Contributed by Gregg L. Semenza, February 15, 2012 (sent for review January 12, 2012)
Chronic systemic hypertension causes cardiac pressure overload leading to increased myocardial O2 consumption. Hypoxia-inducible factor 1 (HIF-1) is a master regulator of O2 homeostasis. Mouse embryos lacking expression of the O2-regulated HIF-1α subunit die at midgestation with severe cardiac malformations and vascular regression. Here we report that Hif1af/f;Tie2-Cre conditional knockout mice, which lack HIF-1α expression only in Tie2+ lineage cells, develop normally, but when subjected to pressure overload induced by transaortic constriction (TAC), they manifest rapid cardiac decompensation, which is accompanied by excess cardiac fibrosis and myocardial hypertrophy, decreased myocardial capillary density, increased myocardial hypoxia and apoptosis, and increased TGF-β signaling through both canonical and noncanonical pathways that activate SMAD2/3 and ERK1/2, respectively, within endothelial cells of cardiac blood vessels. TAC also induces dilatation of the proximal aorta through enhanced TGF-β signaling in Hif1af/f;Tie2-Cre mice. Inhibition of TGF-β signaling by treatment with neutralizing antibody or pharmacologic inhibition of MEK–ERK signaling prevented TAC-induced contractile dysfunction and pathological remodeling. Thus, HIF-1 plays a critical protective role in the adaptation of the heart and aorta to pressure overload by negatively regulating TGF-β signaling in endothelial cells. Treatment of wild-type mice with digoxin, which inhibits HIF-1α synthesis, resulted in rapid cardiac failure after TAC. Although digoxin has been used for decades as an inotropic agent to treat heart failure, it does not improve survival, suggesting that the countertherapeutic effects of digoxin observed in the TAC mouse model may have clinical relevance.
Hypoxia-inducible factor (HIF-1) is a transcriptional activator that functions as a master regulator of oxygen homeostasis in all metazoan species by regulating O2 supply and demand. HIF-1 is a heterodimer composed of HIF-1α and HIF-1β subunits (1), which dimerize and bind to DNA (2). HIF-1α protein levels increase under hypoxic conditions (3) as a result of decreased O2-dependent prolyl hydroxylation, ubiquitination, and degradation (4). HIF-1 activates the transcription of genes that play critical roles in increasing O2 delivery (e.g., by stimulating vascularization) or decreasing O2 consumption (e.g., by inhibiting oxidative phosphorylation and stimulating glycolysis) in response to hypoxia (5). Homozygosity for germ-line null alleles at the Hif1a locus results in lethality by embryonic day 10.5 with cardiac malformations, vascular regression, and impaired erythropoiesis, indicating that all three components of the circulatory system are dependent on HIF-1 for normal development (6, 7).
In the adult, HIF-1 plays a critical role in mediating adaptation to ischemia. To sustain contractile function, the heart requires continuous generation of ATP, over 95% of which is derived from oxidative phosphorylation (8). Myocardial ischemia may result from decreased O2 delivery caused by impaired coronary blood flow or increased O2 consumption caused by pressure overload (9). HIF-1α expression is induced by ischemia in the human heart (10), and polymorphisms at the human HIF1A locus influence the clinical presentation of ischemic heart disease (11, 12). Hif1a+/− mice, which are heterozygous for a knockout allele, develop normally, but the protective effect of ischemic preconditioning on the heart (13) is lost in these mice (14), possibly reflecting the role of HIF-1 in adenosine (15) and nitric oxide production (16).
Chronic systemic hypertension causes cardiac pressure overload. The response to this pathological state involves myocardial hypertrophy, interstitial fibrosis, and microvascular changes that ultimately impair cardiac contractile function, resulting in heart failure (17). Most studies have focused on the pathological effects of pressure overload on cardiomyocytes. When mice were subjected to transaortic constriction (TAC) as a model of pressure overload, HIF-1α expression was induced in the heart by day 3, leading to increased levels of VEGF, which stimulated vascularization so that tissue perfusion was maintained during cardiac hypertrophy; however, by day 14, HIF-1α levels had declined, leading to loss of VEGF expression and a progressive reduction in vascular density and cardiac function between days 14 and 28 (18). Cardiomyocyte-specific knockout of HIF-1α altered baseline cardiac vascularization and contractility (19). Several studies have analyzed the effect of TAC on Hif1a+/− mice and mice with cardiomyocyte-specific knockout of HIF-1α; one study reported reduced vascular density and contractile dysfunction on day 14 after TAC (18), and two other studies reported unchanged (20) or improved (21) cardiac function in knockout compared with WT mice. However, the effects of HIF-1α loss of function in vascular endothelial cells (ECs) on cardiac adaptation to pressure overload have not been reported.
HIF-1 is critical for the production of angiogenic growth factors such as VEGF that activate ECs, but HIF-1 also has important cell-autonomous functions in ECs (22). To explore the role of ECs in the response to pressure overload, we compared the effect of TAC on Hif1af/f;Tie2-Cre (FC) mice and on their Tie2-Cre and Hif1af/f littermates (hereafter, “C mice” and “F mice,” respectively). Surprisingly, within 1 wk after TAC, these mice manifested life-threatening cardiac decompensation that was associated with increased TGF-β signaling via canonical and noncanonical signaling pathways, providing insight into the role of HIF-1 in adaptation of the heart to pressure overload.
FC mice are homozygous for the floxed Hif1a allele and hemizygous for the Tie2-Cre transgene, resulting in Hif1a gene deletion in Tie2+ lineage cells. FC mice developed normally and were indistinguishable from their C or F littermates. Tie2-Cre drives deletion of floxed Hif1a gene sequences in ECs and bone marrow cells (23). Analysis of floxed Hif1a exon 2 sequences by quantitative real-time PCR (qPCR) revealed ˜70% gene deletion in the heart, and analysis of isolated cardiomyocytes revealed ˜60% deletion (Fig. 1A), indicating that cardiomyocytes as well as ECs are affected; these findings are consistent with a previous report of expression in mesenchymal cells in the atrioventricular canal of the developing heart (23). Baseline M-mode echocardiographic studies revealed no significant difference in left ventricle (LV) end systolic diameter (LVESD), LV mass, ejection fraction (EF), or fractional shortening (FS) in FC mice compared with F mice or C mice (Fig. 1B).
Effect of TAC on FC mouse hearts. (A) qPCR analysis of Hif1a exon 2 DNA sequences in hearts from F, C, and FC mice relative to mouse embryo fibroblast (MEF) WT control DNA sample (Left) and isolated cardiomyocytes from FC relative to F mouse hearts (Right). Data are mean ± SEM data (n = 4). *P < 0.05 for FC vs. F, **P < 0.01 vs. F or C. (B) Echocardiographic data (mean ± SEM; n = 4–17) for LVESD, LV mass, EF, and FS before (Basal) and 1 wk after (1W) TAC. *P < 0.05, ***P < 0.001 vs. FC/S, C/T, or F/T. #P < 0.05, ##P < 0.01, ###P < 0.001 vs. FC/S. (C) Mean ± SEM data (n = 4–17) for HW/BW (mg/g) and HW/TL (mg/cm) ratios. **P < 0.01 vs. FC/S. (D) WT mice were treated with daily i.p. injection of saline or digoxin (1 mg/kg) starting 3 d before TAC. Echocardiographic data (mean ± SEM; n = 4–8) were obtained before and 1 wk after TAC. ***P < 0.001 vs. vehicle.
The mice were subjected to an established TAC protocol that reproducibly results in LV hypertrophy after 1 wk, followed by progressive LV dilation and failure over 6 wk (24). However, FC mice subjected to TAC (FC/T mice) decompensated rapidly, with many becoming moribund by day 7. Echocardiography performed before and 1 wk after TAC revealed increased LVESD and decreased EF and FS in FC/T mice compared with F/T or C/T mice (Fig. 1B). The reduction in FS after TAC was greater (from 64.4 ± 2.5% to 34.4 ± 2.8%) in FC/T mice than in F/T mice (from 64.2 ± 1.2% to 48.5 ± 2.2%) or in C/T mice (from 62.3 ± 2.5% to 46.8 ± 3.0%), whereas FC mice subjected to sham surgery (FC/S) showed no significant change in FS (from 62.3 ± 1.2% to 61.0 ± 0.1%). At 1 wk after TAC, LVESD was 50% greater in FC/T hearts than in F/T or C/T hearts and was 200% greater than in FC/S hearts. Mice of all three genotypes showed a similar increase in LV mass, heart weight to body weight (HW:BW) ratio, and HW to tibial length (HW:TL) ratio after TAC (Fig. 1C). Treatment of WT mice with digoxin, which inhibits HIF-1α synthesis (25), also induced heart failure 1 wk after TAC (Fig. 1D).
To explore the cellular basis for heart failure in FC/T mice, heart sections were stained with Masson trichrome, wheat germ agglutinin (WGA), and isolectin B4 for analysis of fibrosis, cardiomyocyte area, and vascular ECs, respectively (Fig. 2A). TAC-induced fibrosis and cardiomyocyte hypertrophy, which were observed in F/T and C/T mice, were potentiated in FC/T mice (Fig. 2B). In addition, hearts from FC/T mice showed a dramatic reduction in vascular ECs that was not observed in F/T or C/T mice.
Histopathological effects of TAC on mouse hearts. (A) One week after sham surgery or TAC, hearts from F, C, and FC mice were sectioned and treated with Masson trichrome stain to identify fibrosis (blue-stained tissue), with fluorescently-labeled WGA (red) to determine myocyte area, or with Griffonia simplicifolia isolectin B4 (green) to identify vascular ECs. (B) Mean ± SEM data (n = 5) for area stained. *P < 0.05, **P < 0.01 vs. F/T or C/T.
To investigate the consequences of reduced capillary density, heart sections were stained with an Ab against activated caspase 3, which revealed increased apoptotic cells in FC/T (2.28 ± 0.24%) as compared with F/T (1.02 ± 0.14%) or C/T (1.05 ± 0.37%) hearts (Fig. 3A). Pimonidazole staining revealed increased myocardial hypoxia 3 d (1.74 ± 0.30%) and 1 wk (3.25 ± 0.27%) after TAC in FC/T mice as compared with F/T mice (0.50 ± 0.10% and 0.97 ± 0.22%, respectively) (Fig. 3B). These results suggest that loss of myocardial blood vessels in FC/T mice resulted in decreased tissue oxygenation, leading to increased cell death.
Effect of TAC on myocardial apoptosis and hypoxia. (A) Analysis of apoptosis 1 wk (1W) after sham surgery or TAC. (Upper) Heart sections were stained with Ab against cleaved caspase 3 (dark brown stain, arrows) to detect apoptosis. (Lower) Mean ± SEM data (n = 4). *P < 0.05 vs. F/T or C/T. (B) Analysis of hypoxia 1wk after sham surgery and 3 d (3D) or 1 wk after TAC. (Upper) Mice were injected with pimonidazole, and heart sections were stained with Ab against pimonidazole to detect hypoxic regions. (Lower) Mean ± SEM data (n = 4 each). *P < 0.05 vs. F/T; #P < 0.05 vs. FC/S; ##P < 0.01 vs. 3D.
In addition to the effects of TAC on cardiac structure and function, FC/T mice also manifested an increase in aortic root diameter of 21.5 ± 2.5% and aortic wall thickening at 1 wk after TAC (Fig. 4 A–C). Doppler mode echocardiography ruled out aortic insufficiency as a cause of diminished EF and FS. Verhoeff–van Gieson staining revealed mild aortic wall thickening with disruption of the internal elastic lamina in some but not all FC/T mice (Fig. 4E). These findings were reminiscent of aortic root abnormalities observed in a mouse model of Marfan syndrome caused by the expression of mutant fibrillin (26, 27).
Pathological effect of TAC on the aortic root in FC mice. (A) Gross pathology demonstrating dilatation of aortic root (arrowheads) in FC/T mice 1 wk after TAC. (B) B-mode and M-mode echocardiography of aorta and heart, respectively. Aortic root diameter and wall thickness are indicated by yellow and white arrows, respectively. (C) Aortic root diameter and wall thickness (mean ± SEM; n = 4–10) were measured before (Basal; open bars) and 1 wk after (1W; closed bars) sham surgery (S) or TAC (T). *P < 0.05 vs. F/T or FC/Basal; **P < 0.01 vs. FC/S-1W, F/T-1W, C/T-1W, and FC/T-Basal. (D) Aortic root diameter was measured before (Basal; open bars) and 1 wk after (1W; closed bars) TAC in mice treated with vehicle, U0126, or neutralizing Ab against TGF-β (NAb). *P < 0.05 vs. FC/T-basal; #P < 0.05 vs. FC/T-1W. (E) Verhoeff–van Gieson staining of proximal aorta revealed wall thickening and disarray of elastic fibers in FC/T mice. (F) Effect of NAb treatment on phosphorylation of ERK1/2. (Left) IB assays of lysates from proximal aorta using Ab that recognizes phosphorylated (p-) or total (t-) ERK1/2 proteins. (Right) Signals from IBs were quantified by densitometry, and the ratio of p-ERK:t-ERK was calculated (mean ± SEM; n = 4–6). ***P < 0.001 vs. F/T; ###P < 0.001 vs. FC/T. (G) Effect of U0126 treatment (U) on phosphorylation of ERK1/2. (Left) IB assays of lysates from proximal aorta using Ab that recognizes p-ERK1/2 or t-ERK1/2 protein. (Right) Signals from IBs were quantified by densitometry, and the ratio of p-ERK:t-ERK was calculated; mean ± SEM data (n = 4–6) are shown. **P < 0.01 vs. F/T or F/T+U; ##P < 0.01 vs. FC/T.
The pathogenesis of aortic dilatation in fibrillin-mutant mice has been shown to involve excessive TGF-β signaling through both the canonical pathway, leading to SMAD2/3 phosphorylation/activation, and the noncanonical pathway, leading to MEK-ERK1/2 phosphorylation/activation (Fig. 5A) (28). Immunoblot (IB) analysis of heart lysates revealed only modest increases in SMAD2 and SMAD3 phosphorylation (Fig. 5B) and ERK1/2 phosphorylation (Fig. 5C) in F/T and C/T mice at 1 wk after TAC; this finding is consistent with late activation of TGF-β signaling that was observed in WT mice with TAC-induced pressure overload (24). However, FC/T mice showed increased SMAD2, SMAD3, and ERK1/2 phosphorylation relative to the two control genotypes (Fig. 5 B and C). In contrast, phosphorylation of JNK and p38 was not increased in FC/T hearts.
Effect of TAC on TGF-β signaling. (A) Schematic of signal transduction in which binding of TGF-β to its receptor (TGF-βR) leads to phosphorylation and activation of the SMAD2 and SMAD3 (p-SMAD2/3) canonical pathway and the MEK1, ERK1, and ERK2 (p-MEK and p-ERK) noncanonical pathway. (B) (Upper) IB assays of heart lysates 1 wk after sham surgery or TAC using Ab that recognizes phosphorylated (p-) or total (t-) SMAD2, SMAD3, and β-actin. (Lower) IB signals were quantified by densitometry, and the ratio of phosphorylated to total SMAD2 and SMAD3 protein (mean ± SEM; n = 4–6) is shown. **P < 0.01 vs. F/T or C/T. (C) (Upper) IB assays of heart lysates using Ab that recognizes p-ERK1/2, t-ERK1/2, and β-actin. (Lower) IB signals were quantified by densitometry, and the ratio of phosphorylated to total ERK1 and ERK2 protein was determined (mean ± SEM; n = 4–6). **P < 0.01 vs. F/T or C/T.
Parenteral administration of a neutralizing Ab against TGF-β (NAb) protected fibrillin-mutant mice from aortic root dilatation (26). In contrast, NAb did not protect WT mice from cardiomyocyte-dependent pathological remodeling in response to TAC because of the failure of NAb to penetrate into the myocardium (24). We hypothesized that aberrant TGF-β signaling in ECs contributed to the severe phenotype of FC/T mice and, if this notion was correct, that NAb administration might be protective. FC mice were treated with a single i.p. injection of NAb (10 mg/kg) at the time of TAC. M-mode echocardiography revealed that NAb protected against pathological effects of TAC on LVESD, EF, and FS in FC/T mice (Fig. 6 A and B) but had no effect on TAC-induced changes in LV mass, HW:BW ratio, or HW:TL ratio (Fig. 6 A and C). NAb also had no effect on TAC-induced fibrosis (Fig. 6D) or cardiomyocyte hypertrophy (WGA; Fig. 6E), whereas the percentage of vascular ECs in the myocardium was increased significantly (Isolectin; Fig. 6E). NAb also prevented aortic root dilatation (Fig. 4D) and disruption of the internal elastic lamina of the aorta (Fig. 4E) in FC/T mice.
Effect of TGF-β NAb on cardiac structure and function. (A and B) Echocardiographic data (mean ± SEM; n = 4–6) before (Basal) and 1 wk after (1W) sham surgery (S) or TAC (T) in mice treated with vehicle (FC/T) or neutralizing Ab (FC/T+NAb). **P < 0.01 vs. FC/T-1W. (C) HW/BW (mg/g) and HW/TL (mg/cm) ratios (mean ± SEM data; n = 4–6). (D) Analysis of fibrosis. (Upper) Masson trichrome staining. (Lower) Mean ± SEM data; n = 4–6. (E) Analysis of cardiomyocyte hypertrophy and capillary density. (Upper) WGA staining of cardiomyocytes and isolectin B4 staining of ECs (Lower) Mean ± SEM data; n = 4–6. *P < 0.05 vs. FC/T.
IB analyses demonstrated that NAb reduced levels of phosphorylated SMAD3, MEK1/2, and ERK1/2 in the myocardium of FC/T mice (Fig. 7A). Immunohistochemistry revealed ERK phosphorylation in the endocardium and blood vessels, which was increased in FC/T (10.50 ± 0.16%) as compared with F/T (3.55 ± 0.03%) hearts and was decreased by NAb (2.20 ± 0.03%) (Fig. 7B). Microarray analysis revealed that expression of two TGF-β–regulated genes, Fst (29) and Ltbp2 (30), which encode follistatin and latent TGF-β–binding protein 2, respectively, was increased in FC/T as compared with F/T hearts and decreased in FC/T+NAb compared with FC/T hearts (Fig. 7C). TGF-β1 and TGF-β2 mRNA expression also was decreased in FC/T+NAb as compared with FC/T hearts. Based on the results presented in Figs. 5–7, we conclude that excess TGF-β signaling in HIF-1α–deficient ECs plays a key role in the pathogenesis of TAC-induced cardiac failure in FC/T mice through disruption of the cardiac microvasculature.
Effect of NAb on TGF-β signaling. (A) IB assays. (Left) Heart lysates were analyzed using Ab against phosphorylated (p-) or total (t-) SMAD3, MEK1/2, and ERK1/2. (Right) IB signals were quantified by densitometry, and ratios of phosphorylated to total SMAD3, MEK, and ERK1/2 (mean ± SEM; n = 4–6) are shown. *P < 0.05, **P < 0.01 vs. F/T. #P < 0.05, ##P < 0.01, ###P < 0.001 vs. FC/T. (B) Immunohistochemistry. (Left) p-ERK1/2 in cardiac vessels and endocardium (dark brown stain, arrows). (Right) Percentage of total area stained was quantified (mean ± SEM; n = 4). ***P < 0.001 vs. F/T. ###P < 0.001 vs. FC/T. (C) Gene expression in hearts 1 wk after TAC. mRNA levels were determined by qRT-PCR (mean ± SEM; n = 3–4). *P < 0.05, **P < 0.01 vs. F/T or C/T. ##P < 0.01 vs. FC/T.
Noncanonical TGF-β signaling plays a critical role in the pathogenesis of aortic dilatation in fibrillin-mutant mice (28). We therefore treated FC mice with U0126, an inhibitor of MEK–ERK signaling, by i.p. injection (7.5 mg/kg) 3 d before, immediately after, and 3 and 6 d after TAC. Echocardiography demonstrated a protective effect of U0126 on LVESD, EF, FS, and LV mass (Fig. 8A). In contrast, U0126 did not alter the modest remodeling observed in WT (F/T) mice. Inhibition of ERK phosphorylation was confirmed by IB (Fig. 8B) and immunohistochemical (Fig. 8C) assays. Immunofluorescence revealed ERK activation specifically in vascular ECs of FC/T hearts (Fig. 8D). Decreased cardiac fibrosis, cardiomyocyte hypertrophy, and caspase activation as well as increased EC density were observed in FC/T hearts treated with U0126 (Fig. 8E). Treatment of FC/T mice with NAb (Fig. 4F) or U0126 (Fig. 4G) also blocked ERK activation in the aorta and prevented aortic root dilatation (Fig. 4D).
Effect of U0126 on cardiac structure and function. (A) Echocardiographic data (mean ± SEM; n = 4–8) before (Basal) or 1 wk after (1W) TAC in F and FC mice treated with U0126 or vehicle. *P < 0.05, **P < 0.01 vs. FC/T. (B) Heart lysates were analyzed for ERK phosphorylation. (Upper) IB assays. (Lower) Densitometric analysis (mean ± SEM; n = 4–8). #P < 0.05, **P < 0.01 vs. F/T; ##P < 0.01 vs. FC/T. (C) ERK activation in heart sections from FC mice subjected to TAC and treated with vehicle (FC/T) or U0126 (FC/T+U). (Left) Immunohistochemistry for p-ERK (brown staining; arrows). (Right) Analysis of stained area as a percentage of total area (mean ± SEM; n = 4–8). **P < 0.01 vs. FC/T. (D) Colocalization of p-ERK and isolectin staining to cardiac vascular ECs by fluorescence microscopy. (E) Sections from hearts of mice treated with vehicle (FC/T) or U0126 (FC/T+U) were stained with Masson trichrome (fibrosis), WGA (myocyte area), isolectin (EC), or anti-activated caspase 3 Ab (apoptosis) and were subjected to image analysis. Mean ± SEM data (n = 4–6) are shown. *P < 0.05 vs. FC/T.
Our data indicate that excess MEK–ERK activation in ECs mediates aortic dilatation and cardiac decompensation in FC/T mice. The high percentage of gene deletion in heart (and lung) suggests that HIF-1α loss of function in cardiac ECs was very efficient, although the lack of a HIF-1α Ab suitable for immunohistochemistry of mouse tissues precluded a more definitive assessment. Although Hif1a gene deletion was observed in 60% of cardiomyocytes, the deletion in ECs is likely to have played a particularly important pathogenic role, based on the observation of increased ERK phosphorylation primarily in ECs and on the dramatic rescue of cardiac function and aortic integrity with NAb or U0126 treatment in FC/T mice. In contrast, NAb treatment failed to protect against the less severe pathological remodeling that occurs following TAC in WT mice, in which TGF-β signaling in cardiomyocytes plays a pivotal role (24). The much less severe post-TAC phenotype of mice with cardiomyocyte-specific knockout of HIF-1α (20, 21) provides further support for the conclusion that HIF-1 loss of function in ECs resulted in cardiac decompensation after TAC.
The key pathological change in FC/T hearts appeared to be a marked reduction of capillary density leading to increased myocardial hypoxia and apoptosis. The effect of HIF-1α deficiency on capillary density played a more critical pathogenic role than the effect on cardiac fibrosis or cardiomyocyte hypertrophy, because treatment with U0126, but not with NAb, corrected the excess fibrosis and hypertrophy that was induced by TAC, whereas both U0126 and NAb prevented the TAC-induced vascular and contractile abnormalities in FC/T mice. In mouse models in which noncanonical TGF-β signaling in cardiomyocytes was implicated in late-onset pathological remodeling, the loss of ECs also appeared to play a role in the loss of contractile function (24). Finally, TAC also induced aortic root dilatation in FC/T mice, another phenotype that is caused by enhanced TGF-β signaling in the vasculature (26, 27).
The failure of NAb and the success of U0126 in inhibiting cardiac fibrosis and cardiomyocyte hypertrophy likely results from the inability of NAb to penetrate into the myocardium (24), whereas the low-molecular-weight compound U0126 is not subject to this limitation. These results suggest that the increased cardiac fibrosis and hypertrophy observed in FC/T mice may be caused by HIF-1α loss of function in cardiomyocytes. However, the ability of NAb to rescue cardiac function in FC/T mice suggests that HIF-1–dependent suppression of TGF-β signaling in ECs is critical for vascular adaptation to pressure overload, and this adaptation in turn is required for maintenance of contractile function. These findings contrast with a report indicating that HIF-1 and TGF-β act cooperatively in cultured ECs subjected to hypoxia (31). It is possible that HIF-1 loss of function in Tie2+ lineage inflammatory cells also contributes to the loss of microvasculature after TAC.
Age-dependent impairment of HIF-1α induction leads to diminished vascular responses to limb ischemia (32) and wound healing (33). Thus, aging may impair the ability of HIF-1 to suppress TGF-β signaling, thereby increasing the risk of heart failure in elderly patients with hypertension. Further studies are needed to test this hypothesis and to determine whether pharmacological activation of HIF-1 using prolyl hydroxylase inhibitors may protect against heart failure. However, cardiomyocyte-specific overexpression of HIF-1α causes a progressive cardiomyopathy (34), indicating the need to regulate its expression tightly to maintain O2 homeostasis and cardiac function. Finally, the inhibitory effect of digoxin on LV function after TAC suggests that this drug may have countertherapeutic effects that explain why it does not increase survival in patients with heart failure (35). In contrast, digoxin blocks pathological pulmonary vascular remodeling and reduces right ventricular pressure in mice subjected to chronic hypoxia (36).
Hif1af/f mice (37) and Tie2-Cre mice (23) were mated to generate Hif1af/f;Tie2-Cre mice, which were maintained by mating with Hif1af/f mice. Animal studies were performed according to protocols approved by The Johns Hopkins University Animal Care and Use Committee in accordance with the National Institutes of Health Guide for Care and Use of Laboratory Animals.
Male mice (age 9–12 wk, 25–30 g) were subjected to TAC as described (24). Mice were anesthetized and placed supine on a 37 °C heating pad. An incision was made at the suprasternal notch. A 7.0 silk suture was passed under the aorta. A 27-gauge needle was placed next to the aortic arch, the suture was tied around the needle and the aorta, and the needle was removed. The chest was closed, and the animal was allowed to recover. Sham-operated mice underwent the same procedure but without constriction.
Mouse hearts were imaged in M-mode using an Acuson Sequoia C256 with a 13-MHz transducer (Siemens) as described (38). Using a Vevo 2100 with a 30-MHz transducer (VisualSonics), the proximal aorta was imaged in B-mode (parasternal long-axis view) Three measurements of maximal internal dimension at the sinus of Valsalva and proximal ascending aorta were made from distinct captured images and were averaged (39). Color Doppler mode was used to rule out aortic insufficiency. Data were obtained and analyzed by operators blinded to genotype and treatment groups.
Genomic DNA was extracted from mouse tail biopsies, heart tissues, and isolated cardiomyocytes using the DNeasy tissue kit (Qiagen) following the manufacturer's protocol. PCR was performed using the following primers: Hif1a deletion, 5′-CGGCGAAGCAAGAGTCTG-3′ and 5′-ATAACTGATGGTGAGCCTCATAAC-3′; Hif1a control, 5′-GTGGTATTATTCAGCACGACTTG-3′ and 5′-GGAGCCAGCAGAGTGAGAG-3′; and Tie2-Cre, 5′-CGAACCATTCAGACTGTG-3′ and 5′-CCCTGTGCTCAGACAGAAATGAGA-3′.
Tissues were fixed in 10% (vol/vol) formalin. Transverse sections of heart and aorta from TAC- and sham-operated mice were stained with Masson trichrome, Alexa Fluor 568-labeled wheat germ agglutinin, Alexa Fluor 488-labeled isolectin B4 (Invitrogen), or Ab against phosphorylated ERK1/2 or cleaved caspase3 (Cell Signaling).
Whole-tissue lysates were prepared from the isolated LV. IB assays were performed with antibodies that specifically recognized phosphorylated SMAD2, SMAD3, ERK1/2 (Thr202/Tyr204), and MEK1/2 (Ser217/Ser221) and antibodies that recognized total SMAD2, SMAD3, ERK, MEK, and β-actin (Cell Signaling Technology).
Total RNA was extracted from the LV using TRIzol (Invitrogen). cDNA synthesis, using an iScript kit (BioRad), and real-time qPCR, using Maxima SYBR Green Master Mix (Fermentas) and iCycler Real-Time PCR Detection System (BioRad), were performed, and the expression of each mRNA relative to 18S rRNA was calculated based on the threshold cycle method (40). A PCR-based mouse TGF-β signaling microarray (SA Biosciences) was used according to the manufacturer's protocol.
Pimonidazole (Hypoxyprobe-1) was administered by i.p. injection (80 mg/kg), and 90 min later the heart was harvested, placed in 10% formalin, and processed for immunohistochemistry using an anti-pimonidazole Ab (Natural Pharmacia International).
TGF-β NAb (R&D Systems) was administered by i.p. injection (10 mg/kg) immediately after TAC. U0126 (LC Laboratories) was administered by i.p. injection at a dose of 7.5 mg/kg 3 d before, immediately after, and 3 and 6 d after TAC.
Data are expressed as mean ± SEM. Differences were analyzed by ANOVA or t test.
This work was supported by National Institutes of Health Grant P01-HL65608, Contracts N01-HV28180 and HHS-N268201000032C, and The Johns Hopkins Institute for Cell Engineering. G.L.S. is the C. Michael Armstrong Professor at The Johns Hopkins University School of Medicine.
Author contributions: H.W. and G.L.S. designed research; H.W., D.B., N.K., D.X., J.C., K.F.-T., R.R., and S.C. performed research; J.W.H., H.C.D., and K.L.G. contributed new reagents/analytic tools; H.W., D.B., R.R., C.S., D.A.K., and G.L.S. analyzed data; and H.W. and G.L.S. wrote the paper.
The authors declare no conflict of interest.
See Author Summary on page 5160 (volume 109, number 14).
