Analyses of several mouse models imply that the phosphodiesterase 5 (PDE5) inhibitor sildenafil (SIL), via increasing cGMP, affords protection against angiotensin II (Ang II)–stimulated cardiac remodeling. However, it is unclear which cell types are involved in these beneficial effects, because Ang II may exert its adverse effects by modulating multiple renovascular and cardiac functions via Ang II type 1 receptors (AT1Rs). To test the hypothesis that SIL/cGMP inhibit cardiac stress provoked by amplified Ang II/AT1R directly in cardiomyocytes (CMs), we studied transgenic mice with CM-specific overexpression of the AT1R under the control of the α-myosin heavy chain promoter (αMHC-AT1Rtg/+). The extent of cardiac growth was assessed in the absence or presence of SIL and defined by referring changes in heart weight to body weight or tibia length. Hypertrophic marker genes, extracellular matrix–regulating factors, and expression patterns of fibrosis markers were examined in αMHC-AT1Rtg/+ ventricles (with or without SIL) and corroborated by investigating different components of the natriuretic peptide/PDE5/cGMP pathway as well as cardiac functions. cGMP levels in heart lysates and intact CMs were measured by competitive immunoassays and Förster resonance energy transfer. We found higher cardiac and CM cGMP levels and upregulation of the cGMP-dependent protein kinase type I with AT1R overexpression. However, even a prolonged SIL treatment regimen did not limit the progressive CM growth, fibrosis, or decline in cardiac functions in the αMHC-AT1Rtg/+ model, suggesting that SIL does not interfere with the pathogenic actions of amplified AT1R signaling in CMs. Hence, the cardiac/noncardiac cells involved in the cross-talk between SIL-sensitive PDE activity and Ang II/AT1R still need to be identified.
Evidence from animal and clinical studies with angiotensin-converting enzyme inhibitors and antagonists of angiotensin II (Ang II) type 1 receptors (AT1Rs) identified the Ang II/AT1 receptor pathway as a major cause of cardiac damage (de Gasparo et al., 2000; López-Sendón et al., 2004; Higuchi et al., 2007; McMurray et al., 2012; Elliott et al., 2014). Kidney cross-transplantation experiments in gene-targeted mice suggest that selective ablation of renal AT1Rs prevents adverse cardiac remodeling, whereas AT1Rs in the heart are not involved in the Ang II–stimulated stress response, i.e., pathologic growth of cardiomyocytes (CMs) and fibrosis (Crowley et al., 2006). In contrast, different rodent models that overexpress the AT1R in CMs reportedly developed hypertrophy, fibrosis, cardiac dysfunction (Paradis et al., 2000; Hoffmann et al., 2001; Ainscough et al., 2009), or premature death (Hein et al., 1997). Moreover, direct growth effects of Ang II on cardiac fibroblasts and smooth muscle cells (SMCs) (Griendling et al., 1997; Leask, 2010) and a strong interplay between the different kinds of cardiac cells are well documented (Fredj et al., 2005; Kakkar and Lee, 2010; Takeda et al., 2010; Tirziu et al., 2010); hence, interaction of Ang II with responsive receptors at different cardiovascular and renal locations may be important to elicit its prohypertrophic and profibrotic effects.
The natriuretic peptide (NP)/cGMP pathway has been shown to play an important cardioprotective role in the cardiac stress response stimulated by various factors, including Ang II (Li et al., 2002; Nishikimi et al., 2006; Kilić et al., 2007; Hofmann et al., 2009; Kinoshita et al., 2010; Bice et al., 2014). Interestingly, a combined strategy to prevent the degradation of several vasoactive peptides, including the NPs and AT1R blockade, has recently given positive results in a large phase III clinical trial in patients with heart failure (McMurray et al., 2014). Among other modes of action, neprilysin inhibition is expected to increase the local bioavailability of endogenous NPs, pointing to a role of the cGMP pathway in this setting. Along these lines, inhibition of the cGMP-hydrolyzing enzyme phosphodiesterase 5 (PDE5) using sildenafil (SIL) was shown to prevent or even reverse pressure overload–induced remodeling by transverse aortic constriction (TAC) in mice (Takimoto et al., 2005), showing beneficial effects in several other cardiac disease conditions such as ischemia/reperfusion injury and cardiotoxicity from doxorubicin (Kass, 2012). Based on these findings, it has been suggested that favorable effects of SIL require cGMP and cGMP-dependent protein kinase type I (cGKI) in the CM (Takimoto et al., 2005, 2009; Koitabashi et al., 2010; Nishida et al., 2010), with the latter one being the major target of NP/cGMP signaling in the cardiovascular system (Hofmann et al., 2009; Takimoto, 2012). Indeed, multiple targets downstream of NP or SIL and cGMP/cGKI were suggested to inhibit adverse cardiac hypertrophy in response to Ang II, elevated afterload, and other causes (Tokudome et al., 2008; Takimoto et al., 2009; Klaiber et al., 2010; Koitabashi et al., 2010; Nishida et al., 2010; Frantz et al., 2013). However, analysis of a murine model that lacks the cGKIα and -Iβ isoenzymes in all non-SMCs, including CMs (cGKIβ rescue mice) has challenged some of these previous findings (Lukowski et al., 2010). Cardiac growth responses to TAC or isoproterenol in cGKIβ rescue mice were normal, suggesting either that non-SMC cGKI is not required to inhibit pressure overload– and β-adrenergic receptor–triggered remodeling or cGMP/cGKI are not appropriately activated under these experimental conditions. Therefore, we previously applied a hypertensive dose of Ang II and confirmed that antifibrotic effects of SIL (Westermann et al., 2012) were indeed related to non-SMC cGKI (Patrucco et al., 2014). The following points regarding the cellular targets of SIL/cGMP/cGKI and Ang II are still open (Lukowski et al., 2014): 1) The vasoactive neurohormone Ang II may provoke cardiac stress by changes in hemodynamics, i.e., an increase in afterload resulting from high blood pressure, and from direct effects on non-CM cells. Therefore, changes in the fibrotic response upon Ang II and SIL coadministration cannot easily be tracked down to a definite type of cell. 2) SIL by itself may signal via different PDE5-expressing cells in the whole-animal system. 3) SIL may affect cardiac cGMP turnover independently of PDE5, e.g., by an effect on PDE1C (Lukowski et al., 2010, 2014).
To this end, we studied mice that carry a CM-specific overexpression of the human AT1R under the control of the α-myosin heavy chain promoter (αMHC-AT1Rtg/+) (Paradis et al., 2000) to discriminate the cross-talk between SIL/cGMP and the Ang II/AT1R during adverse cardiac remodeling. Reportedly, AT1R transgenic mice develop a severe cardiac phenotype, i.e., progressive primary CM hypertrophy with secondary interstitial fibrosis even in the absence of Ang II (Yasuda et al., 2012) and high blood pressure (Paradis et al., 2000; Karnik and Unal, 2012). We tested whether SIL at a previously confirmed antifibrotic dose resulting in a free plasma concentration of ∼70 nM (Adamo et al., 2010; Patrucco et al., 2014) would interfere with amplified in vivo AT1R signaling in the αMHC-AT1Rtg/+ mouse model. Our data indicate that SIL does not prevent cardiac remodeling induced by CM-specific AT1R overexpression, suggesting that other cell types are involved in the potentially favorable effects of this drug.
Materials and Methods
Animal Welfare, Ethical Statements, and Genetic Background of the Experimental Mice.
Experimental mice were bred and maintained at the animal facility of the Institute of Pharmacy, Department of Pharmacology, Toxicology and Clinical Pharmacy, University of Tübingen. All procedures were performed with permission of the local authorities and conducted in accordance with the German legislation on the protection of animals. The mice were kept in temperature- and humidity-controlled cabinets, in cages with wood-chip bedding on a standard 12-hour light/dark cycle with ad libitum access to food (Altromin, Lage, Germany) and water. For experiments, mice with CM-restricted overexpression of the AT1R (genotype, αMHC-AT1Rtg/+) were compared with age- and litter-matched wild-type (WT) mice on a C57BL/6 genetic background (Paradis et al., 2000). Genotyping was performed by a previously established polymerase chain reaction (PCR) protocol using mouse tail genomic DNA and a set of two primers (AT1R forward: 5′-ACC CTT ACC CCA CAT AGA C-3′; AT1R reverse: 5′-ACC ATC TTC AGT AGA AGA GTT G-3′) that amplified the transgenic AT1 allele (400 bp). Experimental mice were studied irrespective of their gender at an age of 8–24 weeks. For organ removal, animals were sacrificed by CO2 inhalation. All studies were performed in accordance with the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guidelines for reporting experiments involving animals.
Long-Term SIL Application Protocol.
Previously assessed cardiac phenotypes of AT1R transgenic mice upon 2 months of losartan treatment (30 mg/kg per day) revealed beneficial effects on various parameters of the adverse remodeling process (Paradis et al., 2000). Using an analogous protocol, 60-day-old AT1R transgenic mice and control animals from the same litters received SIL (400 mg/l) in acidified drinking water (pH 4.5) for 2 months. This dose of SIL was shown to be antifibrotic in an Ang II–stimulated model of cardiac hypertrophy (Patrucco et al., 2014) and yielded an average plasma concentration of 70 nM (Adamo et al., 2010). This concentration is well in the range to inhibit PDE5 (IC50, 10 nM).
Noninvasive echocardiography was performed using an existing protocol with minor variations (Lukowski et al., 2010). In brief, high-resolution images were acquired by the use of an imaging system equipped with a 30-MHz probe (Vevo 2100; VisualSonics, Toronto, ON, Canada). Throughout the procedure, mice were anesthetized by a continuous oxygen/isoflurane (2%) inhalation. The chest was gently shaved, and the area was cleaned with alcohol to minimize ultrasound reduction. For image acquisition, mice were placed on a heated surgical platform in a dorsal position. From good-quality two-dimensional images obtained in the parasternal short-axis view, the fractional shortening and ejection fraction (EF) were assessed in M-mode recordings. Hemodynamic functions of the heart were calculated in three different positions from a total of nine independent measurements per mouse.
Isolation of Adult CMs from WT and αMHC-AT1Rtg/+ Hearts.
Adult CMs were isolated as described previously by a modified protocol of the Alliance for Cellular Signaling (procedure protocol PP00000125) (Lukowski et al., 2010; Soltysinska et al., 2014). After retrograde enzyme perfusion via the aorta, the hearts were minced into small pieces and myocytes were liberated by gently applying mechanical turbulence. For the quantification of the size by planimetry using AxioVision Microscope Software 4.8 (Zeiss, Jena, Germany), cells were plated in six-well format and immediately photographed.
mRNA Expression Analysis Using Real-Time Quantitative PCR.
Total RNA was purified from isolated heart ventricles with 500 μl peqGOLD RNA pure reagent (PEQLab Biotechnologie, Erlangen, Germany) according to the manufacturer's protocol. The RNA concentration was assessed by UV photometry in a NanoPhotometer (Implen, München, Germany) and diluted in diethyl pyrocarbonate–treated ddH2O to a final concentration of 0.1 μg/μl. The RNA (500 ng) was used as a template to synthesize cDNA using the iScript cDNA Synthesis Kit (Bio-Rad Laboratories, Hercules, CA). A 1:10 dilution of the cDNA was amplified using an iQ SYBR Green Supermix (Bio-Rad) in an Opticon real-time quantitative PCR detection system (MJ Research, Watertown, MA) with specific primers. As consistent markers of heart hypertrophy or fibrosis, the expression levels of atrial natriuretic peptide (ANP), brain natriuretic peptide (BNP), sarcoplasmic/endoplasmic Ca2+-ATPase 2 (SERCA2), myosin heavy chain α-isoform (αMHC), connective tissue growth factor (CTGF), α-smooth muscle actin (αSMA), collagen type 1 α-1 chain (Col1A1), collagen type 1 α-2 chain (Col1A2), matrix metalloproteinase 9 (MMP9), and tissue inhibitor of metalloproteinase 1 (TIMP1) were determined and related to hypoxanthine phosphoribosyltransferase 1 (HPRT1) as an internal control for the quantitative PCR reaction. The primer sequences designated were ANP forward: 5′-TGT ACA GTG CGG TGT CCA AC-3′; ANP reverse: 5′-GGG GCA TGA CCT CAT CTT CT-3′; BNP forward: 5′-TCC TCT GGG AAG TCC TAG CC-3′; BNP reverse: 5′-GCC ATT TCC TCC GAC TTT TC-3′; SERCA2 forward: 5′-CTG CTG CAT GGT GGT TCA T-3′; SERCA2 reverse: 5′-TCC ACT CCA TCG AAG TCT GG-3′; αMHC forward: 5′-TGC TGA CAG ATC GGG AGA AT-3′; αMHC reverse: 5′-TGC TGG CAA AGT ACT GGA TG-3′; CTGF forward: 5′-AGG GCC TCT TCT GCG ATT TC-3′; CTGF reverse: 5′-TAC ACC GAC CCA CCG AAG AC-3′; αSMA forward: 5′-AGA GGC ACC ACT GAA CCC TA-3′; αSMA reverse: 5′-GCA TAG AGG GAC AGC ACA GC-3′; Col1A1 forward: 5′-GAG GAA ACT TTG CTT CCC AGA-3′; Col1A1 reverse: 5′-ACC ACG AGG ACC AGA AGG AC-3′; Col1A2 forward: 5′-GTC TGT TGG AGC TGC TGG CCC AT-3′; Col1A2 reverse: 5′-GCA GCA CCA GGG AAG CCA GTC AT-3′; MMP9 forward: 5′-GGA GTT CTC TGG TGT GCC CT-3′; MMP9 reverse: 5′-ACA CGC CAG AAG AAT TTG CCA-3′; TIMP1 forward: 5′-CCA CCC ACA GAC AGC CTT CT-3′; TIMP1 reverse: 5′-CGC TGG TAT AAG GTG GTC TCG-3′; HPRT1 forward: 5′-CAT TAT GCC GAG GAT TTG GA-3′; HPRT1 reverse: 5′-CCT TCA TGA CAT CTC GAG CA-3′. Additionally, we assessed the overexpression of the human AT1R and the expression of PDE5 using the following primer pairs: AT1R forward: 5′-ACA GTA TCA TCT TTG TGG TGG GA-3′; AT1R reverse: 5′-GGC CAC AGT CTT CAG CTT CA-3′; PDE5 forward: 5′-GGA ACA CCA TCA TTT TGA CCA GT-3′, PDE5 reverse: 5′-AGA GGC CAC TGA GAA TCT GGT-3′.
Immunoblotting of Cardiac Proteins.
Western blot analysis was performed with protein extracts obtained from αMHC-AT1Rtg/+ and control mice. Upon dissection and organ removal, the residual blood in the heart chambers and vessels was thoroughly removed via a retrograde injection of ice-cold phosphate-buffered saline (PBS) using a 1-ml syringe combined with a 25-gauge needle. Atria were immediately removed, and ventricles were frozen on liquid nitrogen and stored at −80°C until homogenization was carried out for extraction of the total protein using lysis buffer (20 mM Tris-HCl, pH 8.3; 0.67% SDS; 238 mM β-mercaptoethanol; and 0.2 mM phenylmethanesulfonylfluoride). Proteins (50 μg/lane) were separated by their molecular weight using denaturing 10% SDS-PAGE. Immunodetection was performed using the cGKI common antibody (Methner et al., 2013) (dilution 1:250) and a primary antibody specific for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (dilution 1:1000; Cell Signaling Technology, Danvers, MA). To identify the primary antibody-protein complexes, a Cy5-coupled secondary anti-rabbit antibody (ECL Plex; 1:2500 dilution; GE Healthcare, Little Chalfont, UK) was used. Fluorophores were detected using the Ettan DIGE System (GE Healthcare).
Sirius Red and H&E Staining.
For histology, hearts were dissected, rinsed in PBS, and fixed for 4 hours in 2% paraformaldehyde in PBS. Cryoprotection was accomplished by a series of sucrose solutions with increasing concentrations (5%, 10%, and 20%) in PBS. Neg-50 (Thermo Fisher Scientific, Waltham, MA)–embedded organs were sectioned at 8-μm intervals (Microm HM 560 Cryostat; Thermo Fisher Scientific). H&E staining was performed using standard procedures (Lukowski et al., 2010). The distribution and amount of extracellular collagen fibers were identified using a sirius red staining solution (Direct Red 80 in saturated picric acid) on sequential cardiac sections that were pretreated for 24 hours in Bouin's fixative (5% acetic acid and 9% formaldehyde in saturated aqueous picric acid).
Measurement of CM Size and Fibrosis.
CM size was assessed in photomicrographs of H&E-stained cross-sections using AxioVision Microscope Software 4.8. Cross-sectional areas of ∼300–500 randomly chosen cells with a centrally located nucleus from n = 9–12 hearts per genotype and treatment group were determined. For determining the amount of fibrosis, sirius red–stained sections were scanned using MIRAX Desk Scanner and illustrated with MIRAX Viewer Software (Zeiss). The GSA Image Analyzer software package (Gesellschaft für Softwareentwicklung und Analytik, Rostock, Germany) was used for the quantification of red-stained myocardial tissue and total ventricular areas.
cGMP Determination in the Heart and Aorta.
For cGMP measurements, hearts from transgenic AT1Rtg/+ mice that received either saline or SIL (400 mg/l) for 7 days were removed and flushed with ice-cold PBS. Further cGMP purification was performed as described earlier (Lukowski et al., 2008). In brief, the hearts were weighed before the cyclic nucleotides were extracted by homogenization of the tissue using 1 ml ice-cold 100% EtOH. Resultant homogenates were centrifuged for 7 minutes at 13,000 rpm. The supernatant containing the cyclic nucleotides was concentrated by evaporating the alcohol in a speed vac for 3–4 hours at room temperature. The resulting pellet was resuspended in 115 μl enzyme immunoassay buffer. cGMP concentration in the samples was determined according to the manufacturer’s recommendations of the enzyme immunoassay kit (Cayman Chemical, Ann Arbor, MI).
cGMP-PDE Activity Assay in Heart Cytosolic Preparations.
Whole hearts were homogenized in cold lysis buffer containing 50 mM NaCl; 1 mM EDTA; 2 mM dithiothreitol; 50 mM triethanolamine HCl, pH 7.4; and protease inhibitors (1 μM pepstatin A, 1 mM benzamidine, 1 mM phenylmethanesulfonylfluoride, and 50 μM leupeptin) using a glass-glass homogenizer. After centrifugation (30 minutes, 4°C, 16,000g), PDE activity in the supernatant was measured by the conversion of [32P]cGMP (synthesized from [α-32P]GTP using purified guanylyl cyclase) to guanosine and [32P]phosphate in the presence of alkaline phosphatase at 37°C for 10 minutes. Reaction mixtures contained [32P]cGMP (∼50,000 cpm), 1 μM cGMP, 12 mM MgCl2, 3 mM dithiothreitol, 0.5 mg/ml bovine serum albumin, 1 U alkaline phosphatase, and 50 mM triethanolamine HCl, pH 7.4, in a total volume of 0.1 ml. Reactions were stopped by the addition of 900 μl ice-cold charcoal suspension (30% activated charcoal in 50 mM KH2PO4, pH 2.3). After pelleting the charcoal by centrifugation, [32P]phosphate was measured in the supernatant.
Förster Resonance Energy Transfer–Based cGMP Measurements in Intact CMs.
To analyze CM-specific changes in cGMP levels, we crossed αMHC-AT1Rtg/+ animals with red cGES-DE5 transgenic cGMP sensor mice (Götz et al., 2014). Adult CMs were isolated from double-transgenic animals versus littermate controls expressing the sensor without AT1R overexpression and subjected to cGMP–Förster resonance energy transfer (FRET) measurements under stimulation with C-type natriuretic peptide (CNP) (1 μM) and 3-isobutyl-1-methylxanthine (IBMX) (100 μM) as previously described (Götz et al., 2014).
Data Analysis and Statistical Procedures.
All values are expressed as mean ± S.E.M. Statistical significance was estimated by using Student's t test for unpaired observations with Excel software package (Figs. 1D, 2, C–E, and 3, H–J; Supplemental Figs. 1, C–F, H, and I, and 2, A and B). One-way analysis of variance was used for comparison of more than two groups (Figs. 1B, 3, A–F, 4, B–F, and 5, B–D; Supplemental Fig. 1A), followed by a Tukey-Kramer multiple-comparisons post hoc test using GraphPad InStat 3.0 (GraphPad Software, La Jolla, CA). Statistical differences between genotypes and treatment conditions are indicated as P values. Data were considered to be significant at */#/§P < 0.05, **/##/§§P < 0.01, and ***/###/§§§P < 0.001.
Progressive Heart Hypertrophy and Fibrosis in the αMHC-AT1Rtg/+ Mouse Model.
To revalidate the progressive cardiac stress phenotype of αMHC-AT1Rtg/+ animals, cohorts of transgenic mice and WT age-matched littermates were sacrificed at different time points after birth. Wet weights of the dissected hearts [heart weight (HW)] were measured for every mouse and normalized to either the respective total body weight (BW) or the tibia length (TL) to determine HW/BW and HW/TL ratios (Supplemental Fig. 1A and data not shown). If anything, we observed a decline in both ratios in WT animals with time, whereas the quantification of the αMHC-AT1Rtg/+ ratios defined at 60, 120, and 160 days after birth showed a time-dependent increase of the HW/BW ratio from 5.55 ± 0.05 to 6.04 ± 0.19 and 6.48 ± 0.33 mg/g, respectively (Supplemental Fig. 1A). Prominent increases in CM cross-sectional areas and in the dimensions of primary adult CMs as major causes for the abnormal growth of the hearts could be confirmed (Supplemental Fig. 1, C and D) in presence of AT1R overexpression by more than 150-fold (Supplemental Fig. 1E), specifically in CMs (Supplemental Fig. 1F) (Paradis et al., 2000). Importantly, neither total BW nor any other body proportions, including TLs or naso-anal sizes, were different between αMHC-AT1Rtg/+ and WT litters (data not shown). In addition, the histopathology in the αMHC-AT1Rtg/+ model was characterized by an extensive accumulation of extracellular matrix (ECM) components (Supplemental Fig. 1G) and high transcript levels of genes associated with myocardial fibrosis, which both likely derived from activation of fibroblast-like cells secondary to chronic stimulation of AT1R signal transduction in CMs (Supplemental Fig. 1, G–I). These results are in good agreement with previous reports evaluating pathophysiological properties of the cardiac remodeling process in the αMHC-AT1Rtg/+ mouse line (Paradis et al., 2000; Yasuda et al., 2012). As a potential cause underlying the cardiac phenotype of αMHC-AT1Rtg/+ mice, we considered a change in the relative balance between pro- and antihypertrophic factors in the CMs.
Cardiac cGMP/cGKI Are Stimulated upon AT1R Overexpression in CMs.
We examined the potentially antihypertrophic and antifibrotic roles of cGMP signaling for the (Ang II)/AT1R-provoked defects in the CM (Booz, 2005), as intensive cross-talk between these pathways in different settings of cardiac stresses has been reported (Li et al., 2002; Masuyama et al., 2006; Tokudome et al., 2008; Klaiber et al., 2010; Mokni et al., 2010; Frantz et al., 2013; Patrucco et al., 2014).
Using a genetically encoded cGMP FRET-based sensor in intact adult CMs (Götz et al., 2014), we observed that CNP induced an ∼2-fold-higher cGMP elevation in AT1Rtg/+ cells (isolated from 120-day-old double-transgenic mice) compared with control mice that carried only the FRET-based sensor (Fig. 1A). However, IBMX following CNP treatment attenuated this difference in cGMP between the two genotypes (Fig. 1B). Interestingly, IBMX alone increased the basal cGMP level only in control CMs, with a much less pronounced effect in double-transgenic mouse cells, suggesting a reduction of PDE-mediated effects on cGMP in AT1Rtg/+ CMs under these experimental conditions (Fig. 1, C and D). By using validated cGKI common specific antibodies (Geiselhöringer et al., 2004), we identified a higher abundance of cardiac cGKI in protein lysates derived from 120- and 160-day-old αMHC-AT1Rtg/+ hearts as compared with WT samples (Fig. 2, A–D). In addition to cGKI, basal cGMP concentrations measured in αMHC-AT1Rtg/+ whole-heart lysates were increased (Fig. 2E), confirming activation of the cGMP/cGKI pathway in a setting of amplified Ang II/AT1R in CMs.
Based on these findings, we reasoned that long-term stimulation of cGMP signaling in CMs may ameliorate the cardiac phenotype, i.e., detrimental changes in heart dimension and structure of αMHC-AT1Rtg/+ mice. To test whether PDE5-sensitive cGMP pools are involved in the protection against amplified AT1R signaling, SIL was administered ad libitum via the drinking water at a concentration of 400 mg/l. We treated αMHC-AT1Rtg/+ and WT mice for 60 consecutive days and evaluated the effects of SIL on different histologic and functional parameters in comparison with two cohorts of age-matched littermates from both genotypes that served as untreated controls (Fig. 2, F and G). An analogous long-term SIL regimen previously resulted in an average plasma level of ∼70 nM (Adamo et al., 2010); hence, the drug concentration should be well above the IC50 value to inhibit the PDE5 and ∼7-fold higher than the levels required for the inhibition of TAC-induced cardiac remodeling (Takimoto et al., 2005).
SIL Does Not Counteract the Progressive Cardiac Stress Phenotype in αMHC-AT1Rtg/+ Mice.
Long-term SIL administration affected neither the BW gain nor the survival rates of αMHC-AT1Rtg/+ or WT mice (data not shown). Cardiac remodeling was evaluated at the end of the treatment period in 120-day-old WT and αMHC-AT1Rtg/+ animals (Fig. 3). Quantifications of gross parameters such as the HW/BW and HW/TL ratios did not reveal any beneficial effects of SIL on the extent of heart growth in the αMHC-AT1Rtg/+ model (Fig. 3, A and B). To exclude the possibility that the AT1Rtg/+ overexpression induced early alterations in myocardial structure that were not reversible at 60 days of age (or later), an additional group of αMHC-AT1Rtg/+ animals received SIL for 80 days shortly after weaning, i.e., at 40 days of age. As compared with the respective 60-day SIL treatment group, the prolonged protocol did not show a significant effect on the HW/BW and HW/TL ratios at 120 days of age (Supplemental Fig. 2, A and B). We further evaluated the expression levels of hypertrophic marker genes such as ANP, BNP, αMHC, and SERCA2 (Fig. 3, C–F). As compared with WT hearts, the mRNA levels for ANP and BNP in αMHC-AT1Rtg/+ hearts were ∼4- to 10-fold higher under both control and SIL-treated conditions (Fig. 3, C and D), providing support for the NP/cGMP/cGKI pathway being activated in response to amplified AT1R signaling in CMs. However, neither the mRNA level of the hypertrophic markers ANP and BNP (Fig. 3, C and D) nor the amount of αMHC and SERCA2 mRNA (Fig. 3, E and F) nor the expression level of the human AT1R itself (data not shown) in the αMHC-AT1Rtg/+ model were sensitive to the long-term SIL protocol. If anything, the significant difference in SERCA2 mRNA levels between untreated WT and αMHC-AT1Rtg/+ mice was less pronounced in the presence of SIL (Fig. 3F). Our quantifications of different hypertrophy markers were in excellent agreement with histomorphologic examinations of the CM dimensions (Fig. 3, G and H). Importantly, the mean size of the CMs from αMHC-AT1Rtg/+ mice that either received SIL or remained untreated was identical (Fig. 3, G and H). Because CM-specific defects, i.e., the maladaptive hypertrophic growth in the αMHC-AT1Rtg/+ mouse model, were not inhibited by SIL, we conclude that the rise in both global cardiac and aortic cGMP concentrations as measured at day 7 of the SIL treatment (Fig. 3, I and J) had no impact on the CMs.
Several recent reports suggested an antifibrotic mode of action for SIL and cGMP/cGKI on the cardiac stress response caused by Ang II infusions (Westermann et al., 2012; Patrucco et al., 2014). We tested the effect of long-term SIL on the fibrotic response in the absence of high Ang II, but with amplified AT1R signaling in CMs. We determined the amount of interstitial ECM proteins, i.e., collagens. Cardiac tissue samples from SIL-treated or -untreated αMHC-AT1Rtg/+ mice were examined from 120-day-old animals (Fig. 4, A and B). Histologically, pronounced fibrosis of the papillary muscle and myocardium was apparent in αMHC-AT1Rtg/+ hearts obtained from both control and SIL-treated groups. As expected, the amount of ECM proteins and the levels of the fibrosis markers were in general lower in WT as compared with αMHC-AT1Rtg/+ heart sections (Fig. 4). To further assess the fibrotic response in the two αMHC-AT1Rtg/+ treatment groups, we quantified the transcription levels of several genes associated with ECM turnover and fibrosis, including Col1A1, Col1A2 (Fig. 4, C and D), MMP9, and TIMP1 (Fig. 4, E and F). In line with the histologic quantifications (Fig. 4B), SIL did not affect the transcription levels of the profibrotic genes Col1A1 and Col1A2, whereas the differences in TIMP1 and MMP9 mRNA between the SIL-treated WT and αMHC-AT1Rtg/+ hearts reached significance (compare § groups in Fig. 4, E and F). However, a similar tendency to different turnover rates of the ECM components was also observed in the untreated WT versus αMHC-AT1Rtg/+ groups (Fig. 4, C–F), confirming that SIL did not affect the total amount of fibrosis deposition upon overexpression of the human AT1R in CMs.
Amplified AT1R signaling in the CM reportedly caused a gradual decline in cardiac functions (Paradis et al., 2000; Hoffmann et al., 2001; Yasuda et al., 2012), whereas SIL afforded cardioprotection and improved heart pump functions in different settings of cardiac stress (Takimoto et al., 2005, 2009; Adamo et al., 2010; Zhang et al., 2010; Blanton et al., 2012; Westermann et al., 2012). Hence, we assessed a number of echocardiographic parameters in anesthetized 120-day-old WT and αMHC-AT1Rtg/+ mice that received either SIL or placebo for 60 days (Fig. 5, A, 1 and 2). In agreement with previous studies (Rivard et al., 2011; Yasuda et al., 2012), the contractility of the heart muscle and the cardiac EF were severely disturbed in αMHC-AT1Rtg/+ mice (Fig. 5, B and C), but these parameters were not altered by SIL (Fig. 5, B and C). Moreover, the cardiac PDE5 mRNA expression (Fig. 5D) remained stable over time and did not differ between the two genotypes. Due to this outcome, we reasoned that the inability of SIL to affect function and growth parameters of αMHC-AT1Rtg/+ hearts was not caused by an unexpected regulation of its primary drug target.
The cGMP-Hydrolyzing Activity of αMHC-AT1Rtg/+ Heart Lysates in Response to SIL.
SIL at concentrations that should specifically inhibit PDE5 (≤10 nM) had no effect on the cGMP-hydrolytic activity in WT or hypertrophic αMHC-AT1Rtg/+ heart lysates (Fig. 5, E and F). With excessive amounts of SIL (100–1000 nM), however, a moderate but reproducible inhibition of the cardiac cGMP-hydrolytic activity was observed in both genotypes, indicating inhibition of cardiac cGMP-PDEs different from PDE5 (Lukowski et al., 2014). When total heart PDE activity was determined in the presence of Ca2+/calmodulin, we observed a similar left shift of the inhibitory curves in both genotypes (Fig. 5, E and F), suggesting involvement of cardiac PDE1C. Under these experimental conditions (Fig. 5F), SIL could significantly suppress the cGMP-hydrolyzing activity in the murine hearts only at relatively high concentrations (IC50 > 1000 nM), which were likely not reached by the in vivo drug application protocol used herein (Adamo et al., 2010).
Together, the current data allow us to conclude that in the presence of amplified AT1R signal transduction in murine CMs and hypertrophic heart disease, potentially beneficial effects of cGMP do not involve SIL/PDE5 directly in these cells (Lukowski et al., 2010, 2014; Mokni et al., 2010; Degen et al., 2015).
Herein, we studied normotone transgenic mice that develop myocyte hypertrophy and fibrosis as a result of chronic AT1R overexpression specifically in CMs (Paradis et al., 2000). At baseline, AT1Rtg/+ hearts exhibited high expression of ANP and BNP (Fig. 3) as well as increased levels of cGMP and cGKI (Fig. 2), suggesting an activated NP/cGMP pathway. This was further supported by our finding that CNP induced a higher cGMP level in AT1Rtg/+ compared with control CMs (Fig. 1). Activation of the NP/cGMP pathway has been widely recognized in different cardiac stress models and is usually thought to be part of a counterregulatory mechanism suppressing Gαq-dependent Ca2+ signaling (Nishikimi et al., 2006; Kilić et al., 2007; Tokudome et al., 2008; Takimoto et al., 2009; Kinoshita et al., 2010; Klaiber et al., 2010; Koitabashi et al., 2010; Nishida et al., 2010; Frantz et al., 2013). As expected from previous studies (Adamo et al., 2010; Patrucco et al., 2014), SIL supplemented at high concentrations in drinking water stimulated cGMP levels in the cardiovascular system (Fig. 3, I and J). However, this drug did not affect adverse remodeling processes, expression patterns of hypertrophic and fibrotic marker genes (Figs. 3 and 4), or the contractility of the heart muscle in the αMHC-AT1Rtg/+ mouse model (Fig. 5). Based on these findings, we conclude that PDE5/cGMP does not interfere with amplified Ang II/AT1R signaling in the CM itself; hence, the reported cardioprotective effects of PDE5 inhibition should occur in cells distinct from CMs. Recently, a SIL coadministration regimen was shown to prevent functional decline and adverse remodeling of Ang II–stimulated hypertensive hearts (Westermann et al., 2012). However, this study did not differentiate between distinct cells types and systems that express the AT1R and may benefit from the SIL treatment supporting the concept of a cardioprotective SIL/PDE5/cGMP pathway in nonmyocytes (Lukowski et al., 2014). It seems also unlikely that the blood pressure– and afterload-lowering effects of SIL caused by the drug-induced changes in vascular tone are significant modulators of cardiac remodeling, as a corresponding antihypertensive treatment with hydralazine did not exhibit any signs of protection against the Ang II–induced cardiac pathology (Westermann et al., 2012). We detected a small but significant effect of long-term SIL on the turnover of the ECM components in αMHC-AT1Rtg/+ hearts (compare § groups in Fig. 4, C–F). The implication of this finding and its therapeutic relevance are unclear, in particular because 1) untreated αMHC-AT1Rtg/+ hearts in our study showed a similar overall tendency (Fig. 4, C–F) and 2) SIL did not affect any other parameter of the fibrotic response upon selective overexpression of the AT1Rtg/+ in the CMs. Yet we take this evidence as a rather consistent indicator for the antifibrotic actions attributed to SIL in different rodent models of adverse cardiac remodeling (Table 1).
The presented results from the αMHC-AT1Rtg/+ model seem to be in contrast to some of the previous reports suggesting a cardioprotective role for SIL/PDE5 in preclinical models of TAC- (Takimoto et al., 2005, 2009; Nagayama et al., 2009; Blanton et al., 2012), post-MI– (Ockaili et al., 2002; Salloum et al., 2008), and pulmonary hypertension–induced remodeling and dysfunction (Xie et al., 2012). However, not all aspects of the SIL-dependent cardioprotection were reproduced in other studies (Andersen et al., 2008; Schafer et al., 2009; Kukreja et al., 2014; Lee et al., 2015). It is important to note that in the current study, we assessed the role of SIL for opposing primary defects arising specifically in CMs in the absence of neurohormonal changes and elevated blood pressure.
So far, the cell types in which SIL interferes with Ang II/AT1R-induced cardiac remodeling remain elusive. Exposure of gene-targeted mice that lack cGKI in all non-SMCs with a hypertensive dose of Ang II demonstrated antifibrotic rather than antihypertrophic modes of action for SIL/PDE5/cGMP/cGKI signaling (Patrucco et al., 2014). It seems reasonable that cardiac fibroblasts are the primary target of SIL, as they display the full repertoire for the cross-talk between PDE5/cGMP/cGKI and AT1R, and the presence of PDE5 in murine CMs (Lukowski et al., 2010) and heart (Degen et al., 2015) was recently challenged. Increased cardiac and aortic cGMP concentrations in the presence of SIL (Fig. 3) suggest that the cGMP system in the αMHC-AT1Rtg/+ model is not fully activated; hence, cGMP-elevating compounds that do not act directly on PDE5 may still elicit cardioprotection upon AT1R overexpression. Because short-term SIL efficiently raised both the global cardiac and aortic cGMP levels, we find it very unlikely that the overexpression of AT1R per se can change signaling pathways (e.g., nitric oxide–dependent cGMP production) that render the drug inactive (Lee et al., 2015). However, we cannot rule out that the rather large increase in AT1R expression levels in CMs (Supplemental Fig. 1, E and F) overrides the potentially beneficial effects of SIL by a yet unknown mechanism in vivo, whereas directly blocking AT1Rs with losartan still prevented the progressive remodeling processes with cardiac dysfunctions in αMHC-AT1Rtg/+ mice (Paradis et al., 2000).
For the conditions tested, our global heart cGMP-PDE assay did not reveal any statistical differences in the cGMP-hydrolyzing activity between αMHC-AT1Rtg/+ and WT hearts (Fig. 5, E and F). However, we observed baseline elevation of cGMP/cGKI (Figs. 1 and 2) as well as a reliable increase in the cardiac cGMP levels upon 7 days of SIL in αMHC-AT1Rtg/+ mice (Fig. 2E). Apparently, it requires upstream factors, such as nitric oxide and/or the NPs, and intact cellular compartments to stimulate the cardiac cGMP system via SIL-sensitive PDEs. Because our FRET sensor does not allow reliable measurements of the basal cGMP in CMs, the cGMP-PDE activity under these conditions, i.e., in the absence of exogenous factors that activate the cGMP system, remains elusive (Fig. 1, A and C). However, our single-cell cGMP recordings under combined application of CNP plus IBMX (Fig. 1B) and of IBMX alone (Fig. 1D) suggest a dynamic regulation of the PDE-mediated effects on the NP/cGMP pathway upon a prolonged AT1R overexpression in CMs. As IBMX is a nonspecific PDE inhibitor, it was not possible to identify the isoform responsible for the changes in the cGMP-PDE response seen in intact CMs. Future studies will need to test the role of other PDEs, including the recently identified IBMX-insensitive cGMP-PDE9 (Keravis and Lugnier, 2012), in CMs (Lee et al., 2015) for the progressive heart stress phenotype in the αMHC-AT1Rtg/+ model.
Although we were unable to detect much of the cGMP-hydrolyzing PDE5 activity in total heart lysates (Fig. 5, E and F), the Ca2+/calmodulin sensitivity of the total heart lysates confirmed that cGMP-PDEs of the Ca2+/calmodulin-dependent PDE1 family contribute to the global cGMP-hydrolyzing activity in the healthy and hypertrophic murine heart (Lukowski et al., 2010). In particular, PDE1C may have a role in the integration of cGMP and cAMP signaling because this PDE hydrolyzes both second messengers with similar catalytic rates (Vandeput et al., 2007). Pharmacological inhibition of PDE1 activity significantly reduced cardiac hypertrophy in the isoproterenol-induced mouse model of hypertrophy (Miller et al., 2009), establishing a critical role for CM PDE1 in vivo. In Ang II–infused rats, PDE1 but not PDE5 activation reduced arterial cGMP bioavailability and increased contractile responsiveness (Giachini et al., 2011), suggesting a possible link between Ang II/AT1R and PDE1 signaling. To the best of our knowledge, the role of PDE1C has not been tested so far in an animal model of heart hypertrophy caused by Ang II infusion.
In summary, the results of our study show that the AT1R-induced profibrotic and prohypertrophic signal transduction in CMs is not sensitive to PDE5 inhibition by SIL. Taking into account recent human trials demonstrating clinical failure of SIL treatment in heart failure with preserved EF (Redfield et al., 2013) and diastolic dysfunction after myocardial infarction (Andersen et al., 2013), we believe it might be important to reassess the cell type–specific consequences of PDE5 inhibition in suitable preclinical models of cardiac stress, i.e., in mice that lack PDE5, PDE1C, and/or cGKI specifically in cardiac myocytes and in other cardiovascular cell types. In a clinical setting, the final outcome is certainly one of the most important issues of a drug treatment. If the mechanisms of SIL action in animal models include several cell types, this could well influence the efficacy and side effects of the treatment and how human clinical trials should be set up. From the scientific point of view, it is also important to define the actual molecular targets (and target cells) of SIL in vivo because only such studies can provide detailed insight into the role of, e.g., cGMP, SIL-sensitive PDEs (or other, potentially not yet identified, molecules) for the pathomechanism(s) of hypertrophic heart disease. This year marks more than 10 years of preclinical studies regarding the effects of SIL on the cardiac PDE5/cGMP/cGKI pathway (see Table 1 for an overview). Based on what we have learned so far, it seems reasonable to conclude that the potential of SIL to inhibit hypertrophic growth changes that develop due to a number of cardiovascular diseases may turn out to be rather limited.
The authors thank Tanja Schönberger, Michael Glaser, and Katrin Junger for excellent technical help, and Franz Hofmann (FOR923, Institut für Pharmakologie und Toxikologie, Technische Unversität München, München, Germany) and Peter Ruth (Department of Pharmacology, Toxicology and Clinical Pharmacy, Institute of Pharmacy, University of Tübingen) for helpful discussions.
Participated in research design: Lukowski, Friebe, Nikolaev.
Conducted experiments: Straubinger, Schöttle, Bork, Dünnes, Subramanian.
Contributed new reagents or analytic tools: Gawaz, Nemer, Russwurm.
Performed data analysis: Straubinger, Schöttle, Bork, Dünnes, Subramanian, Friebe, Nikolaev, Lukowski.
Wrote or contributed to the writing of the manuscript: Lukowski, Friebe, Nikolaev.
- Received May 20, 2015.
- Accepted July 7, 2015.
This work was supported by the Deutsche Forschungsgemeinschaft (DFG) Research Unit 2060 “cGMP signaling in cell growth and survival” with grants to R.L., A.F., and V.O.N. and by the Elite Programme financed by the Landesstiftung Baden-Württemberg (to R.L.). The DFG-funded Klinische Forschergruppe (KFO274) contributed to parts of the study.
- Ang II
- angiotensin II
- atrial natriuretic peptide
- angiotensin II type 1 receptor
- brain natriuretic peptide
- body weight
- cGMP-dependent protein kinase type I
- C-type natriuretic peptide
- collagen type 1 α-1 chain
- collagen type 1 α-2 chain
- connective tissue growth factor
- extracellular matrix
- ejection fraction
- Förster resonance energy transfer
- glyceraldehyde-3-phosphate dehydrogenase
- hypoxanthine phosphoribosyltransferase 1
- heart weight
- myosin heavy chain α-isoform
- matrix metalloproteinase 9
- natriuretic peptide
- phosphate-buffered saline
- polymerase chain reaction
- sarcoplasmic/endoplasmic Ca2+-ATPase 2
- α-smooth muscle actin
- smooth muscle cell
- transverse aortic constriction
- tissue inhibitor of metalloproteinase 1
- tibia length
- wild type
- Copyright © 2015 by The American Society for Pharmacology and Experimental Therapeutics