10), as the RVP_sys of the KOSuHx mice exhibited only a marginal elevation relative to normoxic WT and KO controls.

Figure 3 Assessment of hemodynamic and echocardiographic parameters in SuHx-induced WT and KO mice. On day 28, transthoracic echocardiography and right heart catheterization were performed to assess right heart morphology and function. The boxplots (median and interquartile range) represent the RVP_sys; P values for RVP_sys: WTNx vs. KONx: P=0.15; KONx vs. KOSuHx: P=0.10 (A), RV_basal; P values for RV_basal: WTNx vs. KONx: P=0.02; KONx vs. KOSuHx: P=0.06 (B), right atrial area (RA_area); P values for RA_area: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P>0.99 (C) and the TAPSE; P values for TAPSE: WTNx vs. KONx: P=0.94; KONx vs. KOSuHx: P=0.02 (D) of each experimental group (n=26, WTNx n=8, WTSuHx n=8, KONx n=5, KOSuHx n=5). ED-A⁺ Fn, extra-domain A of fibronectin; KO, knockout; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; RV_basal, basal right ventricular diameter; RVP_sys, systolic right ventricular pressure; SuHx, Sugen5416/hypoxia; TAPSE, tricuspid annular plane systolic excursion; WT, wild-type; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

Echocardiographic evaluation

TTE was carried out on day 28 p.i. on all animals. Significant alterations in echocardiographic parameters were observed in the WTSuHx group relative to all other groups. Basal RV diameter (RV_basal) was significantly increased in WTSuHx mice (2.6±0.2 mm) compared to normoxic WTNx (2.1±0.1 mm, P=0.006), normoxic KONx (1.9±0.1 mm, P=0.01), and SuHx-induced KOSuHx (2.1±0.2 mm, P=0.01) mice (Figure 3B). SuHx-induced KO mice (2.14±0.19 mm) showed a trend towards smaller RV_basal compared to WTSuHx (2.57±0.15 mm), but no significant structural changes were observed within the KO groups after PH induction (P=n.s.). For RA_area (Figure 3C), WTSuHx mice (7.8±1.2 mm²) exhibited a significant increase compared to normoxic WTNx (4.3±0.5 mm², P=0.006) and KONx (4.1±0.7 mm², P=0.03), while KOSuHx mice (4±0.9 mm²) did not differ significantly from normoxic WT or KO controls (P=n.s.).

TAPSE, a parameter of RV systolic function, was significantly reduced in both, WTSuHx (0.37±0.04 mm, P=0.006) and KOSuHx (0.41±0.02 mm, P=0.02) mice compared to their respective normoxic controls (Figure 3D), with no significant difference between the two SuHx groups (P=n.s.). Pairwise comparisons of fractional area change (FAC), a functional RV parameter, showed no significant differences between individual groups (P=n.s.), though all SuHx-induced animals (WTSuHx: 18.91%±8.99%, KOSuHx: 21.27%±13.5%) exhibited a trend towards reduced FAC compared to normoxic controls (WTNx: 45.18%±4.59%, KONx: 38.9%±4.03%).

Due to technical limitations during invasive RHC and echocardiographic assessment, complete datasets could not be obtained for all parameters in every animal. For RVP_sys, one measurement was unavailable in the KONx group (n=4). Analyzable data for RA_area were acquired for WTNx (n=6), WTSuHx (n=7), KONx (n=4), and KOSuHx (n=4). For FAC, datasets were available for WTNx (n=7), WTSuHx (n=7), KONx (n=2), and KOSuHx (n=4). Statistical analyses were performed based on the available data for each parameter.

Analysis of tissue damage in the lung

By histological analysis of H&E-stained lung tissue sections using a semiquantitative scoring system, significant lung tissue alterations (Figure 4) in association to the development of PH could be observed in both, WTSuHx (5.63±0.7, P=0.006) and KOSuHx (0.13±0.22, P=0.03) mice compared to their respective normoxic controls (WTNx: 0.06±0.17, KONx: 0±0; Figure 4A). Moreover, lung tissue damage was significantly more pronounced in WTSuHx mice than in both, KONx (P=0.01) and KOSuHx (P=0.01) groups. Following PH induction, WTSuHx mice showed significant hypertrophy of the tunica media in both, peribronchial arteries and small arteries not related to bronchi (Figure 4B), compared to normoxic WT controls (P=0.006 for both) and KONx mice (P=0.01 for both). No significant vascular alterations were observed between normoxic and SuHx-induced KO groups (P=n.s.).

Figure 4 Assessment of histological damage in lung tissue and immunohistochemical staining of α-SMA and CD31 in lung tissue of SuHx-induced WT and KO mice using defined scoring systems. The boxplots (median and interquartile range) depict the sum scores of lung tissue analysis; P values for Sum Score lung: WTNx vs. KONx: P=0.35; KONx vs. KOSuHx: P=0.03 (A) and the quantification of α-SMA/CD31; P values for α-SMA/CD31 scoring: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P>0.99 (C) in lung tissue of each experimental group (n=26, WTNx n=8, WTSuHx n=8, KONx n=5, KOSuHx n=5). Representative images of the H&E-stained lung tissue sections assessed by light microscopy for the different experimental groups are given in (B). Representative images of double immunofluorescence labelling of α-SMA (Cy3, red fluorescence) and CD31 (FITC, green fluorescence) in lung tissue are given in (D). Negative controls (omission of primary antibody) are shown in the upper left corner (DAPI/nuclei, blue fluorescence). α-SMA, alpha-smooth muscle actin; DAPI, 4',6-diamidino-2-phenylindole; ED-A⁺ Fn, extra-domain A of fibronectin; FITC, fluorescein isothiocyanate; H&E, hematoxylin and eosin; KO, knockout; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; SuHx, Sugen5416/hypoxia; TAPSE, tricuspid annular plane systolic excursion; WT, wild-type; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

To further assess the extent of media hypertrophy and PH-induced endothelial dysfunction, double immunofluorescence staining of α-SMA and the endothelial cell marker CD31 was performed (Figure 4D). WTSuHx mice (2.31±0.43) exhibited significantly increased α-SMA expression and disruption of the endothelial cell layer in pulmonary vessels, indicated by discontinuous CD31 staining, compared to WTNx (1±0, P=0.006), KONx (1±0, P=0.01), and KOSuHx (1±0, P=0.01). No significant differences were observed between KONx and KOSuHx mice (P=n.s.; Figure 4C).

Analysis of tissue damage in the RV myocardium

Histopathological analysis of RV myocardial tissue (Figure 5) was conducted on H&E and PSR-stained paraffin sections (Figure 5B and Figure S2) using a semi-quantitative scoring system assessing interstitial cellularity and fibrosis. As expected, myocardial damage in normoxic controls was negligible, whereas significant PH-associated tissue damage was observed in SuHx-induced groups. WTSuHx mice (0.88±0.41) exhibited significantly higher levels of RV tissue damage compared to WTNx controls (0.06±0.17, P=0.006; Figure 5A). Although tissue damage appeared more pronounced in WTSuHx than in KOSuHx mice (0.13±0.22), this difference did not reach statistical significance (P=n.s.). Consistently, no significant differences were observed between the KO groups (KONx: 0±0, P=n.s.).

Figure 5 Assessment of histological damage and immunohistochemical staining of Col3α1 in right ventricular myocardial tissue of SuHx-induced WT and KO mice using defined scoring systems. The boxplots (median and interquartile range) depict the sum scores of right ventricular myocardial tissue analysis; P values for RV Sum Score: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P>0.99 (A) and the quantification of Col3α1; P values for Col3α1 scoring: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P=0.38 (C) in cardiac tissue of each experimental group (n=26, WTNx n=8, WTSuHx n=8, KONx n=5, KOSuHx n=5). Representative images of the H&E (left) and PSR (right) stained cardiac tissue sections assessed by light microscopy for the different experimental groups are given in (B). Representative images of immunofluorescence labelling of Col3α1 (Cy3, red fluorescence) in cardiac tissue are given in (D). Negative controls (omission of primary antibody) are shown in the upper left corner (DAPI/nuclei, blue fluorescence). Col3α1, collagen type III alpha 1 chain; DAPI, 4',6-diamidino-2-phenylindole; ED-A⁺ Fn, extra-domain A of fibronectin; KO, knockout; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; PSR, picrosirius red; RV, right ventricular; SuHx, Sugen5416/hypoxia; TAPSE, tricuspid annular plane systolic excursion; WT, wild-type; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

Concomitantly, immunohistochemical detection of Col3α1, a marker of interstitial activation and fibrosis, revealed a visibly and significantly increased expression in RV myocardial tissue of WTSuHx mice (1.06±0.63) compared to WTNx (0±0) and KONx (0±0) controls (P=0.03 for both; Figure 5C). This increase reflects an interstitial fibrotic response as part of myocardial remodelling and the subsequent right heart strain (pressure overload) induced by PH (Figure 5D and Figure S3). No significant differences in Col3α1 expression were detected between KONx and KOSuHx mice (0.3±0.24, P=n.s.).

Notably, representative left ventricular (LV) myocardial tissue samples showed no relevant histologic alterations after PH induction.

ED-A⁺ Fn detection in lung tissue

Re-expression of ED-A⁺ Fn in lung tissue (Figure 6) was assessed by immunofluorescence staining using the F8 antibody specific for ED-A⁺ Fn. PH induction in the SuHx mouse model led to significant re-expression and pronounced vascular and interstitial deposition of ED-A⁺ Fn in the WTSuHx group (2.75±0,43; Figure 6B), that was significantly higher compared to WTNx (0±0, P=0.006), KONx (0±0, P=0.01) and KOSuHx (0±0, P=0.01) groups (Figure 6A). KO groups lacking the ability to produce ED-A⁺ Fn did not show any re-expression of the target molecule under either normoxic or SuHx conditions.

Figure 6 Assessment of re-expression of ED-A⁺ Fn in lung tissue of SuHx-induced WT and KO mice using a defined scoring system. The boxplots (median and interquartile range) show the quantification of ED-A⁺ Fn in lung tissue; P values for ED-A⁺ Fn scoring: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P>0.99 (A) of each experimental group (n=26, WTNx n=8, WTSuHx n=8, KONx n=5, KOSuHx n=5). For clearer illustrations statistically non-significant P values were left out. Representative images of immunofluorescence labelling of ED-A⁺ Fn (Alexa488, green fluorescence) in lung tissue are given in (B). Negative controls (omission of primary antibody) are shown in the upper left corner (DAPI/nuclei, blue fluorescence). DAPI, 4',6-diamidino-2-phenylindole; ED-A⁺ Fn, extra-domain A containing fibronectin; KO, knockout; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; SuHx, Sugen5416/hypoxia; TAPSE, tricuspid annular plane systolic excursion; WT, wild-type; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

Perivascular inflammation in lung tissue

Perivascular inflammation (Figure 7), another key pathogenetic feature of PH, was assessed by immunohistochemical staining of leukocytes using the surface marker CD45 (Figure 7B) and macrophages via immunofluorescence labelling of CD68⁺ cells (Figure 7D). CD45⁺ cells showed a homogeneous distribution with moderate to intense accumulation in normoxic groups (WTNx and KONx), whereas WTSuHx (2.56±0.53) exhibited a significant increase in leukocyte infiltration and locally accentuated accumulation compared to WTNx (1±0.43, P=0.006), KONx (0.8±0.24, P=0.01), and KOSuHx (0.6±0.2, P=0.01). Although CD45⁺ cells appeared visually increased in KOSuHx versus KONx, this difference was not statistically significant (P=n.s.; Figure 7A). Concomitantly, WTSuHx mice (2.31±0.5) showed a significantly increased accumulation of CD68⁺ macrophages compared to the moderate, homogenous macrophage distribution observed in WTNx (0.88±0.33, P=0.006), KONx (0.7±0.24, P=0.01), and KOSuHx (0.8±0.24, P=0.01). This could not be shown when comparing normoxic and SuHx-induced KO groups (P=n.s.; Figure 7C).

Figure 7 Assessment of immunohistochemical staining of CD45 and CD68 in lung tissue of SuHx-induced WT and KO mice using defined scoring systems. The boxplots (median and interquartile range) show the quantification of CD45⁺ cells; P values for CD45⁺ scoring: WTNx vs. KONx: P=0.62; KONx vs. KOSuHx: P=0.62 (A) and CD68⁺ cells; P values for CD68⁺ scoring: WTNx vs. KONx: P>0.99; KONx vs. KOSuHx: P>0.99 (C) in lung tissue of each experimental group (n=26, WTNx n=8, WTSuHx n=8, KONx n=5, KOSuHx n=5). Representative images of immunofluorescence labelling of CD45 (Cy3, red fluorescence) in lung tissue are given in (B). Representative images of immunofluorescence labelling of CD68 (FITC, green fluorescence) in lung tissue are given in (D). Negative controls (omission of primary antibody) are shown in the upper left corner (DAPI/nuclei, blue fluorescence). DAPI, 4',6-diamidino-2-phenylindole; ED-A⁺ Fn, extra-domain A containing fibronectin; FITC, fluorescein isothiocyanate; KO, knockout; KONx, normoxic ED-A⁺ Fn knockout mice; KOSuHx, Sugen/hypoxia induced PH in ED-A⁺ Fn knockout mice; PH, pulmonary hypertension; SuHx, Sugen5416/hypoxia; TAPSE, tricuspid annular plane systolic excursion; WT, wild-type; WTNx, normoxic wild-type mice; WTSuHx, Sugen/hypoxia induced PH in wild-type mice.

Discussion

Several studies have identified ED-A⁺ Fn as a potential biomarker and therapeutic target in PH, particularly in clinical groups 1, 2 and 3. Elevated serum levels of ED-A⁺ Fn were observed in these patient groups compared to healthy controls. In contrast, no such increase could be shown for PH group 4 (CTEPH), likely reflecting the absence of similar pulmonary vascular remodelling processes. ED-A⁺ Fn serum concentrations correlated positively with key clinical and hemodynamic parameters, especially systolic pulmonary arterial pressure and BNP levels, and inversely with 6-minute walk distance (31), supporting its utility for disease monitoring and early detection, particularly in group 1 of PH. Preclinical studies confirmed the association of elevated ED-A⁺ Fn with increased mPAP and PH severity in MCT-induced animal models (24,26).

In the present study, we employed the SuHx-induced PH mouse model to investigate the causal role of ED-A⁺ Fn in PH pathogenesis and to further evaluate its potential as both, a diagnostic marker and a therapeutic target. The combination of VEGFR inhibition (SU5416) and chronic hypoxia induces severe PH characterized by extensive vascular remodelling and angioobliterative lesions, resembling plexiform lesions observed in PAH patients (32,33). Notably, its pathological effects are largely restricted to the pulmonary vasculature. The SuHx model recapitulates key features of chronic human PH, especially those of PH groups 1 and 3. Histological changes in SuHx-exposed mice closely resemble those observed in both, the MCT model and in human PH as well (24). While MCT-induced PH is associated with acute endothelial toxicity and inflammation, SuHx-induced PH progresses more chronically and is characterized by proliferative pulmonary vascular lesions (32). Due to disease severity, the reproducible development of intimal lesions and its progressive nature, the SuHx model is well-established for preclinical research (34). It allows the use of genetically modified mice, facilitating molecular insights into PH pathogenesis and the identification of therapeutic targets (35). In this study, we utilized ED-A⁺ Fn KO mice to assess the functional role of this fetal molecule in PH pathogenesis.

Key findings

Notably, WTSuHx mice exhibited a relevant clinical phenotype compared to their ED-A⁺ Fn-deficient KO counterparts, in which a significantly attenuated PH induction and a milder disease progression could be proven on hemodynamic, echocardiographic and microscopic level.

When comparing KONx and KOSuHx animals, RVP_sys did not reach statistical significance (P=0.10), despite a marginal increase in mean values in KOSuHx mice compared with normoxic WT and KO controls. While this could raise the question of whether the absence of statistical significance is related to limited sample size, this explanation appears unlikely. Based on hemodynamic and echocardiographic data from our preliminary studies, an a priori power analysis indicated that group sizes of 5–6 animals provide sufficient statistical power (>95%) to detect relevant differences, and that pairwise comparisons remain adequately powered (87.5%). Accordingly, the lack of a significant difference in RVP_sys between KONx and KOSuHx animals more likely reflects the absence of a robust PH phenotype in KOSuHx mice rather than insufficient statistical power. This interpretation is further supported by the consistent pattern observed across all KO groups and by the fact that a significant increase in RVP_sys was exclusively detected in WTSuHx animals compared with both normoxic controls and KOSuHx mice, indicating successful induction of PH only in the WT background.

Although RVP_sys did not increase significantly in KOSuHx mice compared with KONx animals, SuHx exposure in the absence of ED-A⁺ Fn was associated with significant changes in TAPSE and body weight (KONx vs. KOSuHx), indicating mild functional alterations despite the absence of overt PH. Consistent with this notion, echocardiographic assessment revealed a trend toward reduced RV performance in KOSuHx mice without the pronounced RV pressure elevation or structural remodelling observed in WTSuHx mice. We additionally calculated RV-PA coupling as the ratio of TAPSE in mm to RVPsys in mmHg to further elucidate the correlation between pulmonary vascular load and RV function. WTSuHx mice (0.0047 mm/mmHg) demonstrated a marked reduction in RV-PA coupling compared to all other groups (WTNx: 0.0155 mm/mmHg, KONx: 0.018 mm/mmHg, KOSuHx: 0.008 mm/mmHg), reflecting pronounced RV vascular uncoupling in response to SuHx-induced PH. KOSuHx mice exhibited intermediate coupling values, suggesting only partial functional impairment, consistent with the mild or absent PH phenotype observed hemodynamically. Although these differences did not reach statistical significance (P>0.99 for all comparisons), the observed trend indicates that the ED-A⁺ Fn KO provides partial protection from SuHx-induced pulmonary vascular remodelling and RV afterload increase, as evidenced in rather preserved RV-PA coupling despite a modest reduction in TAPSE.

The markedly attenuated disease phenotype in ED-A⁺ Fn deficient mice in the SuHx-induced PH model highlights the pivotal functional role of ED-A⁺ Fn in PH development, primarily by promoting pulmonary vascular and myocardial remodelling, and emphasizes the therapeutic potential of targeting ED-A⁺ Fn to mitigate said remodelling and improve clinical outcomes in PH.

Limitations

Limitations of this study should be acknowledged. Sample sizes were relatively small and uneven between groups (e.g., WT n=8 vs. KO n=5), which may have limited the ability to detect subtle phenotypic differences. Furthermore, group allocation was not randomized, which may have introduced a potential selection bias. In addition, only single-sex (male) mice were chosen to ensure comparability with the existing ED-A⁺ Fn literature, which predominantly uses male-only cohorts. Moreover, the use of a prophylactic genetic KO model precludes conclusions regarding therapeutic efficacy after disease onset. In addition, the mechanistic pathways underlying the observed effects were not investigated in detail and warrant further research. Finally, comprehensive quantitative morphometric analyses of fibrosis, including collagen content and fiber organization, could not be performed due to technical constraints and a limited tissue availability. Therefore, histological evaluation relied on established semi-quantitative scoring methods.

Implications

Previous studies have demonstrated disease-modifying effects in MCT-induced PH models using a function-blocking recombinant antibody specific to ED-A⁺ Fn (F8) (26). This antibody allows targeted delivery of therapeutic agents or direct functional blockade after systemic administration, e.g., intravenously (36). Immunohistochemical co-localization of the F8 generated labelling with α-SMA in murine MCT-induced PH lungs confirmed its selective binding to subendothelial ED-A⁺ Fn (26,37). Moreover, targeted delivery of IL-9 via the F8 antibody has shown to provoke beneficial effects in this model, among others, also on pulmonary vascular remodelling processes (25,38). Finally, F8-IL-9 fusion proteins have been shown to reduce inflammation and pulmonary remodelling in experimental PH (25,38) while F8 alone attenuates macrophage infiltration and improves RV function (26). Given the central role of ED-A⁺ Fn in vascular remodelling, its targeted inhibition, without additional payload, may offer a novel, causative treatment approach in PH, also for the groups 2 and 3, in which specific therapies are not available yet. ED-A⁺ Fn serum levels correlated with RVP_sys and pulmonary histopathology (26), suggesting a role in stratifying disease severity and guiding antibody dosage to neutralize both, circulating and tissue-bound ED-A⁺ Fn effectively. The successful application of F8 in a phase Ib trial in rheumatoid arthritis, where s.c. administration showed clinical benefit without major toxicity (39), further underscores its translational potential. In addition, the fully human anti-inflammatory fusion protein F8-IL10 (Dekavil) effectively inhibited the progression of established arthritis in the collagen-induced mouse model (40,41).

Conclusions

In conclusion, we could show ED-A⁺ Fn to play a certain functional role in PH induction using the SuHx model, qualifying the molecule as a promising diagnostic biomarker and therapeutic target with the potential to mitigate detrimental PH associated tissue remodelling, for instance, through the administration of specific neutralizing antibodies.

Acknowledgments

The authors would like to thank Dr. A.F. Muro [International Centre of Genetic Engineering and Biotechnology (ICGEB), Trieste, Italy] for providing the ED-A⁺ Fn knockout mice and Mrs. Annett Schmidt for excellent technical assistance.

Footnote

Reporting Checklist: The authors have completed the ARRIVE reporting checklist. Available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-443/rc

Data Sharing Statement: Available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-443/dss

Peer Review File: Available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-443/prf

Funding: None.

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://cdt.amegroups.com/article/view/10.21037/cdt-2025-443/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. The experimental protocol received approval from the Thuringian State Office for Food Safety and Consumer Protection (TLLV, Bad Langensalza, Germany) (registration number: UKJ21-010). All experimental procedures were carried out in accordance with the guidelines for the care and use of laboratory animals (National Research Council Committee, Guide for the Care and Use of Laboratory Animals, 2011) and the current version of the German Animal Welfare Act and applicable animal husbandry guidelines.

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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