Pleural Effusion Formation Linked to Altered Transporter Expression Involved in Alveolar Fluid Clearance: Insights From the Monocrotaline Model of Pulmonary Hypertension

ABSTRACT

Pleural effusions (PLEF) in pulmonary arterial hypertension (PAH), particularly in patients with isolated right heart failure, are associated with poor prognosis and increased mortality. This study investigates changes in alveolar fluid clearance (AFC) transporter expression in relation to lung fluid accumulation and PLEF formation during PAH progression, as well as the effects of terbutaline (TER) and riociguat (RIO) treatment. Using a monocrotaline (MCT)‐induced pulmonary hypertension (PH) rat model, we performed a detailed molecular analysis of AFC transporter expression at different disease stages, both before and after PH development. Although only minor changes were observed in the early stages prior to PH onset, a downregulation of key transporters, γ‐ENaC and Na⁺/K⁺‐ATPase subunits Atp1a2 and Atp1b1, was evident in the later stages. This reduction may have contributed to pulmonary oedema, as indicated by histological analysis. TER treatment modestly increased Atp1a2 expression, aligning with the stimulatory effects of β₂‐agonist on oedema clearance. Conversely, RIO showed trends towards fluid accumulation, indicated by perivascular oedema in control animals and reduced oxygen saturation in MCT‐treated rats. These findings support a potential role of impaired AFC in the pathogenesis of PLEF in PAH and suggest that pharmacological interventions may differentially affect lung fluid homeostasis in this setting.

1.

Summary.

• Pleural effusion, or lung fluid buildup, is a serious complication in pulmonary arterial hypertension (PAH), worsening symptoms and survival rates.

• In a rat model, we examined changes in the expression of proteins responsible for fluid clearance from alveoli as PAH progressed.

• We observed altered expression of key proteins involved in fluid removal, which may contribute to fluid buildup as the disease advances.

• Two treatments were tested: one slightly improved expression of these proteins, whereas the other showed trends towards increased lung fluid accumulation.

• Understanding these changes could guide research towards more effective ways to manage PAH complications.

2. Introduction

Pulmonary arterial hypertension (PAH) is a relatively rare, progressive disease characterized by pathological remodelling of the pulmonary vascular bed, which increases pulmonary arterial pressure and vascular resistance. These changes lead to compensatory right ventricular (RV) remodelling and ultimately progress to right heart failure (RHF) [1]. Despite increased awareness and the availability of expanding therapeutic options in recent decades, the long‐term prognosis for patients with PAH remains poor. Even with treatment, the 5‐year survival rate is approximately around 60% [2].

In recent years, several studies have investigated the incidence of pleural effusions (PLEF) in patients with idiopathic or hereditary PAH (14%) [3], PAH associated with connective tissue diseases (33%) [4] or portopulmonary hypertension (30%) [5], where no alternate explanation for their presence, aside from RHF, was identified. This is notable because PLEF in patients with congestive heart failure (CHF) are typically associated with left heart failure (LHF) [6]. Most PLEF in PAH patients were described as trace to small, right‐sided, or bilateral [3, 7] and transudative [3]. Approximately 90% of patients with idiopathic or hereditary PAH and PLEF with no alternate explanation were also diagnosed with isolated RHF. Mortality during follow‐up was significantly higher in patients with PLEF due to RHF compared to those without PLEF (64.8% vs. 27.6%, p ≤ 0.001). Based on their findings, the authors concluded that PLEF in patients with ascites and pericardial effusion likely resulted from RHF itself, rather than from ascites or pericardial effusion, as suggested by the size and localization of the PLEF [3]. However, the mechanism of PLEF formation in these patients remains unclear.

According to the literature, the primary mechanism for PLEF formation in CHF involves fluid entering the pleural space from the lung interstitium [8]. Animal experiments examining PLEF formation due to left atrial pressure elevation have shown that PLEF does not develop unless pulmonary oedema (PE) is present [9]. One of the most important mechanisms for interstitial oedema clearance is the leakage of oedema fluid through the visceral pleura into the pleural space [10, 11], which serves as a safety factor against alveolar flooding [12]. PLEF forms when the rate of pleural fluid formation exceeds the capacity of the parietal lymphatics to absorb it [8].

The formation of cardiogenic pulmonary oedema (CPE) due to LHF has traditionally been attributed to passive fluid filtration across an intact alveocapillary barrier, driven by increased hydrostatic pressure and subsequent alveolar flooding. Although fluid influx into the alveoli increases with elevated vascular pressure (due to elevated left atrial pressure), this mechanism accounts for only 30% of the total net fluid shift observed during hydrostatic PE formation [13]. According to the literature, CPE formation involves two independent components: a pressure‐dependent increase in fluid filtration across the alveocapillary barrier and, predominantly, changes in alveolar fluid transport [14].

Under physiological conditions, the driving force of alveolar fluid clearance (AFC) is primarily active transepithelial transport of sodium and fluid from the alveolar space into the interstitium across the alveolar epithelium [15], followed secondarily by chloride transport to maintain electroneutrality [16]. Apical sodium entry occurs via several sodium channels, primarily the amiloride‐sensitive epithelial sodium channel ENaC, which contributes approximately 50% of AFC in vivo [14], but also through the sodium‐glucose linked transporter SGLT1 [17] and the sodium‐coupled neutral amino acid transporters SNAT2 and SNAT5 [15]. On the basolateral side, sodium extrusion is actively driven through the Na⁺/K⁺‐ATPase [18]. Chloride and water follow the electrochemical gradient through chloride channels—predominantly via the cystic fibrosis transmembrane conductance regulator (CFTR) for chloride—and through aquaporins (AQP) and paracellular pathways for water [19], as shown in Figure 1.

FIGURE 1.

Schematic representation of alveolar fluid clearance under physiological conditions (right) compared to impaired clearance during cardiogenic pulmonary oedema (left). AC, amino acid; AT1, alveolar type I cell; AT2, alveolar type II cell; EC, endothelial cell; glc, glucose.

However, the physiological protection against alveolar flooding provided by intact AFC is largely attenuated in CPE [14, 20]. Increased hydrostatic pressure not only inhibits AFC but also reverses the direction of transepithelial fluid transport, leading to alveolar fluid secretion, which accounts for 70% of the total alveolar fluid influx under elevated hydrostatic pressure [13]. Alveolar fluid secretion results from active transepithelial chloride secretion and secondary fluid influx into the alveolar space. This secretion is primarily triggered by the inhibition of alveolar sodium absorption, particularly through cGMP‐dependent inhibition of ENaC, and secondarily mediated by CFTR and Na⁺/K⁺‐2Cl⁻ cotransporter 1 (NKCC1), providing a mechanistic explanation for the extrarenal effects of furosemide on PE [14]. Elevated lung capillary pressure has been reported to increase endothelial NO production via the PI3K/Akt signalling pathway, which regulates endothelial NO synthase [21]. NO can rapidly diffuse across the thin blood–gas barrier to alveolar epithelial cells, where it inhibits alveolar sodium and liquid absorption via cGMP‐dependent inhibition of ENaC [13]. Notably, the switch from alveolar fluid absorption (i.e., AFC) to secretion has been observed not only due to elevated hydrostatic pressure but also due to ENaC inhibition by amiloride at normal left atrial pressure [14]. Additionally, chronic CHF is associated with pulmonary vascular endothelial dysfunction, caused by dysregulation of vascular smooth muscle tone and structural remodelling [22]. This results in reduced basal production of endogenous NO in the lung tissue, particularly in patients with moderate‐to‐severe HF (corresponding to NYHA Class II or III) [23], questioning the relevance of the NO/cGMP/ENaC pathway in the context of CPE formation. All these findings highlight the need to investigate other potential mechanisms and signalling pathways that may inhibit AFC and, secondarily, contribute to PE formation.

Therefore, we hypothesized that alterations in transporters involved in AFC contribute to fluid accumulation in the lungs and the secondary formation of PLEF. The goal of this study was to provide a comprehensive molecular analysis of these transporters at two distinct stages of a monocrotaline‐induced model of PAH. The analysis included two therapeutic approaches: treatment with terbutaline (TER), a β₂‐agonist expected to enhance the vectorial transport of sodium across the alveolar epithelium and thereby improve AFC [24, 25], and treatment with riociguat (RIO), a stimulator of sGC that may mediate cGMP‐dependent inhibition of ENaC and potentially impair AFC.

3. Materials and Methods

3.1. Animal Experiment Design

A total of 124 male Wistar rats (10–12 weeks old; Department of Toxicology and Laboratory Animals Breeding, Slovak Academy of Sciences, Dobra Voda, Slovak Republic) were used in this experiment. After a 1‐week acclimatization period, the animals were randomized by body weight into six treatment groups (CON, control group; MCT, monocrotaline group; CON + TER, CON group treated with TER; MCT + TER, MCT group treated with TER; CON + RIO, CON group treated with RIO; MCT + RIO, MCT group treated with RIO). All animals were housed under standard conditions with free access to food and water ad libitum. Pulmonary hypertension was induced by a single subcutaneous dose of monocrotaline (60 mg/kg), whereas control animals received a corresponding volume of vehicle (0.9% NaCl). Starting on Day 14, the experimental animals were treated with terbutaline (10 mg/kg/day, Sigma‐Aldrich, St. Louis, Missouri, USA) or riociguat (10 mg/kg/day, Combi‐Blocks Inc., San Diego, USA) in 0.5% methylcellulose solution by oral gavage using a stainless‐steel feeding probe (Fine Science Tools, Vancouver, Canada). The CON group received only vehicle in the same manner. The dose selection was guided by an extensive review of the literature on the use of these substances in rodent experimental models, including RIO [26]. For TER, which had not been previously tested in PH models, the dose was extrapolated from human therapeutic regimens and adjusted for use in rodents [27]. According to previous studies with this model, PAH develops in experimental animals approximately 3 weeks after monocrotaline administration, as confirmed by RV catheterization [28]. To exclude effects related to MCT administration occurring before the development of PH, untreated CON and MCT groups were sacrificed 1 and 2 weeks after MCT or vehicle application. Starting around Day 21, the health status of the experimental animals was monitored twice daily by two independent examiners. Experimental animals displaying signs of acute health deterioration due to rapid PH progression—such as a loss of body weight exceeding 10 g within 24 h, pronounced dyspnoea, low temperature of distal body parts, bristled fur or apathy—were prematurely sacrificed (on Day 23.5 ± 0.3 on average) to minimize suffering. The animals in the CON and MCT groups without signs of acute health deterioration were sacrificed on Day 28, as planned. Thus, according to disease progression, MCT groups were further subdivided into the MCT_4W group—sacrificed 4 weeks after MCT application—and the prematurely terminated MCT group (ptMCT), sacrificed upon acute PAH deterioration.

Haemoglobin oxygen saturation and breath rate measurements were performed with a pulse oximeter (MouseOx, Star Life Sciences, Oakmont, USA) 24 h before sacrifice or shortly before sacrifice in the ptMCT groups. A collar sensor of appropriate size was used to monitor and record vital functions in conscious animals, according to the manufacturer's instructions.

Experimental animals were sacrificed in a CO₂ atmosphere by asphyxiation. The abdominal cavity was opened, and blood samples were collected from the inferior vena cava into EDTA‐treated tubes and immediately centrifuged (1300 g for 15 min at 4°C). Aliquots of the obtained plasma were stored at −80°C until further processing. After blood samples were collected, the pleural cavities were opened, and the presence, size, colour and location of PLEF were noted. The heart and lungs were subsequently harvested and weighed. The heart was dissected to separate the RV free wall from the left ventricle (LV) and the septum (S), allowing assessment of RV hypertrophy by Fulton's index (RV/(LV + S)). The samples from the right lung and RV were snap‐frozen in liquid nitrogen and stored at −80°C for further RT‐qPCR and western blot analysis. The left lung was immediately fixed in 10% formalin for at least 24 h at room temperature and then embedded in paraffin for histological analysis.

A schematic depiction of the experiment is shown in Figure 2. The experimental design, including the order of treatments and measurements performed on experimental animals, was predefined before the study began. To ensure animal welfare, the study was conducted without blinding to allow for prompt identification and early sacrifice of animals showing severe health deterioration due to rapid PH progression, thereby minimizing their suffering.

FIGURE 2.

Schematic illustration of experimental design.

The study was conducted in accordance with the Basic & Clinical Pharmacology & Toxicology policy for experimental and clinical studies [29]. All experimental procedures involving the use of experimental animals were approved by the Ethical Committee of the Faculty of Pharmacy, Comenius University in Bratislava, Slovak Republic, and by the State Veterinary and Food Administration of the Slovak Republic under No. 108/18‐221/3.

3.2. Total RNA Isolation and Quantitative Real‐Time PCR

Quantitative real‐time PCR (RT‐qPCR) was performed on total RNA isolated from lung and RV tissue samples, as previously described [30, 31]. Target‐specific primers were designed using Primer‐BLAST to detect mRNA and were verified to yield a single PCR product with the correct molecular weight (Table S1). The relative expression ratio of the studied genes was calculated according to Pfaffl [32] using hypoxanthine phosphoribosyltransferase 1 (Hprt1) as the endogenous reference gene and calibrating to the appropriate control group.

3.3. Western Blotting

Frozen samples of the middle right lung lobe were homogenized in liquid nitrogen and lysed in Tris–HCl buffer containing protease inhibitors. The protein concentration was measured using the Pierce BCA Protein Assay Kit (Thermo Scientific). After adjusting to the same protein concentration, lysates were solubilized and denatured in Laemmli sample buffer at 95°C for 10 min. Six samples were randomly selected from each experimental group based on the capacity of the available equipment and the number of experimental groups. Subsequently, samples were subjected to sodium dodecyl sulphate‐polyacrylamide gel electrophoresis, and after separation, proteins were transferred to a polyvinylidene fluoride membrane (Immobilon P, Millipore Corporation, USA). Membranes were blocked with 5% non‐fat dried milk in TBST for 1 h at room temperature and subsequently incubated with primary antibodies against the subunits of ENaC (Scnn1a [#bs‐2957R, Bioss], Scnn1b [#ASC‐019, Alomone Labs], Scnn1g [#PA5‐110347, Invitrogen]), the subunits of Na⁺/K⁺‐ATPase (Atp1a1 [#MA3928, Invitrogen], Atp1a2 [#ARC2458, Invitrogen], Atp1b1 [#PA5‐102831, Invitrogen]) and Aqp3 [#PA5‐78811, Invitrogen]. Afterwards, membranes were incubated with appropriate HRP‐conjugated secondary antibodies, specifically goat‐anti‐rabbit IgG (#A6154, Sigma‐Aldrich) and rabbit‐anti‐mouse IgG (#P0260, Dako). Signals were developed using enhanced chemiluminescence (Immobilon Crescendo Western HRP substrate, Millipore) and detected with a chemiluminescence imaging system (UVITEC Imaging Systems, Nine Alliance, Cambridge). Band densities were quantified using densitometry in the UVITEC Imaging Systems software (Nine Alliance, Cambridge). These arbitrary density values were normalized to total protein content determined by scanning densitometry of Ponceau S staining of membranes.

3.4. Histological Analysis

Lung tissue samples were routinely prepared for histological analysis as previously described [33]. Formalin‐fixed, paraffin‐embedded samples were cut into 5‐μm sections. Slides were stained with haematoxylin and eosin for basic histological evaluation, in addition to periodic acid‐Schiff (PAS) staining and Grocott methenamine‐silver (GMS) staining to rule out fungal infection and Gram staining to exclude bacterial infection (none were observed). The slides were evaluated using an ECLIPSE 80i microscope (Nikon, Tokyo, Japan), and images were captured via the built‐in digital camera.

For the quantitative assessment of vascular remodelling, haematoxylin and eosin‐stained lung sections were anonymized prior to evaluation. Arterioles were selected according to established morphological criteria: Both the longest and shortest luminal diameters had to fall within the range of 30–100 μm, which additionally had to exhibit a complete circular lumen, a length‐to‐width ratio below 2.0 and no more than one‐third circumferential wall collapse [34, 35]. The following structural parameters were measured using OPTIKA PROView imaging software (OPTIKA S.r.l., Italy): the longest and shortest luminal diameters, average wall thickness (measured at two points along the short axis to reduce the influence of oblique sectioning), lumen area and total vessel area. Based on these measurements, the percentage of wall area (PWA) was calculated according to the formula: PWA = ((total area − lumen area)/total area) × 100, as described previously [36]. This parameter was used to quantify the degree of vascular wall thickening and remodelling.

3.5. Outcome Measures

The outcome measures in this study included biometric and clinical parameters, such as changes in body weight, lung weight and RV hypertrophy (assessed by normalized RV weight and Fulton's index) due to MCT administration, as well as the incidence of PLEF, haemoglobin oxygen saturation and ventilation rate. Markers of pathological hypertrophy and heart failure were evaluated by assessing mRNA expression of atrial and brain natriuretic peptides (ANP and BNP), encoded by Nppa and Nppb genes, and myosin heavy chain six and seven, encoded by Myh6 and Myh7 genes, in the RV. Additionally, histopathological analysis and the expression of transporters related to AFC in the lungs were examined, including sodium transporters (Scnn1a, Scnn1g, Scnn1b, Atp1a1, Atp1a2, Atp1b1, Snat2, Snat5 and Sglt1), chloride transporters (Cftr and Nkcc1) and aquaporins (Aqp3, Aqp4 and Aqp5).

3.6. Sample Size Determination

The sample size was determined based on multiple outcome measures, particularly mRNA and protein expression of biomarkers and transporters. A fixed‐effect two‐way ANOVA design was used, with monocrotaline‐induced pulmonary hypertension as Factor A (three levels: CON, MCT_4W, ptMCT) and treatment as Factor B (two levels: untreated, treated), resulting in six groups. A priori power analysis was conducted using G*Power Version 3.1.9.7, targeting 80% power at a 0.05 significance level. A large effect size (0.40) was selected based on pilot data. Assumptions included independence of observations, homogeneity of variances and normality of residuals. The required total sample size was n = 64, with at least seven animals per group. The sample size adhered to the principles of the 3Rs (replacement, reduction and refinement), ensuring ethical and responsible use of experimental animals.

3.7. Statistical Analysis

Data were reported as mean ± standard error of the mean (SEM). The Shapiro–Wilk normality test was used to assess the normal distribution of data. Differences among normally distributed data were evaluated by the Student t‐test, one‐way or two‐way analysis of variance (ANOVA), followed by appropriate post hoc tests. For non‐normally distributed data, the Mann–Whitney and Kruskal–Wallis tests were applied. The ROUT method served as an exclusion criterion for outliers. For correlation analyses, the Pearson correlation coefficient was calculated for normally distributed pairs of variables, whereas Spearman correlation was used otherwise. In the case of PLEF incidence, Fisher's exact test and the chi‐square test were used to assess differences in the distribution of a categorical variable between groups. Relative risk and relative risk reduction were also calculated. A value of p < 0.05 was considered statistically significant. All data were analysed using GraphPad Prism 10.2.2 (GraphPad Software, San Diego, CA, USA).

4. Results

4.1. Biometric Parameters and Clinical Characterization

Monocrotaline administration decreased the body weight of experimental animals, particularly in ptMCT groups; this effect was not influenced by TER or RIO treatment. The RV, normalized RV weight and Fulton's index were significantly increased in both untreated and treated MCT groups, with greater RV mass and hypertrophy in end‐stage disease (vs. MCT_4W, p < 0.05). LV weight did not increase under any intervention.

Monocrotaline administration also led to an increase in lung weight, starting from Week 2 after MCT administration (Table S2), which was more pronounced in end‐stage disease (vs. MCT_4W, p < 0.05). In general, TER treatment produced a slightly smaller increase in lung weight (by 43.3% vs. CON, p < 0.05) compared to untreated MCT groups (by 50.0% vs. CON, p < 0.05), whereas RIO treatment resulted in a slightly greater increase in lung weight (by 59.9% vs. CON, p < 0.05). This observation appeared as a trend without a significant difference between the mentioned MCT groups. Furthermore, the difference in lung weight between ptMCT + TER and ptMCT + RIO groups was close to statistical significance (p = 0.0641).

Haemoglobin oxygen saturation was decreased in all MCT groups, including those treated with TER and RIO, and this decrease was compensated by an increased ventilation rate, except in the TER‐treated group. Furthermore, RIO treatment significantly increased heart rate in both CON and MCT groups. All mentioned biometric parameters and other clinical characterizations are summarized in Table 1.

TABLE 1.

Clinical characterics and selected biometric parameters of experimental animals.

	Untreated			Treated with TER			Treated with RIO
	CON	MCT4W	ptMCT	CON	MCT4W	ptMCT	CON	MCT4W	ptMCT
n	10	10	10	10	9	8	10	9	9
Incidence of PLEF	0	2	3	0	1	1	0	2	4
p (CON vs. MCT)	0.1400			0.5157			0.0619
Gravimetric parameters
Body weight (g)	384 ± 7	319 ± 8*	308 ± 8*	378 ± 11	333 ± 9*+	301 ± 6*+−	361 ± 10	343 ± 8*	315 ± 12*○
Lung weight (g)	1.70 ± 0.09	2.33 ± 0.15*	2.78 ± 0.12*#	1.74 ± 0.06	2.22 ± 0.08*+	2.69 ± 0.16*+	1.68 ± 0.12	2.36 ± 0.07*○	3.09 ± 0.16*○●
Lung/BW (mg/g)	4.44 ± 0.24	7.30 ± 0.41*	9.08 ± 0.49*#	4.58 ± 0.20	6.69 ± 0.22*+	8.98 ± 0.61*+−	4.67 ± 0.30	6.92 ± 0.33*○	9.98 ± 0.70*○●
Heart weight (g)	1.15 ± 0.03	1.24 ± 0.06	1.37 ± 0.05*	1.23 ± 0.04	1.24 ± 0.09	1.36 ± 0.06	1.10 ± 0.03	1.22 ± 0.05	1.32 ± 0.05*○
Heart/BW (mg/g)	3.00 ± 0.04	3.92 ± 0.23*	4.44 ± 0.11*	3.26 ± 0.07	3.72 ± 0.23*	4.52 ± 0.20*+−	3.05 ± 0.06	3.54 ± 0.09*○	4.22 ± 0.10*○●
RV weight (g)	0.24 ± 0.01	0.35 ± 0.03*	0.44 ± 0.02*#	0.26 ± 0.01	0.33 ± 0.04*+	0.41 ± 0.01*+−	0.23 ± 0.01	0.34 ± 0.02*○	0.40 ± 0.03*○
RV/BW (mg/g)	0.61 ± 0.01	1.10 ± 0.10*	1.44 ± 0.06*#	0.68 ± 0.03	1.00 ± 0.11*+	1.36 ± 0.06*+−	0.63 ± 0.02	0.98 ± 0.06*○	1.26 ± 0.10*○●
LV weight (g)	0.81 ± 0.02	0.73 ± 0.03	0.73 ± 0.03	0.84 ± 0.03	0.76 ± 0.04	0.74 ± 0.03+	0.76 ± 0.02	0.73 ± 0.03	0.71 ± 0.05
LV/BW (mg/g)	2.10 ± 0.02	2.29 ± 0.07*	2.37 ± 0.05*	2.23 ± 0.04	2.27 ± 0.06	2.45 ± 0.08+	2.10 ± 0.04	2.13 ± 0.05	2.24 ± 0.13
Fulton index (%)	29.20 ± 0.41	47.83 ± 3.83*	60.99 ± 3.30*#	30.54 ± 1.01	43.57 ± 3.83*+	56.14 ± 4.04*+−	30.03 ± 0.75	46.26 ± 2.68*○	58.10 ± 5.69*○●

Vital funtions
	CON	MCT	CON	MCT	CON	MCT
	Untreated		Treated with TER		Treated with RIO
Heart rate (min−1)	397 ± 7	456 ± 7*	395 ± 17	423 ± 8 ♦	443 ± 20*	457 ± 10*
O2 saturation (%)	94.1 ± 0.2	91.8 ± 0.6*	93.1 ± 0.5	89.8 ± 0.6*+	93.9 ± 0.3	89.7 ± 1.0*
Breath rate (min−1)	103.6 ± 2.7	134.5 ± 7.0*	99.3 ± 4.2	124.1 ± 8.9+	112.4 ± 3.8	130.9 ± 6.8*○♦

Note: Vital function measurements could not be performed in some experimental animals due to unexpected sudden health deterioration; therefore, the measured values are presented regardless of disease progression. Statistical significance was assessed for each therapeutic treatment individually compared to the untreated groups. Data are presented as mean ± SEM (*, p < 0.05 vs. CON; #, p < 0.05 vs. MCT _4W ; §, p < 0.05 vs. ptMCT; +, p < 0.05 vs. CON + TER; −, p < 0.05 vs. MCT _4W   + TER; ○, p < 0.05 vs. CON + RIO; ●, p < 0.05 vs. MCT _4W   + RIO; ♦, p < 0.5 vs. MCT).

The PLEF sizes observed in experimental animals ranged from trace to small in untreated and TER‐treated groups and were small in RIO‐treated groups, pale straw‐coloured and bilateral in distribution. The presence of pericardial effusions and ascites was not detected. The overall incidence of PLEF in the experimental model, regardless of therapy, was 23.6% (p < 0.05). Among experimental animals with MCT‐induced PH and PLEF, 61.5% were in end‐stage disease (ns). The incidence of PLEF in RIO‐treated groups was close to statistical significance (p = 0.0619; Table 1). In the RIO‐treated MCT group, significant differences in lung weight were observed between animals with and without PLEF (by 20.9%, p < 0.05), whereas such differences were not noted in the untreated or TER‐treated MCT groups. Additionally, the RIO‐treated MCT group with PLEF showed a significant increase in lung weight (p < 0.05) and a significant decrease in haemoglobin oxygen saturation (p < 0.05) compared to the untreated MCT group with PLEF (Table S3). Notably, the risk of PLEF in the TER‐treated group was reduced by 52.9% compared to the untreated MCT group (p = 0.247), although this reduction did not reach statistical significance.

4.2. Markers of Pathological Hypertrophy and Heart Failure

A 10‐fold increase in Nppb mRNA expression was observed in the RV in the MCT_4W group (vs. CON, p < 0.05); however, its significance did not change with higher disease severity (ptMCT vs. MCT_4W). A similar trend was noted for Nppa mRNA expression in the RV. Moreover, we observed downregulation of Myh6 mRNA expression, with a more pronounced decrease in the end‐stage disease (vs. MCT_4W, p < 0.05). Additionally, Myh7 mRNA expression was upregulated regardless of disease progression (Figure 3). The relative expression of Myh6 was significantly improved by TER treatment (two‐way ANOVA, p < 0.05), and the relative expression of Nppa and Myh6 also improved due to RIO treatment (two‐way ANOVA, p < 0.05) in the RV (Figure S1).

FIGURE 3.

Relative mRNA expression of Nppa (A), Nppb (B), Myh6 (C) and Myh7 (D) in RV in untreated control (CON) and monocrotaline (MCT) groups. Data are presented as mean ± SEM; n = 10 per group; *p < 0.05.

4.3. Histopathological Analysis

Lung histology performed 4 weeks after MCT application or following acute PAH deterioration revealed a noticeable increase in the medial thickness of small pulmonary arterioles in the MCT group compared to healthy controls. In addition to structural remodelling, perivascular and interstitial lung macrophage infiltration indicated the development of pulmonary inflammation. Additionally, MCT‐treated animals exhibited interstitial and alveolar oedema.

Therapeutic intervention with either TER or RIO resulted in reduced inflammatory infiltrates, notably in ptMCT animals. Interestingly, RIO treatment alone induced perivascular interstitial oedema, which was less apparent with MCT administration. RIO treatment in both the CON and MCT groups induced perivascular interstitial oedema; however, it reduced inflammation and macrophage infiltration in the MCT group compared to the untreated MCT group (Figure 4).

FIGURE 4.

Pulmonary tissue histology from experimental animals with pulmonary hypertension induced by monocrotaline (MCT) administration, treated with riociguat (RIO) or terbutaline (TER). Administration of MTC resulted in mild thickening of walls of arteries (a), slight increase in cellularity of the parenchyma, which was more pronounced by increased accumulation of alveolar macrophages and perivascular lymphocyte infiltrations accompanied by alveolar edematous fluid in prematurely deceased animals (ptMCT). Treatment with both TER and RIO lead to a reduction of inflammatory infiltrate, especially in ptMCT animals. Note the perivascular tissue edema (*) in control group treated with RIO (CON + RIO) and (b) bronchus. Haematoxylin and eosin, 100×, n = 6 per group.

Quantitative assessment of pulmonary vascular remodelling based on PWA of small arterioles (Table S4) revealed a significant main effect of PH as well as a significant effect of both TER and RIO treatments, as confirmed by two‐way ANOVA (p < 0.05), even though the increase in PWA in untreated MCT groups compared to controls did not reach statistical significance. Notably, PWA was significantly reduced in TER‐treated healthy controls compared to untreated controls (Tukey's post hoc test, p < 0.05), suggesting a potential modulatory effect of TER on arteriolar wall structure.

4.4. Transporters Involved in AFC

The expression of transporters responsible for active transepithelial sodium transport was assessed in the lungs. The γ subunit of ENaC, encoded by Scnn1g, which plays a critical role in clearing oedema fluid [37], was found to be reduced at the protein level in both the untreated MCT_4W and ptMCT groups (Figure 5E). Pearson correlation analysis showed that the relative Scnn1g mRNA expression in the lungs negatively correlated with lung weight and Fulton's index (Figure S2A,B), as well as with the relative expression of ANP and BNP, encoded by Nppa and Nppb, in the RV (Figure S2C,D). Additionally, it positively correlated with Myh6 mRNA expression in the RV (Figure S2E). No correlation was observed with haemoglobin oxygen saturation (data not shown). However, these findings were not confirmed at the protein level (Figure S3). In contrast, we observed an upregulation of the β‐ENaC subunit, encoded by Scnn1b, at the protein level in the untreated ptMCT group (Figure 5C), whereas α‐ENaC, encoded by Scnn1a, remained unchanged (Figure 5A).

FIGURE 5.

Relative protein expression of Scnn1a (A), Scnn1b (C) and Scnn1g (E) with representative immunoblot and total protein staining (B,D,F) in lungs. Data are presented as mean ± SEM; n = 6 per group; *p < 0.05.

Apical sodium entry is also mediated by Snat2, Snat5 and Sglt1. Although the relative expression of Snat2 remained unchanged across the untreated diseased groups in the lungs (Figure S8), Snat5 expression was significantly decreased in the untreated ptMCT group (Figure S4). The expression of the Sglt1 gene did not reach the detection limit of the qRT‐PCR assay.

Lung liquid clearance is actively driven by the osmotic gradient generated by the Na⁺/K⁺‐ATPase. Alveolar cells express the α₁, α₂ [38] and β₁ [39] subunits of Na⁺/K⁺‐ATPase, whose expression was examined in lung tissue. The expression of the catalytic α₁ subunit (encoded by the Atp1a1 gene) was not detected at the protein level in lungs. However, the expression of the α₂ subunit (encoded by Atp1a2) decreased in untreated MCT groups. TER treatment significantly influenced the relative protein expression of Atp1a2 (two‐way ANOVA, p = 0.0250), most notably by upregulating it in the TER‐treated CON group by approximately 50% compared to the untreated CON group (Figure 6A). The relative protein expression of the β₁ subunit (encoded by Atp1b1) was also downregulated in untreated MCT groups; however, with no significant effect of TER treatment (Figure 6B), we did not observe any significant effect of RIO treatment on either Atp1a2 or Atp1b1 expression (Figure S5).

FIGURE 6.

Relative protein expression of Atp1a2 (A) in untreated and terbutaline (TER) treated monocrotaline (MCT) groups and Atp1b1 (B) in untreated MCT groups with representative immunoblot and total protein staining (C,D) in lung tissue. Data are presented as mean ± SEM; n = 6 per group; *p < 0.05.

In the case of chloride channels, the mRNA expression of Cftr was significantly upregulated in the lungs (Figure S7A), whereas the expression of Nkcc1 remained unchanged across the untreated diseased groups (Figure S7B).

Water transport across pulmonary epithelium is facilitated by aquaporins, namely, Aqp3, Aqp4 and Aqp5. In the untreated diseased groups, we observed a decrease in Aqp3 protein expression, along with reduced mRNA expression of Aqp4 and Aqp5 (Figure S6).

TER treatment normalized the increased expression of Cftr in the MCT_4W and ptMCT groups (Figure S7A) and raised the relative expression of Nkcc1 in the CON and MCT_4W groups (Figure S7B). Moreover, an upregulation of Scnn1g mRNA was noted in the MCT_4W group in response to TER treatment (Figure S7C), although this did not manifest at the protein level (Figure S7D). The expression levels of other previously discussed genes involved in AFC in TER‐treated groups followed a similar pattern to those in untreated groups.

Treatment with RIO resulted in a significant reduction in the relative expression of Snat2 by approximately 50% in the CON, MCT_4W and ptMCT groups (Figure S8). The expression levels of other genes linked to AFC in the RIO‐treated groups showed a similar pattern as in the untreated groups.

Additionally, we did not observe any alterations in the mRNA expression of transporters involved in AFC at 1 and 2 weeks after MCT administration, except for Atp1a2, which showed gradual downregulation beginning in Week 1, and Aqp5, which was downregulated starting in Week 2. For Scnn1a, a slight 10.8% downregulation was observed in Week 1, but this change did not persist into Week 2 (Figure S9).

5. Discussion

This study provides an in‐depth molecular analysis of the transporters involved in AFC in the lungs of a rat model of monocrotaline‐induced PAH, examined at two distinct stages of disease progression. Our hypothesis was based on the assumption that changes in AFC transporter expression during disease progression contribute to fluid accumulation in the lungs and the subsequent formation of PLEF. Additionally, we investigated the impact of different therapeutic approaches, specifically terbutaline and riociguat, on the expression of these transporters.

The rat MCT model is widely utilized in vivo model of PAH, replicating many features of human PAH, including endothelial cell damage in the pulmonary vasculature, subsequent vascular remodelling and pulmonary arterial medial hypertrophy [40]. However, PH development in this model is delayed, occurring approximately 3–4 weeks after a single MCT exposure [28, 41], despite MCT and its metabolites being rapidly metabolized and cleared from the circulation [42]. In addition to causing endothelial cell damage, MCT is also reported to induce acute lung injury [43], resulting in oedema fluid accumulation due to altered microvascular endothelial cell permeability, evident as early as 1‐week post‐administration [44]. For example, in mice, multiple signs of acute lung injury were observed within the first 7–10 days after MCT pyrrole injection, followed by resorption of lung oedema by Days 14–21, with slightly elevated RV systolic pressure 28‐day post‐injection, yet without the development of PH [45].

It is important to note that MCT toxicity depends not only on dose but also varies among species [44]. Thus, we previously analysed selected biometric parameters and the relative mRNA expression of transporters involved in AFC in untreated MCT groups, not only after PH development but also at 1 and 2 weeks' post‐MCT administration, prior to PH development, to differentiate between the acute effects of MCT in early stages and the effects linked to PH progression. Notably, lung weight gradually increased starting from Week 2 after MCT administration. Additionally, we did not observe any alterations in the expression of transporters involved in AFC at 1 and 2 weeks' post‐MCT administration, except for the Atp1a2 gene, which showed gradual downregulation starting in Week 1, and the Aqp5 gene, which began downregulation in Week 2.

The experimental groups with developed PH, namely, the MCT_4W and ptMCT groups, represent distinct stages of the disease and, as expected, differ in the severity of PH symptoms and the extent of target organ damage. The primary determinant of functional state and prognosis in patients with PAH is the RV [46]. In this model, we observed increases in both RV weight and Fulton's index, which were more pronounced in the ptMCT group, indicating a greater degree of RV hypertrophy in this stage.

Although pulmonary vascular remodelling is a hallmark of MCT‐induced PH, the addition of PWA measurements revealed only non‐significant increases in the untreated MCT groups compared to controls. This may reflect the relatively modest degree of structural remodelling in our model at the time points assessed, particularly given the heterogeneity of arteriolar changes in this disease. Notably, PWA was significantly reduced in TER‐treated control animals, suggesting a potential effect of β₂‐adrenergic stimulation on pulmonary arteriolar wall structure in healthy lungs. Furthermore, the ptMCT rats were, in some cases, sacrificed at slightly earlier time points, which might introduce variability in the measured PWA values. Alternatively, the modest arteriolar remodelling phenotype observed in our study may be attributed to strain‐dependent differences in vascular response, possibly associated with the specific substrain of Wistar rats used. Importantly, the absence of overt arteriolar wall thickening in untreated groups during disease progression suggests that alterations in AFC, oedema and PLEF may occur before or partially independently of advanced vascular remodelling, highlighting the complex nature of pathophysiological changes in this model.

The increase in lung weight, particularly pronounced in the ptMCT group, was likely caused by greater accumulation of oedematous fluid, as indicated by histological analysis of lung tissue. In this experimental model, we observed a slightly higher incidence of PLEF (23.6%) than reported in patients with idiopathic or hereditary PAH (14%) [3]. The majority of experimental animals with PLEF (61.5%) were in end‐stage disease, which was associated with greater RV hypertrophy, whereas approximately 90% of PAH patients with PLEF were co‐diagnosed with isolated RHF [3]. However, it is important to note that in the current model, end‐stage disease was defined by acute health deterioration, not by a diagnosis of RHF.

The γ subunit of ENaC is critical for oedema fluid to exert its maximal effect on net fluid absorption by distal lung epithelia [37]; it was found to be downregulated in both MCT_4W and ptMCT groups. Furthermore, we identified a correlation between the relative mRNA expression of the Scnn1g gene not only with increased lung weight but also with Fulton's index and the relative expressions of Nppa, Nppb and Myh6 in the RV. This suggests a connection not only with lung fluid accumulation but also with RV state.

It is known that alveolar sodium and liquid absorption may be suppressed by a cGMP‐dependent inhibition of ENaC [14]. Additionally, experiments using Xenopus 2F3 cells have demonstrated that natriuretic peptide receptor‐A (NPR‐A) is involved in the natriuretic peptide‐mediated regulation of ENaC via cGMP signalling [47]. NPR‐A and NPR‐C were shown to be present in both bronchial and alveolar cells [48]. These studies primarily focused on ENaC activity as an immediate response to given stimuli. However, the long‐term exposure to the same stimulus can lead to changes in the expression of targeted protein. This might explain the observed changes in Scnn1g expression in the experimental model.

Nevertheless, these correlations did not translate to protein expression levels of γ‐ENaC in the lungs. The downregulation of γ‐ENaC was accompanied by an upregulation of the β‐ENaC subunit. Studies using transgenic mice with airway‐specific overexpression of ENaC have shown that Scnn1b overexpression alone—unlike Scnn1a or Scnn1c—increased both basal and amiloride‐sensitive currents across freshly excised tracheal tissues in neonatal and adult Scnn1 transgenic mice compared to their wild‐type counterparts. This indicates that the β subunit of ENaC is rate‐limiting for airway Na⁺ absorption in vivo. However, β‐ENaC overexpression in experimental animals resulted in airway surface liquid depletion, increased postnatal mortality and a phenotype closely resembling the clinical and pathological features of early cystic fibrosis lung disease [49]. We did not observe any significant difference in protein expression of α‐ENaC.

In lung tissue, the α₂ subunit of Na⁺/K⁺‐ATPase is not only expressed in alveolar cells [38] but is preferentially localized in pulmonary vascular smooth muscle cells [50]. Montani et al. proposed a potential association between PAH and channelopathy due to a mutation of the Atp1a2 gene, which is known to cause familial hemiplegic migraine, a rare autosomal dominant disease. It was suggested that mutations in the Atp1a2 gene may contribute to pulmonary arterial remodelling and the development of PAH by disrupting intracellular calcium and potassium concentrations [51].

Given the observed downregulation of Atp1a2 starting from the first week after MCT administration, we propose that this downregulation plays a role in the development of PH in our model. Additionally, we noted a downregulation of the β₁ subunit of Na⁺/K⁺‐ATPase, which is considered essential for restoring AFC in both non‐cardiogenic [52] and cardiogenic [53] pulmonary oedema, as experimentally demonstrated by increased AFC following Atp1b1 overexpression.

The primary driving force of AFC is considered to be the active transepithelial transport of sodium, which facilitates water movement across the alveolar epithelium [15]. Based on calculations of alveolar epithelial membrane potentials and a basal conductance ratio for sodium and potassium, it can be predicted that a partial inhibition of sodium conductance greater than 26.4% is sufficient to reverse the direction of chloride flux, shifting from transepithelial chloride absorption to secretion (calculated according to Solymosi et al. [14]; Supporting Information). This effect is further amplified when sodium uptake is fully blocked [14].

However, despite observing changes in the expression of sodium transporters involved in alveolar transepithelial transport, we cannot precisely predict the rate of sodium current inhibition. Histological analysis, however, revealed the development of interstitial and alveolar oedema in MCT_4W and ptMCT groups, suggesting that the observed changes in expression may have contributed to pulmonary oedema formation.

As previously mentioned, alveolar chloride and fluid secretion, triggered by inhibition of sodium flux, is mediated by CFTR and NKCC1 [14]. Although no changes in NKCC1 expression were observed in untreated MCT groups, we did find a significant upregulation of CFTR. According to the literature, inhibiting CFTR or NKCC1 effectively suppresses alveolar fluid secretion, and CFTR knockout mice have been shown to be protected from developing PE [14].

Aqp5 represents the primary water flux channel in alveolar epithelia [54], and it was found to be downregulated in the MCT model. Significant downregulation of Aqp5 has previously been reported in response to hypoxic stress in mice [55]. However, studies involving AQP‐null mice suggest that AQPs are not essential for AFC [56]. For instance, Aqp1 and Aqp5 knockout mice exhibited no significant impact on AFC at the adult stage [54]. This finding can be explained by water movement following the electrochemical gradient not only through AQPs but also via paracellular pathways [19].

Stimulation of β₂‐adrenergic receptors (β₂ARs) in animal models and human cells enhances vectorial transport of sodium across the alveolar epithelium, thereby improving oedema clearance [24]. β₂ARs regulate several key transporters involved in AFC including ENaC, CFTR and Na⁺/K⁺‐ATPase [57]. The effectiveness of β₂‐agonist therapy for lung oedema clearance has been demonstrated in a clinical trial, where the incidence of high‐altitude pulmonary oedema in a high‐risk group of travellers was reduced by the long‐acting β₂‐agonist salmeterol [58].

In our experimental model, we observed a slightly smaller increase in lung weight with TER treatment compared to the untreated MCT group. Moreover, TER treatment showed a trend towards reducing the risk of PLEF by 52.9% compared to the untreated MCT group, though this observed difference could not be reliably assessed due to the limited sample size.

Previous studies have shown that stimulation of βARs with TER increases the number of ENaCs and their open time in alveolar type II cells by enhancing ENaC trafficking to the cell membrane, increasing ENaC expression, and directly phosphorylating the β and γ subunits of ENaC [57]. In our study, upregulation of Scnn1g mRNA was observed in the MCT_4W group in response to TER treatment, although this was not reflected at the protein level.

Furthermore, upregulation of Na⁺/K⁺‐ATPase by β₂AR signalling has been described to primarily occur through an increased number of Na⁺/K⁺‐ATPase molecules in the cell membrane [57]. In the MCT model, relative expression of Atp1a2 was increased by TER treatment, most notably showing approximately 50% upregulation in the CON group compared to the untreated CON group.

We hypothesized that RIO, as a stimulator of sGC, may be responsible for cGMP‐dependent inhibition of ENaC and therefore negatively impact AFC. However, based on the analysis of ENaC subunit expression alone, we cannot confirm or refute this hypothesis.

In the MCT model, RIO treatment resulted in a slightly greater increase in lung weight compared to the untreated MCT group, which was observed only as a trend. Notably, a similar effect on lung weight was observed with another soluble guanylate cyclase stimulator, BAY 41‐2272, which significantly increased lung weight in a hypoxia‐induced model of PH [59]. Furthermore, the incidence of PLEF in RIO‐treated groups was close to statistical significance. Interestingly, the RIO‐treated MCT group with PLEF exhibited a significant increase in lung weight and a significant reduction in haemoglobin oxygen saturation compared to the untreated MCT group with PLEF. This effect may be attributed to increased fluid build‐up in lung tissue.

Moreover, histological analysis revealed that treatment with RIO‐induced perivascular oedema in CON animals. The summary of product characteristics for the medicinal product Adempas, which contains RIO, stated that shortness of breath during treatment may result from fluid build‐up in the lungs [60]; however, the mechanism underlying this fluid accumulation was not specified. In the MCT model, RIO treatment led to a 50% reduction in the relative expression of Snat2 in the CON, MCT_4W and ptMCT groups.

This study provides a detailed molecular analysis of AFC transporters in a rat model of MCT‐induced PAH, emphasizing the potential role of altered transporter expression in fluid accumulation and pulmonary oedema during disease progression. Key components of AFC, including γ‐ENaC and the Atp1a2 and Atp1b1 subunits of Na⁺/K⁺‐ATPase, were downregulated, potentially contributing to the pulmonary oedema observed in experimental animals. Treatment with TER increased the expression of the α₂ subunit of Na⁺/K⁺‐ATPase, a response associated in the literature with the effects of β₂ agonists on oedema clearance. In contrast, RIO treatment appeared to exacerbate fluid accumulation, as suggested by the presence of perivascular oedema in control animals, increased lung weight and reduced oxygen saturation in MCT‐treated animals.

The present study has several limitations. Although the MCT model reproduces certain features of PAH, it does not fully replicate the complexity of human PAH pathology, mainly due to the acute effects of MCT. Although we analysed the expression of transporters involved in AFC before the development of PAH in the experimental model, we cannot exclude differences in disease progression in the animal model, due to acute MCT toxicity, compared to human PAH. These limitations hinder the direct translation of our findings to human PAH and related therapeutic strategies.

The wet‐to‐dry lung weight ratio, a commonly used method for assessing pulmonary fluid accumulation, could not be performed in this study because the lungs were divided for concurrent histological and molecular analyses, preventing the collection of intact tissue for this measurement.

To reduce animal suffering, right heart catheterization was not conducted in this study. The development of PAH was instead inferred based on an established monocrotaline protocol, previously validated in a parallel study through direct haemodynamic measurements [28]. Additionally, the number of experimental animals was kept low to minimize their suffering, which may have negatively impacted the statistical analysis in cases where the observed effect had a low incidence.

Despite identifying several alterations in AFC transporter expression, PE formation and the incidence of PLEF in the MCT model, further investigation is necessary to better translate these insights to human PAH pathology and potential treatment strategies.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Table S1: Primer sequences for RT‐qPCR.

Table S2: Clinical characterics and selected biometric parameters of experimental animals 1 and 2 weeks after MCT administration. Data are presented as mean ± SEM (*p < 0.05 vs. CON).

Table S3: Clinical characterics and selected biometric parameters of experimental animals according to incidence of PLEF.

Figure S1: Relative mRNA expression of Nppa (A,B), Nppb (C,D), Myh6 (E,F) and Myh7 (G,H) in RV from experimental animals after MCT administration untreated, TER treated or RIO‐treated. Data are presented as mean ± SEM; n = 8–10 per group; *p < 0.05.

Table S4: Quantification of pulmonary vascular remodelling using percentage of wall area (PWA) in untreated controls, MCT‐exposed animals and animals treated with either TER or RIO. Arterioles were selected based on predefined morphometric criteria (diameter 30–100 μm, circular cross‐section, low collapse index). Data are presented as mean ± SEM. *p < 0.05 versus CON.

Figure S2: Correlation between relative mRNA expression of Scnn1g in lungs with selected biometric parameters (A,B), markers of ventricular dysfunction, Nppa and Nppb (C,D) and myosin heavy chain genes, Myh6 and Myh7 (E,F) expressed in RV, in untreated MCT_4W and ptMCT groups. Data are presented as mean ± SEM; n = 10 per group; *p < 0.05.

Figure S3: Correlation between relative protein expression of Scnn1g in lungs with selected biometric parameters (A,B), markers of ventricular dysfunction, Nppa and Nppb (C,D) and myosin heavy chain genes, Myh6 and Myh7 (E,F) expressed in RV, in untreated MCT_4W and ptMCT groups. Data are presented as mean ± SEM; n = 6–10 per group; *p < 0.05.

Figure S4: Relative mRNA expression of Snat5 in lung tissue of experimental animals after MCT administration. Data are presented as mean ± SEM; n = 10 per group; *p < 0.05.

Figure S5: Relative protein expression of Atp1a2 (A) and Atp1b1 (B) with representative immunoblot and total protein staining (C,D) in lung tissue from experimental animals after MCT administration untreated or RIO treated. Data are presented as mean ± SEM; n = 6 per group; *p < 0.05.

Figure S6: Relative protein expression of Aqp3 (A) with representative immunoblot and total protein staining (B) and relative mRNA expression of Aqp4 (C) and Aqp5 (D) in lung tissue of experimental animals after MCT administration. Data are presented as mean ± SEM; n = 10 per group; *p < 0.05.

Figure S7: Relative mRNA expression of Cftr (A) and Nkcc1 (B) and relative expression of Scnn1g on mRNA (C) and protein level (D) with representative immunoblot and total protein staining (E) in lung tissue from experimental animals after MCT administration untreated or TER treated. Data are presented as mean ± SEM; n = 6–10 per group; *p < 0.05.

Figure S8: Relative mRNA expression of Snat2 in lung tissue from experimental animals after MCT administration untreated or RIO treated. Data are presented as mean ± SEM; n = 9–10 per group; *p < 0.05.

Figure S9: Relative mRNA expression of Scnn1a (A), Scnn1b (B), Scnn1g (C), Snat2 (D), Snat5 (E), Atp1a2 (F), Atp1b1 (G), Cftr (H), Nkcc1 (I), Aqp3 (J), Aqp4 (K) and Aqp5 (L) in lung tissue from experimental animals 1 and 2 weeks after MCT administration. Data are presented as mean ± SEM; n = 9–10 per group; *p < 0.05.

Lelkova K., Hadova K., Katanova M., et al., “Pleural Effusion Formation Linked to Altered Transporter Expression Involved in Alveolar Fluid Clearance: Insights From the Monocrotaline Model of Pulmonary Hypertension,” Basic & Clinical Pharmacology & Toxicology 137, no. 4 (2025): e70096, 10.1111/bcpt.70096.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.