blocks hypoxia-induced pulmonary vascular

Dominant negative mutation of the TGF-β receptor blocks hypoxia-induced pulmonary vascular remodeling

Yiu-Fai Chen, Ji-An Feng, Peng Li, Dongqi Xing, Yun Zhang, Rosa Serra, Namasivayam Ambalavanan, Erum Majid-Hassan and Suzanne Oparil J Appl Physiol 100:564-571, 2006. First published 13 October 2005; doi: 10.1152/japplphysiol.00595.2005 You might find this additional info useful... This article cites 35 articles, 15 of which you can access for free at: http://jap.physiology.org/content/100/2/564.full#ref-list-1 This article has been cited by 24 other HighWire-hosted articles: http://jap.physiology.org/content/100/2/564#cited-by Updated information and services including high resolution figures, can be found at: http://jap.physiology.org/content/100/2/564.full

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J Appl Physiol 100: 564 –571, 2006. First published October 13, 2005; doi:10.1152/japplphysiol.00595.2005.

Dominant negative mutation of the TGF-␤ receptor blocks hypoxia-induced pulmonary vascular remodeling Yiu-Fai Chen,1 Ji-An Feng,1 Peng Li,1 Dongqi Xing,1 Yun Zhang,1 Rosa Serra,2 Namasivayam Ambalavanan,3 Erum Majid-Hassan,1 and Suzanne Oparil1 1

Vascular Biology and Hypertension Program, Division of Cardiovascular Disease, Department of Medicine, Department of Cell Biology, and Division of Neonatology, 3Department of Pediatrics, Birmingham, Alabama

2

Submitted 19 May 2005; accepted in final form 7 October 2005

transforming growth factor; vascular hypertrophy and muscularization; alveolar remodeling; extracellular matrix HYPOXIC STRESS RESULTS IN pulmonary vasoconstriction, hypertension, and vascular remodeling, ultimately leading to right heart failure and death by upsetting the balance in the normal relationships between vasoconstrictor and vasodilator and between mitogenic and growth-inhibiting pathways in lung. During chronic hypoxic exposure, vascular smooth muscle cells (VSMCs) resident in normally muscularized arteries undergo hypertrophy and hyperplasia, while new VSMCs appear in intra-acinar arteries that are partially muscularized or nonmuscularized in the normoxic state (24 –26, 34). Extracellular matrix (ECM, e.g., collagen) deposition also contributes to

Address for reprint requests and other correspondence: Y.-F. Chen, 1008 Zeigler Research Bldg., Dept. of Medicine, Univ. of Alabama at Birmingham, UAB Station, Birmingham, AL 35294-0007 (e-mail: [email protected]). 564

pulmonary vascular remodeling during hypoxic exposure (8, 21, 24, 25). Transforming growth factor-␤s (TGF-␤s), a family of multifunctional cytokines consisting of three mammalian isoforms, -␤1, -␤2, and -␤3, are key mediators of pulmonary morphogenesis and pulmonary fibrosis and vascular remodeling (1, 3, 31, 36). Expression and activation of TGF-␤s are increased under stressful conditions, including hypoxia (3, 26, 30), and TGF-␤s stimulate proliferation of pulmonary artery smooth muscle cells (PASMCs) and production of ECM in lung (3, 26). TGF-␤1 is the predominant isoform involved in fibrotic tissue remodeling and is overexpressed in areas of active fibrosis in lung (5), as well as in several animal models of pulmonary hypertension (6). All TGF-␤ isoforms signal through membrane bound heteromeric type I (TGF␤RI) and type II (TGF␤RII) receptors. The kinase domain of TGF␤RII is essential to all TGF-␤ signaling. Activation of TGF-␤ receptors transduces intracellular signals via activation of Smad proteins (13, 19, 20) and mediates tissue remodeling by increasing the production and decreasing the degradation of ECM molecules (3, 36). However, to date studies of the role of TGF-␤s in the pathogenesis of pulmonary hypertension and vascular and/or alveolar parenchymal remodeling in adult animals have been limited by the unavailability of appropriate animal models and reagents. Mice with null mutations of the TGF-␤ isoforms have early lethality, and it can be difficult to distinguish between the primary and secondary effects of the mutations during the development (7). Furthermore, no selective antibodies against TGF-␤ ligands or selective antagonists of TGF-␤ receptors are currently available. In the present study, we used a novel transgenic mouse model that expresses an inducible dominant negative mutation of the TGF-␤ type II receptor gene (DnTGF␤RII mouse) (28) and thus lacks a functional TGF-␤ type II receptor response to all three TGF-␤ isoforms to test the hypothesis that increased pulmonary TGF-␤ signaling during hypoxia participates in the pathogenesis of chronic hypoxia-induced pulmonary hypertension and vascular and/or alveolar remodeling and that disruption of the TGF-␤ signaling cascade attenuates the development of hypoxia-induced pulmonary hypertension and vascular and/or alveolar remodeling. Nontransgenic (NTG) mice exposed to hypoxia or room air were used as controls. MATERIALS AND METHODS

Transgenic mice and animal preparation. DnTGF␤RII transgenic mice originally generated in the laboratory of Dr. Rosa Serra (28) as The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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Chen, Yiu-Fai, Ji-An Feng, Peng Li, Dongqi Xing, Yun Zhang, Rosa Serra, Namasivayam Ambalavanan, Erum Majid-Hassan, and Suzanne Oparil. Dominant negative mutation of the TGF-␤ receptor blocks hypoxia-induced pulmonary vascular remodeling. J Appl Physiol 100: 564 –571, 2006. First published October 13, 2005; doi:10.1152/japplphysiol.00595.2005.—The present study utilized a novel transgenic mouse model that expresses an inducible dominant negative mutation of the transforming growth factor (TGF)-␤ type II receptor (DnTGF␤RII mouse) to test the hypothesis that TGF-␤ signaling plays an important role in the pathogenesis of chronic hypoxia-induced increases in pulmonary arterial pressure and vascular and alveolar remodeling. Nine- to 10-wk-old male DnTGF␤RII and control nontransgenic (NTG) mice were exposed to normobaric hypoxia (10% O2) or air for 6 wk. Expression of DnTGF␤RII was induced by drinking 25 mM ZnSO4 water beginning 1 wk before hypoxic exposure. Hypoxia-induced increases in right ventricular pressure, right ventricular mass, pulmonary arterial remodeling, and muscularization were greatly attenuated in DnTGF␤RII mice compared with NTG controls. Furthermore, the stimulatory effects of hypoxic exposure on pulmonary arterial and alveolar collagen content, appearance of ␣-smooth muscle actin-positive cells in alveolar parenchyma, and expression of extracellular matrix molecule (including collagen I and III, periostin, and osteopontin) mRNA in whole lung were abrogated in DnTGF␤RII mice compared with NTG controls. Hypoxic exposure had no effect on systemic arterial pressure or heart rate in either strain. These data support the hypothesis that endogenous TGF-␤ plays an important role in pulmonary vascular adaptation to chronic hypoxia and that disruption of TGF-␤ signaling attenuates hypoxia-induced pulmonary hypertension, right ventricular hypertrophy, pulmonary arterial hypertrophy and muscularization, alveolar remodeling, and expression of extracellular matrix mRNA in whole lung.

ROLE OF TGF-␤ IN HYPOXIA-INDUCED PULMONARY HYPERTENSION AND REMODELING

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analyzer (Radiometer, Copenhagen, Denmark). For the hypoxic groups, blood pressure measurement and blood sampling were done inside the hypoxic chamber to avoid complications due to normoxia. Histological and immunohistochemical analyses. Paraformaldehyde-fixed lungs were paraffin embedded and sectioned for examination of collagen content by picrosirius red staining or for fibronectin and ␣-smooth muscle actin (␣-SMA) expression by using the selective AB2033 anti-fibronectin antibody (Chemicon, Temecula, CA) and clone1A4 anti-␣-SMA antibody (Dako, Carpinteria, CA), respectively, as previously described (35). Quantitative morphometric analysis of pulmonary vascular and parenchymal content of collagen and ␣-SMA was carried out by light microscopy with a Qimaging QiCam fast cooled color charge-coupled device 12-bit camera (Qimaging, Burnaby, BC, Canada) interfaced with a computer system running MetaMorph 6.2v4 software (Universal Imaging, Downingtown, PA) with high-resolution color images (1,392 ⫻ 1,040 pixels). At least 36 pulmonary arteries (defined as vessels that accompanied airways, ranging in external diameter from 15 to 200 ␮m) and 12 parenchymal areas from different ⫻400 fields were evaluated from each lung section. Pulmonary arterial wall thickness was measured along the shortest curvature of the lumen diameter. Lumen diameter (distance within internal elastic lamella), external vessel diameter (distance within external elastic lamella), and medial thickness (distance between external and internal elastic lamellae) were measured. Percentage medial wall thickness [% WT ⫽ (2 ⫻ medial WT)/external diameter ⫻ 100] was calculated (9). Pulmonary arterial muscularization was assessed using ␣-SMAimmunostained lung sections according to the method of Jones et al. (10). Arterial muscularization was defined according to the degree of muscularization: muscularized arteries (with two distinct elastic laminae and complete medial coat); partially muscularized arteries (with a continuous external elastic lamina and an incomplete medial coat); and nonmuscularized vessels (with only one single elastic lamina but no VSMC apparent) were distinguished by observation and counted. The percentage of muscularization of each pulmonary artery relative to its size (15–200 ␮m in diameter) was calculated as an index of pulmonary arterial muscularization. For quantitative evaluation of alveolar remodeling, mean linear intercepts (MLI, used as an index of alveolar size) (18), number of ␣-SMA positive cells, and amount of interstitial collagen and fibronectin in alveolar parenchyma were measured by using the Metamorph image-analysis system, and the volume percent collagen and fibronectin were calculated as described previously (2). All slides were stained at the same time to minimize variation in staining intensity to avoid interobserver variation. Morphometric analysis was carried out by two independent examiners who were blinded with respect to the treatment assignment of the tissue samples examined. RNA extraction for RT-PCR analysis of DnTGF␤RII mRNA expression and Northern blot analysis of ECM molecule mRNAs. RNA was isolated from snap-frozen tissue by a modification of the acid guanidinium thiocyanate extraction method as described previously (32). For RT-PCR analysis, RNA was treated with RNase-free DNase for 30 min at 37°C to remove contaminating genomic DNA. cDNA was synthesized from 1 ␮g of total RNA pooled from tissue using random primers as described in the GeneAmp RNA PCR kit (Perkin Elmer, Norwalk, CT). PCR amplification of DnTGF␤RII cDNA was performed by using 2 ␮l of the cDNA mix as described previously (28). The conditions and primers described above for the amplification of DnTGF␤RII from genomic DNA were used. Samples from reactions performed in the absence of reverse transcriptase were amplified to demonstrate that there was no contaminating genomic DNA in the RNA samples. Northern analysis was performed using 32P-labeled selective cDNA probes for the ECM molecules collagen I (Col I), collagen III (Col III), periostin (PN), and osteopontin (OPN) that had been generated in our laboratory by reverse transcription (RT) followed by

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well as NTG wild-type mice of the C57BL/6 strain were studied. All mice were raised in our resident colonies, which were founded with pathogen-free breeding pairs. Genotypes were identified by polymerase chain reaction (PCR) assay of genomic DNA from tail snips after weaning (3, 11, 32). The two primer pairs for PCR were 5⬘-ATCGTC-ATC-GTC-TTT-GTA-GTC-3⬘ and 5⬘-TCC-CAC-CGC-ACGTTC-AGA-AG-3⬘ for the DnTGF␤RII allele. Because the DnTGF␤RII mouse overexpresses the DnTGF␤RII gene under the control of a metallothionein-derived promoter, expression of DnTGF␤RII gene was induced by giving 25 mM ZnSO4 in the drinking water beginning 1 wk before hypoxic exposure and continuing throughout the study. NTG mice drinking either 25 mM ZnSO4 water or tap water served as controls. All mice were housed three or four per cage, maintained at constant humidity (60 ⫾ 5%), temperature (24 ⫾ 1°C), and light cycle (6 AM to 6 PM), and fed a standard diet (0.55% NaCl, Harlan-Teklad, Madison, WI). All protocols were approved by the Institutional Animal Care and Use Committee at the University of Alabama at Birmingham and were consistent with the Guide for Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH publication no. 96-01, revised 1996). Hypoxic exposure. Male homozygous DnTGF␤RII (drinking 25 mM ZnSO4 water) and NTG (half drinking 25 mM ZnSO4 water and half drinking tap water) mice were exposed to hypoxia in a 800-liter model 818GBB Plexiglas glove box (Plas Labs, Lansing, MI) beginning at age 8 –9 wk for a total of 6 wk as previously described (32). Hypoxic exposures (range 10.0 ⫾ 0.5% O2) were accomplished by intermittently adding N2 gas to the chamber from a liquid N2 reservoir, controlled by the recorder output of a model 1630 O2 controller (Engineered System & Designs). A baralyme CO2 scrubber kept the CO2 concentration at ⬍0.2%. Relative humidity within the chamber was kept at ⬍60% with anhydrous CaSO4 and a constant temperature circulator (Polyscience, Niles, IL). Boric acid was used to keep NH3 levels within the chamber at a minimum. Daily animal maintenance was carried out without interruption of the exposures through double ports in the chamber. Normoxic (air) control animals were caged similarly and were exposed to filtered room air for identical periods. Right ventricular pressure measurement and tissue collection. After 6 wk of hypoxic or normoxic exposure, mice were weighed and anesthetized with a mixture of ketamine (80 mg/kg ip) and xylazine (12 mg/kg ip). Right ventricles (RV) were cannulated in situ by a closed-chest technique for RV pressure determination as an index of pulmonary hypertension as described previously (32). Mice were then cervically dislocated, and organs were removed for weighing and histological analysis. To assess the development of RV hypertrophy, hearts were dissected free, and the RV was carefully separated from the left ventricle and septum (LV⫹S). Ventricles were blotted dry then weighed separately to determine the index of RV hypertrophy based on the RV free wall weight (adjusted to body weight), left ventricle (LV) weight, and RV to LV⫹S ratio. Half of the lungs from hypoxia-adapted and air control mice were fixed in situ (without weighing) in the distended state by infusion of 10% buffered formalin into the pulmonary artery (at 25 mmHg pressure) and trachea for 1 min, and then they were placed in a bath of 4% paraformaldehyde for 24 h for immunohistochemical analysis. The remaining lungs and brain, heart, intestine, kidney, liver, pancreas, and spleen were immediately frozen in liquid nitrogen for RNA isolation. Systemic arterial pressure, heart rate, blood gas, pH, electrolytes, and hematocrit analyses. In separate groups of hypoxic and air control DnTGF␤RII and NTG mice, PE-10 catheters were implanted into the left common carotid artery through the left external carotid artery under ketamine-xylazine anesthesia. After mice recovered from anesthesia, mean arterial pressure and heart rate were measured simultaneously as described previously (33). In additional groups of mice, 500 ␮l of blood were withdrawn from arterial catheter for blood-gas and hematocrit measurement. Arterial pH, arterial PO2, arterial PCO2, and hematocrit were measured with a model ABL 1640 blood-gas

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RESULTS

Effects of Zn2⫹ on DnTGF␤RII mRNA in tissues. The DnTGF␤RII transgene contains metal-responsive elements and the expression of DnTGF␤RII mRNA was induced in lung and all other tissues examined, including brain, kidney, liver, intestine, pancreas, spleen, and LV by having the mice drink 25 mM ZnSO4 water for 1 wk. Effects of chronic hypoxia on RV pressure and weight. No differences in RV or LV⫹S weight, RV-to-LV⫹S ratios (Fulton index) (Table 1), or mean RV pressure (MRVP) between NTG mice drinking 25 mM ZnSO4 water and NTG mice drinking tap water were observed under either normoxic or hypoxic conditions. Therefore, data from these two groups were combined (Fig. 1). After 6 wk of hypoxic exposure, MRVP, the index of pulmonary arterial pressure, was significantly lower in the DnTGF␤RII than in the NTG mice (by

Fig. 1. Effects of 6-wk hypoxic exposure (10% O2, 1 atm) on mean right ventricular pressure (MRVP; A), and right ventricular (RV)-to-left ventricle and septum (LV⫹S) weight ratio (B) in transforming growth factor-␤ type II receptor dominant negative (DnTGF␤RII) mice and nontransgenic (NTG) wild-type mice. Half of the NTG mice were given 25 mM ZnSO4 in their drinking water throughout the study. Because ZnSO4 does not affect MRVP or RV weight (Table 1), data were combined. Results are means ⫾ SE; n, number of mice per group. *P ⬍ 0.05 vs. respective air control groups. #P ⬍ 0.05 vs. respective NTG groups.

Table 1. Effects of 6-wk hypoxic exposure (10% O2, 1 atm) and/or 25 mM ZnSO4 in drinking water on tissue weights, collagen volume in media of pulmonary arteries, MAP, HR, hematocrit, arterial blood pH, PaO2, PaCO2, and electrolytes of DnTGF␤RII and NTG mice NTG ⫹ ZnSO4

NTG

DnTGF␤RII ⫹ ZnSO4

Treatment

Air

Hypoxia

Air

Hypoxia

Air

Hypoxia

RV, mg LV⫹S, mg RV/(LV⫹S) Kidney, mg Liver, mg Collagen volume on pulmonary artery, % MAP, mmHg HR, beats/min Hematocrit, % PaO2, Torr PaCO2, Torr pH Na⫹, mM K⫹, mM

22.7⫾0.4 89.3⫾1.4 0.25⫾.01 346⫾11 1,303⫾64

33.5⫾0.6* 87.0⫾2.5 0.38⫾.02* 334⫾11 1,245⫾65

24.6⫾0.8 85.8⫾2.9 0.28⫾.02 354⫾10 1,055⫾30 4.98⫾.63 107⫾4 378⫾12 44.8⫾1.2 105.7⫾3.1 32.4⫾3.3 7.37⫾.03 154⫾3 3.4⫾0.1

34.6⫾1.4* 85.9⫾1.9 0.41⫾.02* 325⫾13 1,067⫾23 6.86⫾.42* 109⫾4 390⫾19 65.3⫾1.1* 50.2⫾3.0* 19.8⫾1.4* 7.40⫾0.01 154⫾1 3.6⫾0.1

25.0⫾0.8 93.9⫾0.6 0.27⫾.01 484⫾14 1,293⫾25 2.31⫾.23† 108⫾2 418⫾30 46.9⫾1.0 99.1⫾2.2 31.7⫾2.1 7.39⫾.01 150⫾2 4.1⫾.3

30.3⫾1.6*† 93.1⫾1.4 0.32⫾.02*† 420⫾10* 1,221⫾23 3.43⫾.57† 103⫾3 407⫾16 59.9⫾1.1*† 47.9⫾4.4* 21.2⫾1.4* 7.37⫾0.01 155⫾1 3.6⫾0.2

Values are means ⫾ SE; n ⫽ 7–10 mice per group. Pulmonary arteries were ⬎75 ␮m in external diameter, n ⫽ 24 vessels per mouse were measured. RV, right ventricle; LV⫹S, left ventricle⫹septum; MAP, mean systemic arterial pressure; HR, heart rate; PaO2 and PaCO2, arterial PO2 and PCO2, respectively. DnTGF␤RII, transforming growth factor-␤ type II receptor dominant negative mutation mice. Adjusted tissue weights were determined by analysis of covariance with body weight as a covariate. *P ⬍ 0.05 vs. respective air control groups. †P ⬍ 0.05 vs. respective nontransgenic (NTG) groups. J Appl Physiol • VOL

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the DNA polymerase chain reaction (PCR) using lung RNA as the template, as previously described (32, 33). A 32P-labeled 18S rRNAoligonucleotide (5⬘-ACGGTATCTGATCGTCTTCGAACC-3⬘) was used as the control probe to normalize data. Autoradiographic signals were scanned with an optical densitometer (model GS-670 Imaging Densitometer, Bio-Rad, Hercules, CA). To estimate steady-state specific mRNA levels, Col I, Col III, PN, and OPN mRNA-to-18S rRNA ratios were determined by dividing the absorbance corresponding to the specific cDNA probe hybridization by the absorbance corresponding to the 18S rRNA probe hybridization. Statistical analysis. Results were expressed as means ⫾ SE. Tissue weights were adjusted by analysis of covariance with body weight as the covariate (23). Statistical analyses were carried out using the SigmaStat statistical package (Jandel Scientific Software, San Rafael, CA). The data were analyzed by two-way ANOVA to test for separate and combined effects of genotype and hypoxia on tissue weights, RV pressure, pulmonary arterial wall thickness, and muscularization. One-way ANOVA followed by the Newman-Keuls test was used to test the effects of genotype on the above variables within the hypoxic and normoxic groups. The unpaired t-test was used to test the effects of hypoxia on the above variables within each genotype. Differences were reported as significant if the P value was ⬍0.05.

ROLE OF TGF-␤ IN HYPOXIA-INDUCED PULMONARY HYPERTENSION AND REMODELING

0.63, P ⬍ 0.05). The hypoxia-induced increase in wall thickness of small pulmonary arteries (⬍25–100 ␮m in diameter) was significantly less in DnTGF␤RII mice than in normoxic NTG mice (P ⬍ 0.05, by two-way ANOVA, Fig. 2, B–D). The degree of hypoxia-induced muscularization in small pulmonary arteries (⬍25 to 100 ␮m in diameter) was parallel to the extent of pulmonary arterial wall thickening (Figs. 2 and 3). The large pulmonary arteries (⬎100 ␮m) were fully muscularized (with two distinct elastic laminae and a complete medial coat) in both genotypes under both normoxic and hypoxic conditions. After 6-wk hypoxic exposure, muscularization extended into smaller arteries accompanying terminal and respiratory bronchioles (25- to 100-␮m-diameter vessels) and alveolar walls in both genotypes. Hypoxia-induced muscularization was less marked in arteries (25–100 ␮m) of DnTGF␤RII mice than in NTG mice (Fig. 3, B and C). More of the precapillary pulmonary arterioles (⬍25 ␮m) located in the alveolar ducts and alveolar walls were muscularized in hypoxic NTG mice than in hypoxic DnTGF␤RII mice (Fig. 3D). Hypoxia-induced muscularization (both in percentage and number) of precapillary pulmonary arterioles (⬍25 ␮m) was significantly attenuated in DnTGF␤RII mice, compared with NTG mice (P ⬍ 0.05, by two-way ANOVA, Fig. 3D). Effects of chronic hypoxia on ␣-SMA expression and collagen and fibronectin content in alveolar parenchyma and pulmonary artery. The density of ␣-SMA-positive cells in alveolar parenchyma in the two genotypes was parallel to the alveolar parenchymal collagen and fibronectin content (Figs. 4 and 5). Under air control conditions, the density of ␣-SMApositive cells in alveolar parenchyma (expressed as ␣-SMA cell numbers/100 alveoli) was low in both genotypes (Figs. 4A, 4C, and 5A). Exposure to hypoxia significantly increased the density of ␣-SMA-positive cells in alveolar parenchyma of

Fig. 2. Effects of 6-wk hypoxic exposure (10% O2, 1 atm) on wall thickness (Wt) of pulmonary arteries ranging between ⬎100 ␮m (external diameter; A), 51–100 ␮m (B), 25–50 ␮m (C), and ⬍25 ␮m (D) in DnTGF␤RII and NTG mice. Results are means ⫾ SE; n ⫽ number of vessels from total 7–9 mice per group. *P ⬍ 0.05 vs. respective air control groups. #P ⬍ 0.05 vs. respective NTG groups.

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one-way ANOVA). The absolute increase in MRVP with hypoxic exposure was significantly less in DnTGF␤RII mice (a 36% or 2.7 mmHg increase compared with its air control), than in NTG mice (a 63% or 4.9-mmHg increase compared with its air control) (P ⬍ 0.05, by two-way ANOVA). These findings reflect an attenuated hypoxia-induced increase in pulmonary arterial pressure in DnTGF␤RII mice compared with hypoxiaadapted NTG mice. No differences in mean arterial pressure, heart rate, blood pH, or sodium and potassium concentrations between DnTGF␤RII mice were observed under either normoxic or hypoxic conditions (Table 1). Hypoxic exposure caused similar decreases in arterial PO2 and increases in arterial PCO2 in DnTGF␤RII and NTG mice (Table 1). Hematocrit increased in both strains in response to hypoxia, but to significantly lower levels in the DnTGF␤RII than in the NTG mice (Table 1, by one-way ANOVA). The mechanism for the attenuated hypoxia-induced polycythemic response in DnTGF␤II mice is unclear. The hypoxia-induced increase in RV weight was significantly less in DnTGF␤RII mice (an 18% or 4.5-mg increase compared with its air control) than in NTG mice (a 44% or 10.4-mg increase compared with its air control) (P ⬍ 0.05, by two-way ANOVA), again reflecting strain differences in the severity of the hypoxia-induced pulmonary hypertension. Chronic hypoxic exposure did not alter LV weight in any genotype (Table 1). Effects of chronic hypoxia on pulmonary arterial hypertrophy and muscularization. Under normoxic conditions, DnTGF␤RII mice had thinner small (51–100 ␮m in diameter) pulmonary arteries, as reflected in the wall thickness index, than NTG mice (Fig. 2B). Chronic hypoxic exposure was associated with increases in pulmonary arterial wall thickness that correlated with the rise in RV pressure (r ⫽ 0.72, P ⬍ 0.05, all mice combined) and the increase in RV weight (r ⫽

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Fig. 3. Effects of 6-wk hypoxic exposure (10% O2, 1 atm) on percentage of muscularization of pulmonary arteries ranging between ⬎100 ␮m (A), 51–100 ␮m (B), 25–50 ␮m (C), and ⬍25 ␮m (D) in external diameter in DnTGF␤RII and NTG mice. Results are means ⫾ SE; n ⫽ number of vessels from total 7–9 mice per group. *P ⬍ 0.05 vs. respective air control groups. #P ⬍ 0.05 vs. respective NTG groups.

induced increase in collagen content in pulmonary artery was abrogated in DnTGF␤RII mice compared with NTG (Table 1). Exposure to chronic hypoxia significantly increased collagen and fibronectin volume in alveolar parenchyma of NTG mice but had no stimulatory effect on collagen content in DnTGF␤RII mice (Fig. 5, C and D). Effects of chronic hypoxia on ECM molecule mRNA expression in lung. Baseline (air control) Col I and III, PN, and OPN mRNA levels in lung were similar in both genotypes (Fig. 6). Six-week normobaric hypoxic exposure significantly increased

Fig. 4. Representative light micrographs of lungs from DnTGF␤RII (A and B) and NTG (C and D) mice exposed to hypoxia (10% O2, 1 atm) or room air for 6 wk. Lung sections were immunostained with ␣-smooth muscle actin (␣-SMA) antibody and counterstained with hematoxylin. Arrows indicate representative ␣-SMApositive alveolar cells.

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NTG mice (5-fold increase, Figs. 4D and 5A), but not of DnTGF␤RII mice (Figs. 4B and 5A), indicating that alveolar myofibroblast transformation was attenuated in DnTGF␤RII mice compared with the control strain under hypoxic conditions. Baseline (air control) average alveolar diameter, represented by the MLI of alveolar parenchyma, was similar in both genotypes (Fig. 5B). Chronic hypoxic exposure had no significant effect on MLI in either DnTGF␤RII or NTG mice. Baseline (air control) pulmonary arterial collagen content was significantly lower in DnTGF␤RII mice, and the hypoxia-

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Col I (⫹46%), Col III (⫹33%), PN (⫹41%), and OPN (⫹45%) in the NTG control strain, compared with their respective air control groups. In contrast, chronic hypoxic exposure had no significant effect on expression of these ECM molecules in lung of DnTGF␤RII mice. DISCUSSION

The present study was the first to test the pulmonary vascular response to chronic hypoxic stress in an animal with an inducible defect in the TGF-␤ signaling pathway. The most striking finding of the study is that disruption of TGF-␤ type II receptor expression in lung greatly attenuates the chronic hypoxia-induced phenotypic changes (pulmonary hypertension, RV enlargement, pulmonary arterial remodeling and muscularization, myofibroblast transformation of alveolar fibroblasts, and enhanced ECM mRNA and protein expression) in DnTGF␤RII mice compared with NTG control mice. These dramatic findings in the DnTGF␤RII model support our hypothesis that endogenous TGF-␤ plays an important role in the pulmonary response to chronic hypoxic stress. All three isoforms of TGF-␤ as well as TGF␤RI and -RII are expressed at high levels during normal lung development, and expression of TGF-␤1 and -␤3 is increased in pathological conditions, including hypoxia-induced pulmonary hypertension in adult animals (3, 4, 26, 36). However, because null mutations of TGF-␤s and TGF␤RII are embryonic lethal (7, 22), previous whole animal studies of the functional role of TGF-␤ in normal pulmonary morphogenesis and in the pathogenesis of pulmonary fibrosis and vascular remodeling were limited. The DnTGF␤RII mouse offers the important advantage of not disrupting critical TGF-␤ signaling pathways during development and enables the in vivo functions of J Appl Physiol • VOL

TGF-␤ to be rigorously analyzed in the adult lung under stress conditions. Active TGF-␤ ligands bind to a membrane-bound complex of TGF␤RI and TGF␤RII kinase that transduces intracellular signals via activation of Smad proteins (13, 19, 20). When overexpressed, a cytoplasmically truncated TGF␤RII receptor can compete with endogenous receptors for heterodimeric complex (TGF␤RI and -RII) formation, thereby acting as a dominant-negative mutant. The DnTGF␤RII mouse overexpresses the DnTGF␤RII gene under the control of a metallothionein-derived promoter that can be induced by Zn2⫹ in adult animals, thus offering the important advantage of transient disruption of TGF-␤ signaling in adult mice, without permanently terminating critical TGF-␤ signaling pathways in the developing lung (3). The DnTGF␤RII mouse model has been used in studies of bone development and mammary gland morphogenesis (28, 29) but never to test the role of TGF-␤ in cardiopulmonary disorders. We were the first to systematically assess the organ distribution of DnTGF␤RII mRNA transcripts in Zn2⫹-treated DnTGF␤RII mice. We observed robust expression of this gene transcript in lung of air control DnTGF␤RII mice 1 wk after Zn2⫹ treatment. Therefore, this novel animal model is suitable for assessing the role of TGF-␤ signaling in pulmonary responses to hypoxic stress. We found that NTG mice exposed to chronic hypoxia exhibit increased RV pressure and hypertrophy in association with hypertrophy and muscularization of distal pulmonary arterioles. These findings are consistent with previous reports that the development of hypoxia-induced pulmonary hypertension is associated with structural changes in the pulmonary vascular bed with increased medial muscular wall thickness in normally muscular pulmonary arteries and with extension of

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Fig. 5. Effects of 6-wk hypoxic exposure (10% O2, 1 atm) on density of ␣-SMA-positive cells in alveoli (A), mean linear intercept (MLI), representing the average alveolar diameter (B), collagen volume (C), and fibronectin volume (D) in alveolar parenchyma of DnTGF␤RII and NTG mice. n, Number of mice per group. Twelve parenchymal areas per mouse were measured. *P ⬍ 0.05 vs. respective air control groups. #P ⬍ 0.05 vs. respective NTG groups (bottom).

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ROLE OF TGF-␤ IN HYPOXIA-INDUCED PULMONARY HYPERTENSION AND REMODELING

muscularization into smaller and more peripheral arteries in humans and in animal models (8, 17, 24). We observed dramatic attenuation of RV hypertension, pulmonary vascular hypertrophy, and muscularization in DnTGF␤RII mice compared with NTG controls under hypoxic conditions, supporting the hypothesis that signaling through the TGF␤RII plays a critical role in mediating pulmonary vascular remodeling in hypoxia-adapted animals. Using quantitative morphometry, we found that expression of ␣-SMA, a first marker of smooth muscle cell differentiation, was attenuated in alveolar parenchyma of DnTGF␤RII mice compared with NTG mice exposed to chronic hypoxia. These results are consistent with our in vitro observations of robust TGF-␤-mediated myofibroblast transformation of mouse cardiac fibroblasts (36). The present finding suggests that TGF␤-stimulated alveolar myofibroblast transformation contributes to pulmonary parenchymal remodeling in response to chronic hypoxic stress. TGF-␤ mediates fibrotic tissue remodeling by increasing the production and decreasing the degradation of ECM molecules (3, 13, 27, 36). The present study demonstrated increased J Appl Physiol • VOL

expression of Col I, Col III, OPN, and PN, a novel ECM molecule originally described in bone (14, 16), in lung of NTG mice adapted to hypoxia as part of a generalized ECM response to hypoxic stress. We have found a similar functionally significant stimulatory effects of TGF-␤ or hypoxia on ECM expression in isolated PASMCs in vitro (14, 15). We demonstrated that TGF-␤1 dose dependently enhanced expression of PN and OPN mRNA in isolated rat PASMCs (15). These findings, coupled with our present observation that the hypoxia-induced ECM mRNA expression was abolished in lung of DnTGF␤RII mice, suggest that TGF-␤ signaling is a critical mediator of ECM gene expression in hypoxic lung. Consistent with the attenuation in vascular remodeling and ECM mRNA expression in whole lung in hypoxia-adapted DnTGF␤RII mice, we found no hypoxia-induced increase in pulmonary arterial collagen content or alveolar parenchymal collagen or fibronectin volume in this strain. This contrasts with the significant increases in all of these parameters that we observed in the control NTG strain and with published reports of increased collagen and fibronectin expression in pulmonary arteries in a variety of experimental models of pulmonary

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Fig. 6. Effects of 6-wk hypoxic exposure (10% O2, 1 atm) on steady-state collagen I (Col I; A), collagen III (Col III; B), periostin (PN; C), and osteopontin (OPN; D) mRNA expression in DnTGF␤RII and NTG mouse lung. Northern blot analysis was carried out with 15 ␮g of total lung RNA from each mouse. The Northern blot membrane was probed with PN, OPN, Col I, Col III, and 18S rRNA sequentially. The mRNA loading was normalized by 18S rRNA. The mRNAs from air control and hypoxic DnTGF␤RII and NTG mice were run in pairs on the same gel. mRNAto-18S RNA ratios were normalized and standardized to mean mRNA level of NTG air control mice. Results are means ⫾ SE; n, number of mice. *P ⬍ 0.05 vs. respective air control groups. #P ⬍ 0.05 vs. respective NTG groups. Representative Northern blot images are shown at top.

ROLE OF TGF-␤ IN HYPOXIA-INDUCED PULMONARY HYPERTENSION AND REMODELING

hypertension (12, 21). It is likely that the reduced deposition of collagen and other ECM molecules in the pulmonary arterioles of the DnTGF␤RII mice contributed to the maintenance of near-normal compliance and resistance characteristics of the pulmonary circulation under hypoxic conditions, thus minimizing the development of pulmonary hypertension. In summary, the present study provides the first evidence that chronic hypoxia-induced pulmonary hypertension and vascular and parenchymal remodeling are attenuated in absence of TGF-␤ signaling. These data support the hypothesis that endogenous TGF-␤ plays an important role in regulating pulmonary artery pressure, ECM production, and pulmonary vascular remodeling in response to hypoxic stress. The DnTGF␤RII mouse provides a novel model with which to study the role of TGF-␤ signaling in the lung under stress conditions.

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ACKNOWLEDGMENTS 21.

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This work was supported in part by National Institutes of Health Grants HL-44195, HL-50147, HL-45990, HL-07457, HL-56046, and HD-46513, and American Heart Association Grant 0455197B.

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