BMP type II receptor deficiency confers resistance to growth inhibition by TGF-β in pulmonary artery smooth muscle cells: role of proinflammatory cytokines

Abstract

Mutations in the bone morphogenetic protein (BMP) type II receptor (BMPR-II) underlie most cases of heritable pulmonary arterial hypertension (HPAH) and a significant proportion of sporadic cases. Pulmonary artery smooth muscle cells (PASMCs) from patients with pulmonary arterial hypertension (PAH) not only exhibit attenuated growth suppression by BMPs, but an abnormal mitogenic response to transforming growth factor (TGF)-β1. We sought to define the mechanism underlying this loss of the antiproliferative effects of TGF-β1 in BMPR-II-deficient PASMCs. The effect of TGF-β1 on PASMC proliferation was characterized in three different models of BMPR-II dysfunction: 1) HPAH PASMCs, 2) Bmpr2^+/− mouse PASMCs, and 3) control human PASMCs transfected with BMPR-II small interfering RNA. BMPR-II reduction consistently conferred insensitivity to growth inhibition by TGF-β1. This was not associated with altered canonical TGF-β1/Smad signaling but was associated with a secreted factor. Microarray analysis revealed that the transcriptional responses to TGF-β1 differed between control and HPAH PASMCs, particularly regarding genes associated with interleukins and inflammation. HPAH PASMCs exhibited enhanced IL-6 and IL-8 induction by TGF-β1, an effect reversed by NF-κB inhibition. Moreover, neutralizing antibodies to IL-6 or IL-8 restored the antiproliferative effect of TGF-β1 in HPAH PASMCs. This study establishes that BMPR-II deficiency leads to failed growth suppression by TGF-β1 in PASMCs. This effect is Smad-independent but is associated with inappropriately altered NF-κB signaling and enhanced induction of IL-6 and IL-8 expression. Our study provides a rationale to test anti-interleukin therapies as an intervention to neutralize this inappropriate response and restore the antiproliferative response to TGF-β1.

pulmonary arterial hypertension (PAH) is a life-limiting condition resulting from the remodeling of the small- to medium-sized distal pulmonary arteries, leading to elevated pulmonary vascular resistance and, ultimately, death from right heart failure (39). The molecular mechanisms underlying the pathogenesis of PAH remain poorly defined, although substantial evidence implicates dysregulated signaling by members of the transforming growth factor (TGF)-β superfamily (33, 44). For example, mutations in the gene encoding the bone morphogenetic protein (BMP) type II receptor (BMPR-II), a TGF-β superfamily receptor, underlie most cases of heritable PAH (HPAH) (23). Interestingly, fewer than half of Bmpr2 mutation carriers develop HPAH, suggesting that additional “hits,” such as inflammation, are required for disease initiation and progression (23).

TGF-β superfamily ligands bind and activate heteromeric complexes of type I and type II receptors. For example, BMPs can activate complexes comprising the type II receptor, BMPR-II, in complex with the type I receptors, ALK1, ALK2, ALK3, and ALK6. TGF-βs bind a different type II receptor, TGF-β type II receptor, in complex with the type I receptor, ALK5. Upon activation, TGF superfamily receptor complexes phosphorylate the canonical second messengers, Smads, according to the particular ligand-receptor response (1, 25). BMP ligands generally signal via Smad1, Smad5, and Smad8, whereas TGF-β1 typically activates Smad2 and Smad3. The activated Smads translocate from the cytosol to the nucleus and form complexes with other transcription factors to bind and activate the expression of target genes (1, 25). In addition, TGF-β superfamily receptors can also signal through noncanonical pathways, such as MAP kinases (49). HPAH pulmonary artery smooth muscle cells (PASMCs) from patients with defined BMPR2 mutations have reduced levels of functional BMPR-II, resulting in reduced Smad1/5/8 activation in response to BMP4 (33, 47). One important functional consequence of this is a reduced antiproliferative response to BMP4 (47).

Recent studies support a major role for TGF-β1 in the pathogenesis of PAH (33, 44, 48). We reported that PASMCs harvested from patients with idiopathic PAH, of unknown BMPR-II status, exhibit a blunted antiproliferative response to TGF-β1 (33). Furthermore, TGF-β1 is implicated in the pathogenesis of monocrotaline (MCT)-induced PAH (MCT-PAH) in rats, as three independent studies reported that small-molecule ALK5 inhibitors prevent and reverse the pulmonary vascular remodeling in MCT-PAH (27, 44, 48). Depending on the context, TGF-β1 may mediate pro- or anti-inflammatory responses, and its role in the development of PAH may be related to this interaction with inflammatory pathways. Human and animal models of PAH demonstrate abnormal levels of several inflammatory mediators, including IL-1 and IL-6 (4, 8, 15, 17). IL-6 appears to play a key role, since homozygous IL-6-null mice do not develop raised pulmonary artery pressures when challenged with hypoxia (40). Also, administration of dexamethasone to MCT-PAH rats reduces aberrant IL-6 release and prevents the development of vascular remodeling (2). Moreover, transgenic mice overexpressing a dominant-negative Bmpr2 exhibit increased IL-6 release and pulmonary hypertension (15).

We initially hypothesized that the loss of TGF-β1-mediated growth repression in HPAH PASMCs would result from disrupted Smad signaling. However, activation of the canonical TGF-β Smad2/3 signaling pathway was unaffected in HPAH PASMCs. Instead, comprehensive gene expression profiling of the TGF-β1 response in HPAH PASMCs with defined BMPR2 mutations and controls, coupled with gene set enrichment analysis (GSEA), identified an increased frequency of gene sets associated with inflammation in HPAH PASMCs. We confirmed enhanced NF-κB activation and expression of the proinflammatory cytokines IL-6 and IL-8 in HPAH PASMCs. Neutralization of these cytokines restored the antiproliferative effects of TGF-β1. Our findings suggest that BMPR-II dysfunction leads to enhanced basal and TGF-β1-stimulated secretion of proinflammatory cytokines, which antagonizes the antiproliferative effects of TGF-β1. This mechanism is likely to contribute to the abnormal accumulation of PASMCs that characterizes the vascular remodeling in PAH and provides a rationale for testing anti-interleukin therapies for the treatment of PAH.

METHODS

Isolation and culture of PASMCs.

Explant-derived PASMCs were obtained from proximal segments of human pulmonary artery and from peripheral pulmonary arteries (<2 mm diameter) obtained from patients undergoing lung or heart-lung transplantation for HPAH (n = 4). All HPAH isolates harbored disease-associated mutations (C347R, C347Y, N903S, and W9X) in BMPR-II. Control samples were obtained from unused donors for transplantation (n = 5). The Papworth Hospital Ethical Review Committee approved the study, and subjects gave informed written consent.

Segments of lobar pulmonary artery were cut to expose the luminal surface. The endothelium was removed by gentle scraping with a scalpel blade, and the media was peeled away from the underlying adventitial layer. The medial explants were cut into ∼4- to 9-mm² sections, plated into T25 flasks, and allowed to adhere for 2 h.

For peripheral explants, the lung parenchyma was dissected away from a pulmonary arteriole, following the arteriolar tree, to isolate 0.5- to 2-mm-diameter vessels. These were dissected out and cut into small fragments, which were plated in T25 flasks and left to adhere for 2 h. A section of the pulmonary arteriole was collected, fixed in formalin, and embedded in paraffin, and sections were analyzed to ensure that the vessel was of pulmonary origin. Cells were used between passages 4 and 8.

Once adhered, the explants were incubated in DMEM supplemented with 20% FBS and 1% antibiotic-antimycotic (Invitrogen) until cells had grown out and were forming confluent monolayers. PASMCs were trypsinized, and subsequent passages were propagated in DMEM supplemented with 10% heat-inactivated FBS and 1% antibiotic-antimycotic (DMEM-10% FBS) and maintained at 37°C in 95% air-5% CO₂. PASMCs were used between passages 4 and 8. The smooth muscle phenotype of isolated cells was confirmed by positive immunofluorescence with antibodies (Sigma) to anti-smooth muscle α-actin (IA4) and anti-smooth muscle-specific myosin (hsm-v).

Mouse PASMCs were isolated from Bmpr2^+/− mice and their wild-type (WT) littermate controls by explant culture, as previously described (28). Mouse PASMCs were maintained in culture in DMEM-10% FBS and used between passages 2 and 4.

Growth studies.

Human and mouse PASMCs were plated at a density of 15,000 cells per well in 24-well plates, maintained in DMEM-10% FBS for 24 h, and then serum-starved for a further 24 h. The cells were incubated in DMEM-10% FBS with or without TGF-β1 (10 ng/ml; R & D Systems, Abingdon, UK) and then trypsinized and counted using trypan blue (Sigma) exclusion on days 0, 2, 4, and 6 (n = 4 wells per treatment). TGF-β1 was refreshed every 48 h. For studies involving transfer of conditioned media, cells were grown to confluence and made quiescent as described above. The media were exchanged every 48 h for fresh DMEM-0.1% FBS with or without TGF-β1 (10 ng/ml). Media from control (n = 3) or HPAH (n = 3) PASMCs were pooled, and the conditioned media were incubated with fresh control or HPAH PASMCs seeded in 24-well plates. The number of viable cells was determined 6 days later. Each isolate was studied at least twice under each condition, and the mean values were taken from all studies conducted with any one isolate.

For neutralizing antibody studies, neutralizing antibodies (R&D Systems) to human IL-6 and human IL-8 or an IgG control were used at the indicated concentrations. PASMCs from control and HPAH patients were treated as described above in the presence and absence of TGF-β1 (10 ng/ml) and/or neutralizing antibodies. The effects on proliferation were assessed 6 days later, as described above.

BMPR-II small interfering RNA.

Human PASMCs were seeded in 24-well plates (15,000 cells per well) for growth studies, 6-well plates (60,000 cells per well) for RNA studies, and 6-cm dishes (180,000 cells per dish) for protein extraction. Cells were grown for 2 days in DMEM-10% FBS and then preincubated in OptiMEM I for 3 h. PASMCs were transfected with 10 nM small interfering RNA [siRNA; Dharmacon BMPR-II siGenome Smartpool (siBMPR-II) or Dharmacon siGLO (siGLO)] in complex with siFECTamine (ICVEC, London UK; 4 μl per well for 6-well plate or 8.75 μl per well for 6-cm dish). Cells were incubated with the complexes for 4 h at 37°C; then the medium was changed to DMEM-10% FBS for a further 24 h. Prior to treatment, cells were serum-restricted in DMEM-0.1% FBS for 24 h. The cells were then treated in DMEM-10% FBS with or without TGF-β1 (10 ng/ml) for growth studies or DMEM-0.1% FBS for the RNA and protein extraction studies. To confirm siRNA knockdown, cells were transfected in parallel to the growth curve, and protein extraction and specific reduction of the relevant RNA were confirmed using real-time PCR. The efficiency of BMPR-II siRNA targeting was confirmed by real-time PCR, and expression levels were normalized to β-actin using the cycle threshold (ΔΔC_T) method (26). BMPR-II protein reduction was confirmed by Western blotting.

Reporter gene assays and constructs.

Control or HPAH PASMCs were plated in 12-well plates at a density of 35,000 cells per well in 10% FBS-DMEM and grown for 48 h and then incubated in OptiMEM I for 3 h. Cells were transfected with the Smad1/5-responsive reporter BMP response element (BRE)-luciferase or the Smad3/4-responsive luciferase reporter promoter construct CAGA₁₂-luciferase. Cells were transfected with 4 μg of plasmid in complex with Lipofectamine 2000 (Invitrogen) prepared according to the manufacturer's protocol. Cells were incubated for 4 h with the complexes; then the medium was changed to 10% FBS overnight. For 3 h prior to treatment, cells were serum-deprived in 0.1% FBS and then treated for 24 h in 10% FBS with or without recombinant TGF-β1 or BMP4 (R & D Systems). Luciferase activity in the cells was assessed using a luciferase reporter assay kit (Roche).

Gene array analysis.

To evaluate the differences in responses of HPAH and control PASMCs to TGF-β1, gene array analysis was performed. Briefly, proximal PASMCs were grown to confluence. Subsequently, cells were maintained in DMEM-0.1% FBS for 48 h before the medium was exchanged for fresh DMEM-0.1% FBS with or without TGF-β1 (2 ng TGF-β1/ml) for 4 h. At the end of the treatment period, total RNA was harvested (RNeasy, Qiagen) and quantified, and integrity was verified by denaturing gel electrophoresis. Equal amounts of identically treated RNA were reverse-transcribed (Invitrogen) into cDNA, which was then in vitro-transcribed into biotinylated cRNA. The target cRNA was fragmented and hybridized to the Affymetrix human U133A array, following the Affymetrix (Santa Clara, CA) protocol. Hybridization of cRNA to Affymetrix human U133A chips, signal amplification, and data collection were performed using an Affymetrix fluidics station and chip reader. Chip files were scaled to an average intensity of 100 per gene and analyzed using Affymetrix version 5.0 (MAS5) comparison analysis software. Experiments were performed twice, and genes induced more than threefold (Table 1) are an average of these independent studies. Briefly, transcripts were defined as upregulated by TGF-β1 only when identified as “present” by the Affymetrix detection algorithm and “significantly increased” as determined by the Affymetrix change algorithm. Changes were defined as significant when P < 0.05. A ≥1.5-fold change between treated and untreated samples was required to identify a transcript as being altered. These criteria had to be met in both sets of experiments.

Table 1.

GSEA list of gene sets differentially regulated by TGF-β1 in HPAH and control PASMCs

HPAH PASMCs (0 h vs. 4 h TGF-β1)	Q Value, log10	Gene Sets Over- or Underrepresented
Protein biosynthesis	−16.4	↓
Cell structure and motility	12.5	↑
Cell motility	11.6	↑
Electron transport	−11.6	↓
Oxidative phosphorylation	−10.8	↓
Human osteoarthritis-interleukin signaling pathway	9.2	↑
Osteoarthritis-interleukin signaling pathway	8.3	↑
Cytoskeletal regulation by Rho GTPase	7.7	↑
Rheumatoid arthritis-interleukin signaling pathway	7.6	↑
Integrin outside-in signaling	6.7	↑
Propanoate metabolism	−5.1	↓
DNA repair	−4.8	↓
Human rheumatoid arthritis-interleukin signaling pathway	4.8	↑
MAPK signaling pathway	4.5	↑
Ubiquinone metabolism	−4.5	↓
Muckle Wells-responsive genes	4.3	↑
Human rheumatoid arthritis-MAPK-mediated pathway	4.1	↑

Pathways in italics were altered in control and heritable pulmonary arterial hypertension (HPAH) gene sets.

PASMCs, pulmonary artery smooth muscle cells; GSEA, gene set enrichment analysis; TGF-β1, transforming growth factor-β1.

GSEA pathway analysis.

In addition to the single gene analyses, we used the GSEA algorithm, which is a microarray data analysis method that uses predefined gene sets to identify significant biological changes in microarray data sets (43). GSEA is especially useful when the gene expression changes in a given microarray data set are relatively small. To define the gene sets, an in-house implementation of GSEA was performed using the gene chip microarray data, as previously described (30, 42). As input, GSEA requires microarray data from two conditions (e.g., unstimulated vs. TGF-β1-stimulated HPAH PASMCs). Briefly, genes significantly (1.5-fold, P < 0.05) induced by TGF-β1, with expression of average intensity >100 on the gene chips, were used for comparisons. Differential expression for each probe set was estimated by its expression ratio between pairs of conditions (presence or absence of TGF-β1). Gene sets differentially modulated were identified using Wilcoxon's rank sum test, and false discovery rate was set to gene sets with Q > 0.001 induced by TGF-β1 in HPAH PASMCs.

Real-time PCR analysis.

To investigate the temporal expression of genes, PASMCs from HPAH patients (n = 4) and control (n = 5) PASMCs were grown to confluence and serum-starved for 18 h and then stimulated with TGF-β1 (1 ng/ml) for 1 and 4 h. RNA was isolated using RNeasy mini-columns (Qiagen). DNase-digested total RNA (1 μg) was reverse-transcribed using a high-capacity cDNA reverse transcription kit (Applied Biosystems). Real-time PCRs (25 μl/reaction) were set up using 75 ng of cDNA with SYBR Green Jumpstart Taq ReadyMix (Sigma) containing the relevant sense and antisense primers at 200 nM each and 10 nM fluorescein (Invitrogen) and amplified on an iCycler (Bio-Rad Laboratories). Samples were analyzed using Qiagen Quantitect primers for IL-6, IL-8, plasminogen activator inhibitor 1 (PAI-1), β₂-microglobulin, and hypoxanthine phosphoribosyltransferase or custom primers for β-actin [5′-GCACCACACCTTCTACAATGA-3′ (sense) and 5′-GTCATCTTCTCGCGGTTGGC-3′(antisense)] or BMPR-II [5′-CAAATCTGTGAGCCCAACAGTCAA-3′ (sense) and 5′-GAGGAAGAATAATCTGGATAAGGACCAAT-3′ (antisense)]. The IL-6 and IL-8 expression levels were normalized to β-actin, hypoxanthine phosphoribosyltransferase, and β₂-microglobulin using the GeNorm normalization program (46). Real-time PCR was performed to confirm siRNA targeting of BMPR-II, and expression levels were normalized to β-actin using the ΔΔC_T method (26).

Inhibition of NF-κB activity.

Confluent PASMCs from HPAH patients (n = 2) and healthy controls (n = 2) in six-well plates were serum-starved in DMEM-0.1% FBS for 24 h. To ascertain the role of NF-κB on TGF-β1 induction of IL-6, cells were incubated with the IκB kinase inhibitor BMS-345541 (1 μmol/l; Calbiochem) or the peptide inhibitor of NF-κB nuclear translocation SN-50 (10 μg/ml; Calbiochem) for 45 min prior to TGF-β1 addition (2 ng/ml). Conditioned media were collected, and IL-6 levels were determined by ELISA using a commercial dual-antibody kit (R & D Systems), according to the manufacturer's instructions.

Western blot analysis.

Control and mutant human PASMCs were plated in 10-cm cell culture dishes and incubated in DMEM-10% FBS until 80% confluent. After serum deprivation in DMEM-0.1% FBS for 24 h, cells were treated with DMEM-0.1% FBS with or without TGF-β1 (10 ng/ml) or BMP4 (50 ng/ml) for 1 h. Cells were snap-frozen on an ethanol-dry ice bath. Cells were then lysed with 150 μl of lysis buffer containing 125 mM Tris (pH 7.4), 2% SDS, 10% glycerol, and an EDTA-free protease inhibitor cocktail (Roche). Samples were sonicated for two periods of 5 s and stored at −20°C. For Western blotting, samples were subjected to SDS-PAGE (12%) and transferred by semidry blotting to nitrocellulose membranes and then, in later experiments, polyvinylidene difluoride membranes (Bio-Rad). Protein was loaded at 40–60 μg/lane. Lysates were immunoblotted with COOH-terminal phosphorylated Smad1/5 rabbit monoclonal, COOH-terminal phosphorylated Smad2 rabbit polyclonal, phosphorylated NF-κB p65 (Ser⁵³⁶) rabbit polyclonal (Cell Signaling Technology), COOH-terminal phosphorylated Smad3 rabbit polyclonal (R & D Systems), or BMPR-II mouse monoclonal (BD Transduction Laboratories) antibody. All blots were reprobed with anti-human β-actin mouse monoclonal antibody (Sigma).

Statistical analysis.

Values are means ± SE. Data were analyzed with GraphPad Prism (version 3.0). Comparisons were made by Student's t-test or one-way ANOVA, with Tukey's post hoc honestly significant difference test when appropriate. P ≤ 0.05 indicated statistical significance.

RESULTS

BMPR-II deficiency renders PASMCs insensitive to growth inhibition by TGF-β1.

We previously reported that TGF-β1 inhibits serum-induced proliferation of proximal PASMCs from healthy individuals, whereas cells from idiopathic PAH patients proliferate to TGF-β1 (33). Vascular remodeling and occlusion of the vasculature, synonymous with the development of HPAH, occur within the small- to medium-diameter arterioles (6, 18). As BMPR-II deficiency predisposes patients to HPAH, we examined the potential link between BMPR-II deficiency and failed growth inhibition by TGF-β1. Consistent with our previous findings, proliferation of control distal PASMCs was significantly repressed (Fig. 1A) by exogenous TGF-β1 (10 ng/ml), whereas HPAH cells with defined BMPR-II mutations were refractory to the antiproliferative effects of TGF-β1 (Fig. 1B). To establish if reduced BMPR-II expression was directly associated with this resistance to TGF-β1, we compared this response in WT and Bmpr2^+/− PASMCs. Again, the proliferation of WT mouse PASMCs was significantly repressed (P < 0.05) by exogenous TGF-β1 (Fig. 1C), whereas Bmpr2^+/− mouse PASMCs were insensitive to growth inhibition by TGF-β1 (Fig. 1D). Furthermore, siRNA targeting of BMPR-II (siB) significantly (P < 0.05) reversed the antiproliferative effects of TGF-β1 (Fig. 1E), which was intact in PASMCs treated with transfection reagent alone (siF) or transfected with a nontargeting siRNA (siG). Using real-time PCR, BMPR-II mRNA was reduced by >70% of endogenous gene levels, and protein loss was confirmed by Western blotting (Fig. 1F).

Fig. 1.

Reduced bone morphogenetic protein (BMP) type II receptor (BMPR-II) in pulmonary artery smooth muscle cells (PASMCs) leads to loss of the growth-inhibitory effect of transforming growth factor (TGF)-β1. Effect of TGF-β1 (10 ng/ml) on PASMC proliferation in 0.1% or 10% FBS was examined by cell counting on days 0, 2, 4, and 6 (n = 4 wells per treatment). Treatments were replenished every 48 h. A: control human PASMCs (n = 3). **P < 0.01. ***P < 0.001. B: PASMCs from patients with heritable pulmonary arterial hypertension (HPAH) harboring Bmpr2 mutations (n = 3). *P < 0.05. C: pooled PASMC isolates from wild-type (WT) mice (n = 2 isolates). **P < 0.01. ***P < 0.001. D: pooled PASMC isolates from Bmpr2^+/− mice (n = 2 isolates). Values are means ± SE of 3 independent experiments. *P < 0.05 vs. respective control (Student's t-test). E: TGF-β1 (10 ng/ml) inhibited proliferation of control human PASMCs transfected with vehicle (siF) or control small interfering RNA (siRNA, siG), but not cells transfected with BMPR-II siRNA (siB). *P < 0.05. F: BMPR-II protein was reduced by BMPR-II siRNA compared with cells transfected with vehicle or control siRNA.

BMPR-II deficiency does not alter TGF-β1-induced Smad signaling or transcriptional responses.

TGF-β superfamily ligands activate the canonical Smad signaling pathways, so we examined whether reduced BMPR-II levels alter the BMP-activated Smads, Smad1 and Smad5, or the TGF-β1-activated Smads, Smad2 and Smad3. BMP4-mediated COOH-terminal phosphorylation of Smad1/5 was reduced in HPAH PASMCs compared with controls (Fig. 2A). Consistent with this, the intensity of the BMP4-induced phosphorylated Smad1 band detected by the phosphorylated Smad1/3 antibody was also diminished. In contrast, similar activation of Smad2 or Smad3 was seen in response to TGF-β1 in HPAH and control PASMCs (Fig. 2A). Consistent with our findings in HPAH PASMCs, BMPR-II siRNA transfection significantly reduced Smad1/5 phosphorylation by BMP4, whereas the TGF-β1-mediated phosphorylation of Smad2 and Smad3 was not affected (Fig. 2B). We also examined the impact of reduced BMPR-II expression using Smad-responsive luciferase reporter constructs. BMP4 significantly (P < 0.05) and selectively induced the BRE-luciferase response, and TGF-β1 selectively induced CAGA₁₂-luciferase activity in control human PASMCs (Fig. 2C). Consistent with the Smad immunoblot data, HPAH cells exhibited reduced BRE-luciferase activity compared with control cells, whereas TGF-β1-induced CAGA₁₂-luciferase activity (P < 0.05) was intact in HPAH PASMCs (Fig. 2D). In addition, the induction of PAI-1, a transcriptional target of TGF-β1, did not differ between control and HPAH PASMCs (Fig. 2E). We also questioned whether the temporal activation of Smad2 or Smad3 was altered in the setting of reduced BMPR-II expression. When compared directly, the rate of decay of the phosphorylated Smad2 and phosphorylated Smad3 signals did not differ between control and HPAH PASMCs (Fig. 2F).

Fig. 2.

Activation status of Smads in PASMCs. A and B: activation of the canonical Smad signaling pathways in response to BMP4 or TGF-β1 in control (C) or HPAH (M) PASMCs with a defined Bmpr2 (W9X) mutation (A) or control PASMCs transfected with BMPR-II siRNA (siB) or a nontargeting siRNA (siG) using siFectamine (siF) (B). Serum-starved cells were treated with TGF-β1 (10 ng/ml), BMP4 (50 ng/ml), or media alone for 1 h. Cell lysates were immunoblotted for phosphorylated Smad1/5 (pSmad1/5), Smad2 (pSmad2), and Smad3 (pSmad3). C and D: Smad-dependent transcriptional responses to BMP4 and TGF-β1 in control PASMCs (C; n = 3) and HPAH PASMCs (D; n = 3) transfected with the Smad3-responsive CAGA₁₂-luciferase (CAGA₁₂-Luc) reporter or the Smad1/5/8-responsive BMP response element (BRE)-luciferase (BRE-Luc) reporter. *P < 0.05. E: transcriptional induction of plasminogen activator inhibitor (PAI)-1 by TGF-β1 in control (n = 3) and HPAH (n = 3) PASMCs treated with TGF-β1 (1 ng/ml) for 4 h. RNA was reverse-transcribed and analyzed for PAI-1 expression using real-time PCR. Gene expression changes were normalized to 3 housekeeping genes (β₂-microglobulin, hypoxanthine phosphoribosyltransferase, and β-actin). F: longevity of the TGF-β1-driven Smad response in control (C) and HPAH (M) PASMCs. Cells were incubated with TGF-β1 for 1 h; then the medium was removed, and DMEM-0.1% FBS containing the ALK5 inhibitor SD-208 (2 μM) was added. Cells were lysed at the times indicated and immunoblotted for phosphorylated Smad2 and phosphorylated Smad3.

A secreted factor released by HPAH PASMCs antagonizes TGF-β1-mediated growth suppression.

Having demonstrated that abnormal TGF-β responses in HPAH cells are unlikely to be due to alterations in canonical Smad signaling, we sought to determine whether the loss of the antiproliferative effects of TGF-β1 in HPAH PASMCs was due to a secreted mediator. To address this, we assessed the proliferative potential of media from HPAH or control PASMCs grown in the presence or absence of TGF-β1. Conditioned media from control PASMCs exposed to TGF-β1 significantly inhibited (P < 0.05) control PASMC proliferation compared with media conditioned without exogenous TGF-β1 (Fig. 3A). The growth of control PASMCs in media from HPAH PASMCs was identical to that in media from control PASMCs. Intriguingly, the proliferation of control PASMCs was not inhibited by media from HPAH PASMCs conditioned with TGF-β1. Furthermore, HPAH PASMCs grown in control or HPAH PASMC-conditioned media were refractory to the antiproliferative effects of TGF-β1 (Fig. 3A). Thus the loss of TGF-β1 growth-inhibitory effects appeared to be mediated by a soluble factor(s) secreted in response to TGF-β1 in HPAH PASMCs.

Fig. 3.

Conditioned medium from HPAH PASMCs inhibits TGF-β1 antiproliferative effects on control PASMCs. Conditioned media were collected from control (C-CM) and HPAH (H-CM) PASMCs grown in DMEM-10% FBS in the presence or absence of TGF-β1 (10 ng/ml) for 48 h. A: conditioned media were transferred to fresh control or HPAH PASMCs and replaced every 48 h, and cells were counted on day 6. Values are means ± SE (n = 4 wells) from a representative experiment for a control and HPAH PASMC line treated in parallel. *P < 0.05 (Student's t-test). B: conditioned media from control and HPAH PASMCs treated as described in A were assayed for induction of CAGA₁₂-luciferase activity, confirming that TGF-β1 activity was comparable. ***P < 0.001. C: TGF-β1 induced CAGA₁₂-luciferase activity in a concentration-dependent manner in mink lung stable transfectants. ***P < 0.001.

It was possible that HPAH PASMCs produce a factor that may be neutralizing or inactivating the exogenous TGF-β1. Therefore, active TGF-β1 was assayed in conditioned media from control or HPAH PASMCs incubated with or without exogenous TGF-β1 using mink lung cells stably expressing CAGA₁₂-luciferase (kindly provided by D. B. Rifkin) (45). Media from untreated control and HPAH PASMCs stimulated a low response, whereas similar levels of luciferase activity were stimulated by conditioned media from TGF-β1-stimulated cells, confirming that the TGF-β1 was still active.

TGF-β1 induced differential gene expression in HPAH PASMCs.

On the basis of these data, we surmised that TGF-β1 was activating different transcriptional responses in control and HPAH PASMCs. In cells treated with TGF-β1 for 4 h, the expression changes of genes induced >3-fold revealed that the majority (>70%) of transcripts were similarly regulated by TGF-β1 in control and HPAH PASMCs (Table 1). In contrast, when we compared the gene transcripts significantly (by >1.5-fold) changed by TGF-β1, marked differences were revealed between control and HPAH PASMCs. For example, 513 genes were significantly altered >1.5-fold in HPAH PASMCs (P < 0.05, 372 increased and 141 decreased) compared with 339 genes in HPAH PASMCs (P < 0.05, 237 increased and 102 decreased).

To assess the differences in transcriptional responses between control and HPAH PASMCs, GSEA was performed using the gene chip microarray data, as previously described (30). Briefly, we compared genes significantly (1.5-fold, P < 0.05) induced in HPAH and control PASMCs in response to 4 h of TGF-β1 stimulation. Gene sets as defined on the basis of pathway or biological processes in HPAH PASMCs are shown in Table 2. Gene sets associated with inflammatory pathways, including the interleukin signaling pathways, were overrepresented in response to TGF-β1, whereas gene sets associated with “oxidative phosphorylation” were underrepresented.

Table 2.

Genes induced or repressed more than threefold in HPAH and control PASMCs

HPAH PASMCs			Control PASMCs
Gene ID	Description	Change	Gene ID	Description	Change
HBEGF	Heparin-binding EGF-like growth factor	51.71	HBEGF	Heparin-binding EGF-like growth factor	25.69
PMEPA1	Transmembrane, prostate androgen-induced RNA	29.32	PMEPA1	Transmembrane, prostate androgen-induced RNA	18.74
ANGPTL4	Angiopoietin-like 4	14.29	ELN	Elastin	18.22
HBEGF	Heparin-binding EGF-like growth factor	12.00	FOSB	FBJ murine osteosarcoma viral oncogene homolog B	14.32
IL11	Interleukin 11	11.06	IL11	Interleukin 11	12.81
FOSB	FBJ murine osteosarcoma viral oncogene homolog B	10.58	CCDC99	Coiled-coil domain containing 99	11.06
NR4A3	Nuclear receptor subfamily 4, group A, member 3	10.34	COMP	Cartilage oligomeric matrix protein	9.98
PGDB5	PiggyBac transposable element-derived 5	9.91	NEDD9	Neural precursor cell-expressed, developmentally downregulated 9	8.32
CCDC99	Coiled-coil domain-containing 99	7.78	FGF18	Fibroblast growth factor 18	8.16
SLC22A3	Solute carrier family 22	7.41	PCDH9	Protocadherin 9	7.26
PTHLH	Parathyroid hormone-like hormone	7.29	LIF	Leukemia inhibitory factor	7.20
NEDD9	Neural precursor cell-expressed, developmentally downregulated 9	7.23	KIAA1644	KIAA 1644 protein	6.70
MFAP3L	Microfibrillar-associated protein 3-like	7.06	HBEGF	Heparin-binding EGF-like growth factor	6.26
F2RL1	Coagulation factor II (thrombin) receptor-like 1	5.77	PTHLH	Parathyroid hormone-like hormone	6.25
EIF1AY	Eukaryotic translation initiation factor 1A, Y-linked	5.69	PRR5L	Proline-rich 5-like	6.03
TUBB2A	Tubulin, β2A	5.38	SPHK1	Sphingosine kinase 1	6.01
KIAA1644	KIAA1644 protein	5.19	GADD45B	Growth arrest and DNA-damage-inducible, β	5.89
GADD45B	Growth arrest and DNA damage-inducible, β	5.18	PTHLH	Parathyroid hormone-like hormone	5.84
UPP1	Uridine phosphorylase 1	5.05	PDGFA	Platelet-derived growth factor-α polypeptide	5.66
FGF2	Fibroblast growth factor 2 (basic)	5.05	HAS2	Hyaluronan synthase 2	5.52
TNFRSF12	TNF receptor superfamily, member 12A	4.90	GADD45B	Growth arrest and DNA damage-inducible, β	5.32
PTHLH	Parathyroid hormone-like hormone	4.86	NOX4	NADPH oxidase 4	5.02
PDGFA	Platelet-derived growth factor-α polypeptide	4.84	ERG	v-ets	4.95
GADD45B	Growth arrest and DNA damage-inducible, β	4.74	LIMK2	LIM domain kinase 2	4.57
HOMER1	Homer homolog 1 (Drosophila)	4.67	NFATC1	Nuclear factor of activated T cells, calcineurin-dependent 1	4.44
TSPAN13	Tetraspanin 13	4.53	XYLT1	Xylosyltransferase I	4.41
SPHK1	Sphingosine kinase 1	4.40	GADD45B	Growth arrest and DNA damage-inducible, β	4.08
ATP13A3	ATPase type 13A3	4.24	TNFRSF12	TNF receptor superfamily, member 12A	3.94
FGF2	Fibroblast growth factor 2 (basic)	4.15	PTHLH	Parathyroid hormone-like hormone	3.93
LMCD1	LIM and cysteine-rich domains 1	4.11	KLF10	Kruppel-like factor 10	3.86
F2RL1	Coagulation factor II (thrombin) receptor-like 1	4.08	SMAD7	SMAD family member 7	3.66
NUPL1	Nucleoporin-like 1	4.02	PDLIM4	PDZ and LIM domain 4	3.60
CLCF1	Cardiotrophin-like cytokine factor 1	4.00	FGF2	Fibroblast growth factor 2 (basic)	3.57
NOX4	NADPH oxidase 4	3.97	PDGFA	Platelet-derived growth factor-α polypeptide	3.56
ENC1	Ectodermal-neural cortex (with BTB-like domain)	3.95	FGF2	Fibroblast growth factor 2 (basic)	3.56
NR4A3	Nuclear receptor subfamily 4, group A, member 3	3.83	ZNF124	Zinc finger protein 124	3.56
TNS1	Tensin 1	3.78	PRMT2	Protein arginine methyltransferase 2	3.50
SERPINE1	Serpin peptidase inhibitor, clade E (nexin, plasminogen activator inhibitor type 1), member 1	3.76	MYCN	v-myc myelocytomatosis viral-related oncogene	3.31
GADD45B	Growth arrest and DNA damage-inducible, β	3.76	KIR	Killer cell immunoglobulin-like receptor,	3.23
PDLIM4	PDZ and LIM domain 4	3.76	MAP2K3	MAPK kinase 3	3.13
LIMK2	LIM domain kinase 2	3.75	PNP	Nucleoside phosphorylase	3.11
ATP13A3	ATPase type 13 A3	3.70	IL11	Interleukin 11	3.09
CTGF	Connective tissue growth factor	3.63	UAP1	UDP-N-acteylglucosamine pyrophosphorylase 1	3.04
PDGFA	Platelet-derived growth factor-α polypeptide	3.58	DACT1	Dapper, antagonist of β-catenin, homolog 1	3.04
RHOB	ras homolog gene family, member B	3.55	THBD1	Thrombomodulin	3.01
PNP	Nucleoside phosphorylase	3.52
NFATC1	Nuclear factor of activated T cells, calcineurin-dependent 1	3.46
AMIGO2	Adhesion molecule with Ig-like domain 2	3.44
PCDH9	Protocadherin 9	3.43
HMGA2	High-mobility group AT-hook 2	3.30
PTHLH	Parathyroid hormone-like hormone	3.28
IGFBP7	Insulin-like growth factor binding protein 7	3.21
CCNF	Cyclin F	3.20
STK38L	Ser/Thr kinase 38-like	3.17
CBFB	Core-binding factor, β-subunit	3.16
BHLHE40	Basic helix-loop-helix domain-containing, class B, 2	3.16
KLF10	Kruppel-like factor 10	3.16
PSG1	Pregnancy-specific-β1 glycoprotein 1	3.16
NEDD9	Neural precursor cell-expressed, developmentally downregulated 9	3.10
SMAD7	SMAD family member 7	3.08
VEGFA	Vascular endothelial growth factor A	3.05
BMP6	Bone morphogenetic protein 6	3.03
PLAUR	Plasminogen activator, urokinase receptor	3.01
INHBE	Inhibin, βE	−3.20	MAGIX	MAGI family member, X-linked	−3.07
BFSP1	Beaded filament structural protein 1, filensin	−3.21	ZMYM5	Zinc finger, MYM-type 5	−3.08
C13orf8	Chromosome 13 open reading frame 18	−3.24	LOC729799	SEC14-like 1 pseudogene	−3.14
PHKA2	Phosphorylase kinase, α2 (liver)	−3.81	CXCL1	Chemokine (C-X-C motif) ligand 1	−3.19
POLD1	Polymerase, δ1, catalytic subunit 125 kDa	−4.09	—	Clone 24629 mRNA sequence	−3.22
RGS12	Regulator of G protein signaling 12	−4.36	KLHL23	Kelch-like 23 (Drosophila)	−3.33
GAS1	Growth arrest-specific 1	−4.36	NEK1	NIMA-related kinase 1	−4.07
C5orf4	Chromosome 5 open reading frame 4	−4.36		Hypothetical protein FLJ23556	−4.56
SRGAP2	SLIT-ROBO Rho GTPase-activating protein 2	−4.47	RPL7	Ribosomal protein L7	−4.61
CDS1	CDP-diacylglycerol synthase 1	−4.48	CEBPD	CCAAT/enhancer-binding protein (C/EBP), δ	−4.61
ADM	Adrenomedullin	−5.03	PLK1S1	Polo-like kinase 1 substrate 1	−5.48
ALDH4A1	Aldehyde dehydrogenase 4 family, member A1	−5.34	GAS1	Growth arrest-specific 1	−5.75
GAS1	Growth arrest-specific 1	−6.63	MED13L	Mediator complex subunit 13-like	−6.12
C10orf10	Chromosome 10 open reading frame 10	−6.95	CASC1	Cancer susceptibility candidate 1	−6.44
BMP4	Bone morphogenetic protein 4	−10.99	BTF3P11	Basic transcription factor 3, like 1	−6.49
			STEAP4	STEAP family member 4	−13.09

Cells were incubated for 4 h in DMEM-0.1% FBS with or without TGF-β1 (2 ng/ml), and extracted RNA was subjected to microarray analysis. Values (means from 2 experimental repeats) are fold changes of expression in TGF-β1-treated cells compared with their untreated controls. EGF, epidermal growth factor. KIR probe detects multiple KIR isoforms.

TGF-β1 enhances interleukin gene expression in HPAH PASMCs.

As TGF-β1-stimulated gene sets associated with the interleukin pathway were overrepresented in HPAH PASMCs compared with control PASMCs, we questioned whether the soluble factor might be an interleukin. Therefore, we first profiled the temporal induction of IL-6 and IL-8 in HPAH and control PASMCs in response to TGF-β1. TGF-β1 promoted a greater transcriptional induction of IL-6 at 4 h in HPAH PASMCs than control PASMCs, but this did not achieve significance (Fig. 4A). In contrast, TGF-β1 significantly (P < 0.05) induced a greater transcriptional induction of IL-8 in HPAH PASMCs than control PASMCs at 4 h (Fig. 4B).

Fig. 4.

Enhanced interleukin expression in proximal HPAH PASMCs. Confluent, serum-starved HPAH (n = 4) and control (n = 5) PASMCs were treated for 1 and 4 h in the presence or absence of TGF-β1 (2 ng/ml). Total RNA was isolated from each sample and reverse-transcribed, and cDNA was subjected to real-time PCR analysis for IL-6 (A) and IL-8 (B), and expression was normalized to 3 housekeeping genes (β₂-microglobulin, hypoxanthine phosphoribosyltransferase, and β-actin). Data are expressed as fold induction by TGF-β1 compared with 0.1% control for each time point. Each data point represents a different PASMC isolate. *P < 0.05.

Elevated NF-κB activation in HPAH PASMC nuclear extracts.

Several pathways mediate transcriptional induction of interleukins, including members of the activator protein 1 (AP-1) and NF-κB transcription factor families (9, 10, 22, 38). We found a constitutively higher phosphorylation of the p65/RelA NF-κB signal in HPAH PASMCs than control cells (Fig. 5,A and B). To further define the importance of the differential activation of NF-κB/p65 in HPAH PASMCs, the impact of the NF-κB inhibitors SN-50 and BMS-345541 (3) on IL-6 secretion was determined (Fig. 5C). TGF-β1 induced a significant (P < 0.001) increase in secretion of IL-6 in PASMCs. Inhibition of NF-κB significantly (P < 0.05) reduced TGF-β1-induced IL-6 secretion in HPAH PASMCs, whereas an increase (P < 0.05) was observed in control PASMCs. The specificity of BMS-345541 was confirmed using a multimerized NF-κB reporter construct stimulated with TNF-α (data not shown). These data suggest that the role of NF-κB in control PASMCs may be to restrict IL-6 responses to TGF-β1, whereas inappropriate NF-κB activation in HPAH PASMCs contributes to the enhanced secretion of IL-6 by these cells.

Fig. 5.

NF-κB/p65 activity is dysregulated in HPAH PASMCs. A: confluent serum-starved control (n = 3) or HPAH (n = 3) PASMCs (N903S, C347Y, and W9X mutations) were lysed and immunoblotted for phosphorylated p65 (pp65). B: when analyzed by densitometry, basal phosphorylated p65 was significantly increased in HPAH compared with control human PASMCs. *P < 0.05. C: serum-starved control and HPAH PASMCs were incubated for 1 h in the presence or absence of BMS-345541 (BMS, 1 μmol/l) or SN-50 (10 μg/ml) and then stimulated with TGF-β1 (2 ng/ml). Media were collected 24 h later, and IL-6 levels were determined by ELISA in triplicate. Values are means ± SE. *P < 0.05 (Student's t-test). ***P < 0.001.

Inhibition of IL-6 and IL-8 restores the antiproliferative effects of TGF-β1 on HPAH PASMCs.

Several proproliferative growth factors, including IL-6, are induced by TGF-β1 in PASMCs (31). To examine if the loss of the antiproliferative effects of TGF-β1 on HPAH PASMCs is mediated by the induction of interleukins, we assessed the effects of neutralizing antibodies to IL-6 and IL-8 on proliferation. Neutralizing antibodies against IL-6, IL-8, or an IgG control antibody did not alter the antiproliferative effects of TGF-β1 on control PASMCs (Fig. 6A). In contrast, neutralization of IL-6 or IL-8, but not the IgG control, significantly (P < 0.05) restored TGF-β1 antiproliferative effects on PASMCs from HPAH patients (Fig. 6B). These data are consistent with the notion that the loss of TGF-β1 antiproliferative effects on HPAH PASMCs is mediated via the enhanced secretion of IL-6 and IL-8.

Fig. 6.

Inhibition of IL-6 or IL-8 restores antiproliferative effects of TGF-β1 on HPAH PASMCs. Serum-starved control (A; n = 3 lines) or HPAH (B; n = 3 lines) PASMCs were treated for 6 days in the presence (+) or absence (−) of TGF-β1 (10 ng/ml) with neutralizing anti-human IL-6 (IL-6 Ab), anti-human IL-8 (IL-8 Ab), or IgG control. Cell counts from a representative experiment for 1 control and 1 HPAH PASMC line are shown. Values are means ± SE (n = 4 wells per treatment). *P < 0.05 (Student's t-test).

DISCUSSION

We have identified a link between the loss of BMPR-II, inappropriate NF-κB signaling, and increased proinflammatory cytokine secretion by PASMCs in response to TGF-β1 that overrides the normal antiproliferative effects of TGF-β1 on PASMCs. Proinflammatory cytokines have previously been proposed as a “second hit” in the development of PAH. Therefore, we provide a mechanism by which BMPR-II mutations in HPAH cause susceptibility to an inflammatory second hit through the altered cellular response to TGF-β1 (Fig. 7).

Fig. 7.

Proposed mechanism by which enhanced IL-6 and IL-8 secretion by HPAH PASMCs may inhibit the antiproliferative response to TGF-β1. A: in control PASMCs, BMPs, signaling via a receptor complex containing BMPR-II, repress proliferation. TGFβR, TGF-β receptor. B: TGF-β1, acting through TGF receptors, also inhibits proliferation, and these effects combine to promote vessel wall homeostasis. TGF also induces IL-6 production, an effect negatively regulated by cellular NF-κB signaling. Loss of BMPR-II leads to altered NF-κB signaling in HPAH PASMCs. This leads to a permissive effect of dysregulated TGF-β1 signaling through NF-κB and enhanced IL-6 and IL-8 secretion. These inhibit the antiproliferative effect of TGF-β1, leading to PASMC proliferation and increased muscularization of the vessel wall. This supports further investigation of anti-IL-6/IL-8 as potential therapy for blockade of this pathological process.

We previously reported that proximal PASMCs from patients with idiopathic PAH of unknown BMPR-II status are insensitive to the antiproliferative effects of TGF-β1 (33). We hypothesized that resistance to the antiproliferative effects of TGF-β1 is a general feature of HPAH PASMCs as a direct consequence of reduced BMPR-II expression. We show that HPAH PASMCs, from the disease-relevant distal arterioles, exhibited a similar level of insensitivity to TGF-β1 treatment. As PASMCs from HPAH patients may differ from control cells because of disease pathology, we confirmed our observations in two models of BMPR-II deficiency without overt disease, Bmpr2^+/− mouse PASMCs and control human PASMCs transfected with BMPR-II siRNA. Our data confirm that the loss of TGF-β1 repressive effects on PASMC proliferation is directly associated with reduced BMPR-II.

We hypothesized that the insensitivity of HPAH PASMCs to growth repression may be through altered canonical TGF-β1 signaling. However, we did not observe changes in Smad2 or Smad3 phosphorylation or Smad3 transcriptional responses. A previous study of prostate cancer cells reported that Smad2 synergizes with other pathways to mediate the induction of IL-6 by TGF-β1 (38), but whether this mechanism is relevant to normal cells is unclear. We do not see any overt change in Smad2, but we cannot rule out a switch in Smad2 utilization as a contributing factor to the refractory response to TGF-β1 in HPAH cells.

We established that a secreted factor from HPAH PASMCs was rendering PASMCs insensitive to the growth-inhibitory effect of TGF-β1. It was possible that the cells may be releasing a factor that inactivated or neutralized the exogenous TGF-β1 in the media. A study reported that expression of the BMP ligand trap Gremlin is increased in the lungs of PAH patients, so it was possible that other ligand traps could be expressed by HPAH PASMCs (5). We observed no difference in TGF-β1 activity between conditioned media from control and HPAH PASMCs, suggesting that ligand neutralization was not the mechanism.

To identify pathways potentially linked to this phenomenon, we employed gene chip profiling and GSEA on transcripts induced by TGF-β1 in PASMCs with defined BMPR-II mutations. TGF-β1 induced several gene set changes common to control and HPAH PASMCs, including those associated with cell structure and motility. Interestingly, TGF-β1 induced gene sets associated with the Rho and MAPK signaling pathways in HPAH PASMCs, but not control PASMCs. Previous studies implicated Rho kinase (7, 14, 24, 37) and MAPK pathways (15, 32, 47) in the development of PAH. Indeed, Rho kinase is a potential therapeutic target for PAH, and animal studies have proven favorable (24, 36). However, our data highlighted a potential role for one or more soluble secreted factors, and the most striking relevant gene sets enhanced in HPAH PASMC were those associated with interleukins and inflammation.

We sought to confirm this overrepresentation of gene sets associated with interleukins in HPAH cells treated with TGF-β1 by comparing the responses of cells from controls and HPAH patients. On the basis of our previous report that circulating IL-6 and IL-8 levels are significantly raised in HPAH patients and correlate with reduced survival, we chose to analyze these cytokines (41). We confirm that HPAH PASMCs show enhanced transcription of IL-8 and IL-6 in response to TGF-β1 compared with control PASMCs. The regulation of these cytokines is dependent on the cell context and may differ between normal and disease states. In lung fibroblasts, TGF-β1 induces IL-6 via AP-1 (9), whereas in vascular smooth muscle cells, TGF-β1 represses cytokine-mediated IL-6 expression via a Smad3-dependent mechanism (11). In prostate cancer cells, the induction of IL-6 by TGF-β1 is mediated through synergistic interactions of the Smad2, p38-NF-κB, JNK, and Ras pathways (38). No functional Smad binding site has been identified in the IL-8 promoter region, implying that the TGF-β1 responses we observed are not focused on dysregulated Smad2/3 signaling. Similar to IL-6, NF-κB and AP-1 binding sites are present in the IL-8 promoter (9). We focused primarily on a potential role for inappropriate NF-κB signaling in the dysregulated interleukin responses observed in HPAH PASMCs. Consistent with this hypothesis, elevated basal NF-κB phosphorylated p65 was evident in HPAH PASMCs. Furthermore, pharmacological inhibition of NF-κB abrogated the heightened induction of IL-6 in HPAH PASMCs. Intriguingly, NF-κB blockade in control cells led to an enhancement of the IL-6 response to TGF-β1, suggesting that NF-κB may normally be a negative regulator of TGF-β1-mediated IL-6 signaling. Our data showing opposing effects of NF-κB blockade on TGF-β1-mediated IL-6 production by control and HPAH PASMCs suggest that NF-κB signaling is misdirected as a consequence of BMPR-II insufficiency. The regulation of NF-κB signaling is very complex, as components of this pathway can be phosphorylated on several serine residues to mediate selective responses (16). Furthermore, other modifications, such as acetylation of NF-κB subunits, also regulate their activity (16). Indeed, in epithelial cells, TGF-β1 can enhance NF-κB activation elicited by exposure to bacteria via p300 acetyltransferase in concert with a Smad3/4-dependent mechanism (19). Establishing the nature of the change in NF-κB response would be interesting, but as shown by our present results, targeting of the enhanced cytokine production provides a more accessible target for potential therapeutic intervention.

Significant evidence in the literature supports the role of inflammation in the initiation and development of HPAH (8, 15, 17, 20, 29). For example, HPAH patients have increased circulating levels of inflammatory cytokines, including monocyte chemoattractant protein-1, IL-1β, IL-6, IL-8, and TNF-α (17, 20, 20). We recently reported that circulating IL-6 and IL-8 levels are significantly raised in HPAH patients and correlate with reduced survival (41). Animal studies support the pathological role of inflammatory cytokines in the initiation and development of PAH. For example, IL-6 is elevated in several animal models of experimental PAH (2, 29) and in inducible targeted SM22-tet-Bmpr2^delx4+ mice (15). Additionally, recombinant IL-6 injection induces mild PAH in animals and augments hypoxia-driven PAH (13, 29). Moreover, Rho kinase is reported to mediate the induction of IL-6 in osteoblasts, so it would be interesting to examine if Rho kinase inhibitors reduce IL-6 in animal models of PAH (21). Consistent with a pathological role for proinflammatory cytokines, we found that neutralization of IL-6 or IL-8 restored the antiproliferative effects of TGF-β1 on HPAH PASMCs. Anti-IL-6 antibody treatment has previously been shown to restore the antiproliferative and proapoptotic effects of TGF-β1 in human prostate cancer cell lines (38). Furthermore, anti-IL-6 therapies have proven clinically safe and effective for treating rheumatoid arthritis and other disorders (34, 35). Administration of tocilizumab, a humanized anti-IL-6 monoclonal antibody, has improved the PAH symptoms in a patient with mixed connective tissue disease and severe PAH (12). Our data suggest that anti-IL-6 therapy may be a suitable intervention in PAH to target the effects of inflammation while also restoring the normal antiproliferative response to TGF-β1.

In summary, this study is the first to demonstrate a direct link between BMPR-II deficiency and the failure of TGF-β1 to inhibit the growth of HPAH PASMCs. We show that the enhanced IL expression profile in HPAH PASMCs corresponds to enhanced IL-6 and IL-8 secretion, the inhibition of which restores the normal antiproliferative effects of TGF-β1 to HPAH PASMCs. IL-8 is commonly known as a neutrophil chemoattractant and proangiogenic factor, whereas IL-6 is a pleiotropic cytokine with various biological activities, including promotion of PASMC proliferation. Excessive production of IL-6 and IL-8 has been implicated in the pathogenesis of several human conditions, including chronic inflammatory diseases, such as rheumatoid arthritis. Antibody-based therapies are currently being tested clinically; thus IL-6 and/or IL-8 represent targets for the treatment of the inflammation and vascular remodeling underpinning the pathology of HPAH.

GRANTS

This work was supported through British Heart Foundation Programme Grant RG/08/002/24718 (awarded to N. W. Morrell). R. J. Davies is supported by Wellcome Trust Clinical Research Training Fellowship GR077167MA.

DISCLOSURES

N. W. Morrell and P. D. Upton have received funding from Novartis in the form of a Novartis strategic external collaboration grant. A. M. Holmes, L. Barker, C. Walker, and D. C. Budd were employed by Novartis during the course of these studies.

AUTHOR CONTRIBUTIONS

R.J.D., A.M.H., C.W., D.C.B., P.D.U., and N.W.M. are responsible for conception and design of the research; R.J.D., A.M.H., J.D., L.L., X.Y., L.B., and P.D.U. performed the experiments; R.J.D., A.M.H., J.D., and P.D.U. analyzed the data; R.J.D., A.M.H., C.W., D.C.B., P.D.U., and N.W.M. interpreted the results of the experiments; R.J.D., A.M.H., and P.D.U. prepared the figures; R.J.D. and A.M.H. drafted the manuscript; R.J.D., A.M.H., P.D.U., and N.W.M. edited and revised the manuscript; R.J.D., A.M.H., J.D., L.L., D.C.B., P.D.U., and N.W.M. approved the final version of the manuscript.