Abstract
The aim of this study was to identify the role of chymase in the conversion of exogenously administered Big endothelin-1 in the mouse in vivo. Real-time polymerase chain reaction analysis detected mRNA of mucosal mast cell chymases 4 and 5, endothelin-converting enzyme 1a, and neutral endopeptidase 24.11 in pulmonary, cardiac, and aorta homogenates derived from C57BL/6J mice, with the latter tissue expressing the highest levels of both chymase isoforms. Furthermore, hydrolysis of a fluorogenic peptide substrate, Suc-Leu-Leu-Val-Tyr-7-amino-4-methylcoumarin, was sensitive to the chymase inhibitors Suc-Val-Pro-PheP(OPh)2 (200 μM) and chymostatin [(S)-1-carboxy-2-phenylethyl]-carbamoyl-α-[2-iminohexahydro-4(S)-pyrimidyl]-(S)-Gly-X-Phe-al, where X can be the amino acid Leu, Val, or Ile) (100 μM) in supernatants extracted from the same tissue homogenates. In anesthetized mice, Big endothelin-1, endothelin-1 (1–31), and endothelin-1 triggered pressor responses (ED50s, 0.67, 0.89, and 0.16 nmol/kg) that were all reduced or potentiated by selective endothelin ETA or ETB receptor antagonists, respectively, BQ-123 (cyclo[d-Asp-Pro-d-Val-Leu-d-Trp]) or BQ-788 (N-[N-[N-[(2,6-dimethyl-1-piperidinyl)carbonyl]-4-methyl-l-leucyl]-1-(methoxycarbonyl)-d-tryptophyl]-d-norleucine sodium salt), each at 1 mg/kg. The pressor responses to big endothelin-1 were significantly reduced by the neutral endopeptidase inhibitor thiorphan (dl-3-mercapto-2-benzylpropanoylglycine) (1 mg/kg) or the endothelin-converting enzyme inhibitor CGS 35066 [α-[(S)-(phosphonomethyl)amino]-3-dibenzofuranopropanoic acid] (0.1 mg/kg). In contrast, the responses to endothelin-1 (1–31) were abolished by thiorphan but unaffected by CGS 35066. In addition, Suc-Val-Pro-PheP(OPh)2 (20–40 mg/kg) reduced, by more than 60%, the hemodynamic response to big endothelin-1 but not to endothelin-1 (1–31) and endothelin-1. Finally, intravenous administration of big endothelin-1 induced Suc-Val-Pro-PheP-(OPh)2-sensitive increases in plasma-immunoreactive levels of endothelin-1 (1–31) and endothelin-1. The present study suggests that chymase plays a pivotal role in the conversion and cardiovascular properties of big endothelin-1 in vivo.
The endothelin-converting enzyme (ECE) produces the potent vasoactive peptide endothelin (ET)-1 (Yanagisawa et al., 1988) by hydrolyzing the tryptophan 21-valine 22 bond of its precursor, big endothelin-1 (1–38) [Big ET-1 (1–38)] (McMahon et al., 1991; D'Orléans-Juste et al., 2003). This ECE-dependent process had been shown initially to be sensitive to phosphoramidon [N-(α-rhamno-pyranosyl-oxy-hydroxy-phosphinyl)-Leu-Trp disodium salt], a neutral endopeptidase 24.11 (NEP)/ECE inhibitor (McMahon et al., 1991; D'Orléans-Juste et al., 2003). It is noteworthy that tissues derived from mice knockdown for both ECE-1 and ECE-2, which are nonviable past late gestational stages, still contain two thirds of mature endothelin peptides found in wild-type congeners, thus suggesting an important role for other proteases in the production of the 21-amino acid peptides (Yanagisawa et al., 2000).
Chymase, a mast cell-derived serine protease, also hydrolyzes Big ET-1 toward the intermediate peptide endothelin-1 (1–31), with no further hydrolysis products in rat tracheal rings (Nakano et al., 1997). In addition, the pivotal role of the NEP has been shown subsequently in the conversion of ET-1 (1–31) to ET-1 in human bronchial smooth muscle cells (Hayasaki-Kajiwara et al., 1999). Wypij et al. (1992) also reported a chymase-dependent processing of Big ET-1 to ET-1 in rat perfused lungs. Furthermore, a significant role for chymase in the vasculature was demonstrated by the chymostatin-sensitive constriction of rat aortic rings to Big ET-1 (Watts et al., 2007). On the other hand, our group has shown that intracardiac administration of Big ET-1 increases circulating levels of ET-1 (1–31), which in turn is readily converted by NEP to ET-1 in anesthetized rabbits (Fecteau et al., 2005). The contribution of chymase, however, in that alternate processing of Big ET-1 in vivo was not determined in the latter study (Fecteau et al., 2005).
Chymase is predominantly involved in the intramural production of angiotensin II in the human heart, arteries, and lungs, whereas the angiotensin-converting enzyme is responsible for genesis of the vasoactive peptide in the circulation (Fleming, 2006). Moreover, chymase is involved in extracellular matrix remodeling (Kielty et al., 1993; Kofford et al., 1997) and in the degradation of apolipoprotein E (Lee et al., 2002), thrombin, and fibrin (Tchougounova et al., 2003). The interstitial angiotensin II-forming characteristic of chymase is explained by the fact that this enzyme is interfered within the circulation by blood-borne endogenous protease inhibitors, such as α1 trypsin and α1 macroglobulin (Lindstedt et al., 2001). Intimal-located remnant granules derived from degranulated mast cells, however, release a heparin-bonded chymase that reduces the interfering properties of these high-molecular weight protease inhibitors on chymasic activity (Lindstedt et al., 2001; Miyazaki et al., 2006).
There are two known isoforms of chymase, namely α- and β-chymases. The former is mainly found in humans, monkeys, sheep, and hamsters, whereas the latter is also found in rats and mice (Doggrell and Wanstall, 2004). In mice, chymases are members of the murine mast cell protease family (mMCPs). Although mMCP-1, 2, and 4 are β-chymases, mMCP-5 is considered an α-chymase based on structural comparison or phylogenetic analysis (Gallwitz and Hellman, 2006). It is interesting that the single human α-chymase shares most of its functional homology and cleavage specificity with the mouse MCP-4, a β-chymase (Andersson et al., 2008).
A role for chymase-dependent production of angiotensin II in cardiovascular and inflammatory diseases in animal models and humans has been pharmacologically identified with specific inhibitors such as Suc-Val-Pro-PheP(OPh)2, NK-3201, SUN-C8257, and TY51184 (for review, see Miyazaki et al., 2006) and with the chymotrypsin-like inhibitor, chymostatin (Doggrell and Wanstall, 2004). This latter molecule, however, also nonspecifically reduces the activity of cathepsin G, cysteine proteases, and high-molecular weight proteasomes (Suga et al., 1993; Miyazaki et al., 2006). In the present study, we have characterized the processing of Big ET-1 in vivo with chymostatin and with a specific chymase inhibitor validated in different animal models of cardiovascular and inflammatory diseases, namely Suc-Val-Pro-PheP(OPh)2 (Oleksyszyn and Powers, 1994; Konno et al., 2005), a metabolically stable molecule with a biological half-life of more than 20 h (Soga et al., 2004).
Materials and Methods
Animals. C57BL/6J mice (16–19-week old males) were purchased from Charles River Canada (Montreal, QC, Canada). All animals were kept at constant room temperature (23°C) and humidity (78%) under a controlled light/dark cycle (6:00 AM–6:00 PM). Animal care and experiments were approved by the Ethic Committee on Animal Research of the Université de Sherbrooke in respect of the Canadian Council on Animal Care guidelines.
RNA Extraction and Real-Time PCR. Pulmonary, cardiac, and aortic tissues from C57BL/6J mice were homogenized in TRIzol reagent (Invitrogen Canada Inc., Burlington, ON, Canada) by using a tissue homogenizer (IKA Works, Wilmington, NC) or a tissue grinder (Kontes Glass, Vineland, NJ). The extraction of total RNA was performed in phenol/chloroform, precipitated in isopropanol, and washed in 75% ethanol. The RNA was dried and redissolved in diethyl-pyrocarbonate-treated water. A deoxyribonuclease treatment (DNase I; Invitrogen Canada Inc.) was effectuated on 1 μg of total RNA. Finally, a reverse transcription was performed using 25 units of avian myeloblastosis virus reverse transcriptase (Roche Diagnostics, Laval, QC, Canada) at 42°C for 60 min.
Each PCR contained 2 μl (∼100 ng) of cDNA and 300 nM primers in 25 μl of reactive mixture with 12.5 μl of SYBR Green master mix (Stratagene, La Jolla, CA). Quantitative PCR for actin, ECE-1a, mMCP-4, mMCP-5, and NEP were performed by monitoring in real time the fluorescence increase of SYBR Green using the MX3000P Multiplex Quantitative PCR System (Stratagene). Actin served as internal control.
The cycle profile was: 10 min at 95°C, followed by 40 cycles of denaturation for 30 s at 95°C, annealing for 1 min at 60°C, and extension for 30 s at 72°C. The length of amplified fragments for each PCR was: 190 bp for actin, 293 bp for ECE-1a (Lindenau et al., 2006), 77 bp for mMCP-4 (Kitaura-Inenaga et al., 2003), 71 bp for mMCP-5 (Kitaura-Inenaga et al., 2003), and 163 bp for NEP.
The sequences used were: actin F, 5′-GGG AAA TCG TGC GTG ACA TCA AAG-3′; actin R, 5′-CAT ACC CAA GAA GGA AGG CTG GAA-3′; mMCP-4 F, 5′-GAA GTG AAA AGC CTG ACC TGC-3′; mMCP-4 R, 5′-CAT GCT TTG TTG AAC CCA AGG-3′; mMCP-5 F, 5′-TTG CCA GCC TGT GAG GAA A-3′; mMCP-5 R, 5′-TAC AGA CAG GCC AGA TCG CAT-3′; ECE 1a F, 5′-CCC TGG TCT CAT GGT CTC GCT-3′; ECE 1a R: 5′-CGT AGC TGA AGA AGT CCT GGC A-3′; NEP F, 5′-GAG ACG GTG TGC TAA CTA CG-3′; and NEP R, 5′-ATC CAT CCA AGT AAG GTC ATC C-3′.
mRNA expression levels were calculated using the ΔCt method. In brief, 2-ΔCt was calculated to normalize gene of interest expression levels based on actin mRNA expression. Amplification reactions were tested for equal efficiency by real-time PCR of serial dilution (1:5). Primer quality and specificity (lack of primer-dimer and formation of a single product) were confirmed by melting curve analysis and migration on polyacrylamide/ethidium bromide gels.
Enzymatic Assays for the Determination of 7-Amino-4-Methylcoumarin-Forming Activity and Chymase-Like Activity. Chymase-like activity was determined in supernatants derived from C57BL/6J mice cardiac (whole heart), pulmonary (right and left lobes), and aortic (thoracic and abdominal) homogenates. The organs were dissected in small pieces and placed in a potassium phosphate buffer (0.1 g/400 ml), pH 8.0, at 4°C and homogenized using a tissue grinder (Kontes Glass). The homogenates were centrifuged at 14,000g for 20 min at 4°C. The pellets were discarded, and the supernatant was used for the assay. The substrate-specific enzymatic activity was measured at 37°C in a 0.2-ml reaction mixture comprising 100 mM fluorogenic peptide substrate, Suc-Leu-Leu-Val-Tyr-7-amino-4-methylcoumarin (MCA), in 100 mM potassium phosphate buffer, pH 8. The amount of MCA liberated from the substrate was determined fluorometrically (λex, 370; λem, 460 nm) with a fluorescence spectrophotometer (Molecular Devices, Sunnyvale, CA).
Hemodynamic Studies. All mice were induced and maintained under ketamine/xylazine (87/13 mg/kg i.m.) anesthesia throughout hemodynamic experiments. The left jugular vein and the right carotid artery were cannulated using a polyethylene-10 catheter, the former for intravenous injection and the latter for blood pressure monitoring or blood sampling. Mean arterial blood pressure (MAP) and heart rate (HR) were monitored using a Blood Pressure Analyzer-200A (Micro-Med, Tustin, CA). Basal MAP was averaged at 60.5 ± 1.7 mm Hg and HR at 163.8 ± 9.8 beats per min in all anesthetized mice tested in the present study (n = 130). Anesthetized animals were allowed to stabilize from the surgical procedure for 20 min before pharmacological interventions, and data were then recorded for 15 min after the intravenous administration of Big ET-1 (1–38), ET-1 (1–31), or ET-1 in each experiment. ED50 values were expressed as the dose (nanomoles per kilogram) of agonist necessary to induce 50% of the maximal pressor response and calculated using a fit curve analysis.
In another series of experiments, BQ-123 (ETA antagonist), BQ-788 (ETB antagonist), and phosphoramidon (ECE/NEP inhibitor) (Fukuroda et al., 1990) were administered intravenously 5 min before Big ET-1 or its derivatives. Thiorphan (NEP inhibitor) (Roques et al., 1980) and CGS 35066 (ECE inhibitor) (Jeng et al., 2002), were intravenously administered 15 and 20 min, respectively, before Big ET-1 and its derivatives. Finally, chymostatin (nonspecific chymase inhibitor) and Suc-Val-Pro-PheP(OPh)2 (specific chymase inhibitor) were given 20 min before the agonists via intraperitoneal injections.
Incubation times for antagonists (5 min) and inhibitors (15–20 min) were selected in accordance with previous reports (Honoré et al., 2002; Fecteau et al., 2005). Furthermore, vehicles used to dissolve both antagonists and inhibitors were systematically administered before each agonist for control purposes and were without effect on all parameters studied in each series of experiments.
Blood Sample Preparation. In separate groups of mice pretreated with vehicle, chymostatin, or Suc-Val-Pro-PheP(OPh)2, blood samples were collected 1 min after intravenous injection of Big ET-1 for the measurement of immunoreactive ET-1 (1–31) or ET-1. Blood samples (100 μl) were collected in trisodium citrate (3.5%) in a 9:1 ratio (v/v), immediately centrifuged for 1 min at 19,800g for blood content separation, and 50 μl of plasma samples was stored at -80°C until assayed by high-performance liquid chromatography (HPLC).
HPLC Separation and Radioimmunoassay and Enzyme Immunoassay Analysis. A reversed-phase HPLC analysis was performed using a Zorbax 300SB-C18 column (Agilent Technologies, Santa Clara, CA). Samples were eluted with a 35-min linear gradient of acetonitrile (from 28 to 40%) in 0.1% trifluoroacetic acid at a flow rate of 1 ml/min using a Waters model 510 pump (Waters, Milford, MA) as previously reported (Fecteau et al., 2005). Thirty-second fractions (500 μl) were separated in two, dried using a speed-vak evaporator (Thermo Fisher Scientific, Waltham, MA), and subsequently analyzed by radioimmunoassay (RIA) and enzyme immunoassay (EIA) for determination of ET-1 and ET-1 (1–31) levels, respectively. For calibration purposes, synthetic ET-1, ET-1 (1–31), or Big ET-1 were run separately or conjointly, and positions of the eluted standards were determined by ultraviolet absorbance (214 nm) and confirmed by RIA and EIA.
Immunoreactive (IR)-ET was measured by an RIA kit (RPA 555; GE Healthcare Bio-Sciences, Little Chalfont, Buckinghamshire, UK) as previously reported (Gratton et al., 1997). IR-ET-1 (1–31) was monitored with an EIA kit (Phoenix Biotech Corporation, Mississauga, ON, Canada). Plasma peptide concentrations are shown uncorrected for extraction recovery.
Drugs and Solutions. ET-1, ET-1 (1–31), Big ET-1, and phosphoramidon were purchased from Peptides International Inc. (Louisville, KY). BQ-788 was from American Peptide Co., Inc. (Sunnyvale, CA), and thiorphan and chymostatin were from Sigma Diagnostics Canada (Mississauga, ON, Canada). BQ-123 was synthesized by Dr. Witold Neugebauer (Department of Pharmacology, Université de Sherbrooke) (Brosseau et al., 2005). Finally, CGS 35066 was supplied by Tocris Bioscience (Ellisville, MO).
Drugs were dissolved in phosphate-buffered saline (PBS), pH 7.4, except for BQ-123, BQ-788, thiorphan, chymostatin, Suc-Val-Pro-PheP(OPh)2, and ET-1 (1–31), which were diluted in dimethyl sulfoxide and subsequently in PBS (10% dimethyl sulfoxide/PBS solution). CGS 35066 was diluted in 0.25 mM NaHCO3/PBS.
Statistical Analysis. Data are expressed as mean ± S.E.M. of n experiments. Statistical analyses were performed by Student's t test for two-group comparison, whereas analysis of variance followed by the Tukey's post hoc test was used for multiple comparisons in all applicable figures. Values of p ≤ 0.05 were considered significant.
Results
Analysis of mMCP4 and mMCP5 mRNA Levels.Table 1 shows the relative quantities of mMCP-4 and 5, ECE-1a, and NEP mRNAs from cardiac, pulmonary, and aorta homogenates. Relative to ECE1a, mMCP-4 and 5 mRNA levels were found in lower quantities in the heart and lungs but in much higher quantities in aortic homogenates. In contrast, ECE-1a mRNA levels were found at highest densities in cardiac homogenates. Finally, NEP was markedly expressed in all three organs tested but most importantly in the lungs and aorta (Table 1).
Monitoring of Chymase-Like Activity. The mouse aorta, heart, and lungs were analyzed for their MCA-forming activity. Suc-Val-Pro-PheP(OPh)2 and chymostatin, but not phosphoramidon, significantly reduced the MCA-forming activity in all mouse tissues (Fig. 1). Suc-Val-Pro-PheP(OPh)2 reduced by more than 20% the MCA-forming activity in supernatants derived from the lungs and heart and by over 40% the same hydrolytic process in that of aorta homogenates. As a final control, chymostatin efficiently reduced MCA-forming activity (by 60–70%) in all tissues tested (Fig. 1).
It is noteworthy that Suc-Val-Pro-PheP(OPh)2-sensitive chymase activity was higher in aortas (870 ± 50.8 fmol/min/mg) versus lungs (413.8 ± 17.1 fmol/min/mg) and heart (166.4 ± 10.4 fmol/min/mg) homogenates (*, p ≤ 0.05 aortas vs. lungs and heart). Each point represents the mean ± S.E.M. of seven experiments.
Big ET-1-, ET-1 (1–31)-, and ET-1-Induced Increases in MAP in the Anesthetized Mouse. To compare the effects of Big ET-1 and its derivatives on MAP, increasing doses of Big ET-1, ET-1 (1–31), or ET-1 were administered intravenously in anesthetized mice. Dose-dependent pressor response curves were performed with each of the three agonists with ET-1 being more potent than ET-1 (1–31) or Big ET-1 (ED50, 0.16 ± 0.14, 0.67 ± 0.36, and 0.89 ± 0.14 nmol/kg, respectively) (Fig. 2A). None of the doses tested for each of the three agonists induced significant changes in HR.
It is noteworthy that onset and maximal pressor responses were attained at an earlier time point with ET-1 than ET-1 (1–31) or Big ET-1. Maximal pressor response was attained at 1.5 min for ET-1, 9 min for ET-1 (1–31), and 15 min for Big ET-1 (Fig. 2B). Doses higher than 1 (for ET-1) or 10 nmol/kg [for Big ET-1 and ET-1 (1–31)] were not used to avoid lethality.
Contribution of ETA and ETB Receptors in Endothelin-Induced Pressor Responses. Big ET-1, ET-1 (1–31), or ET-1 administered at 1, 1, or 0.5 nmol/kg, respectively, induced similar pressor responses that were assessed in the absence or presence of selective ETA or ETB antagonists, BQ-123 and BQ-788, respectively. BQ-123 and BQ-788 significantly repressed and potentiated, respectively, the hemodynamic responses afforded by the three peptides tested (Fig. 3). As a control, the pressor response to a selective ETB agonist (IRL 1620) was unaffected by BQ-123 but markedly reduced by BQ-788 (vehicle, 23.4 ± 0.9 mm Hg; BQ-123, 18.3 ± 2.2 mm Hg; BQ-788, 12.5 ± 1.2 mm Hg; *, p ≤ 0.05, BQ 788 versus vehicle; n = 4).
Role of the ECE and NEP in the Big ET-1- and ET-1 (1–31)-Induced Pressor Responses. The increase in MAP triggered by Big ET-1 was reduced in a dose-dependent fashion by phosphoramidon, CGS 35066, and thiorphan (ECE/NEP-, ECE-, and NEP-selective inhibitors, respectively) (Fig. 4A). CGS 35066 was 10- to 100-fold more potent than thiorphan and phosphoramidon (0.1 versus 1 and 10 mg/kg) as an inhibitor of Big ET-1-induced responses (Fig. 4A). Thiorphan was not tested at higher doses because of cardiotoxic effects. Figure 4B, in contrast, shows that the pressor response to ET-1 (1–31) was abolished by phosphoramidon and thiorphan but unaltered by the ECE-selective inhibitor CGS 35066. As a final control, none of the three above-mentioned inhibitors interfered with the pressor response to ET-1 (Fig. 4B).
Big ET-1 Induces Chymase Inhibitor-Sensitive Pressor Response. The pressor responses to Big ET-1, ET-1 (1–31), and ET-1 were also tested in mice pretreated with chymostastin (20 mg/kg). The nonspecific chymase inhibitor reduced the pressor response to Big ET-1 (1 nmol/kg) without affecting those of ET-1 (1–31) (1 nmol/kg) and ET-1 (0.5 nmol/kg) (Fig. 5A).
In another series of experiments, Suc-Val-Pro-PheP(OPh)2 markedly reduced the pressor response induced by Big ET-1 (Fig. 5A). Moreover, the specific chymase inhibitor did not alter those afforded by ET-1 (1–31) or ET-1 (Fig. 5B). In addition, dual ECE/chymase inhibition after coadministration of CGS 35066 and Suc-Val-Pro-PheP(OPh)2 further inhibited the maximal increase of MAP observed with systemically administered Big ET-1 (Fig. 5A). Finally, Fig. 5C shows that the bolus administration of Suc-Val-Pro-PheP(OPh)2 induces a long-lasting inhibition of the pressor response to Big ET-1 (1 nmol/kg).
Chymase-Dependent Conversion of Big ET-1 to ET-1 in Vivo. Plasma levels of both ET-1 (1–31) and ET-1 after systemic administration of Big ET-1 were measured by EIA and RIA, respectively, in mice pretreated with vehicle or chymase inhibitors. Figure 6A shows that the reverse HPLC gradient used was able to effectively separate a high amount of synthetic Big ET-1 from ET-1 (1–31) and ET-1 in extracted plasma via ultraviolet absorbance detection. Moreover, when Big ET-1 was systemically administered in anesthetized mice, the HPLC approach combined with immunodetection confirmed the separation of ET-1 (1–31) and ET-1 produced de novo (Fig. 6B). Basal levels of both ET-1 (1–31) and ET-1 were under detection threshold levels because of the limited systemic blood volumes than can be acutely drawn from mice without inducing hypovolemia.
Furthermore, the nonspecific chymase inhibitor, chymostatin, abolished the increase in plasma levels of ET-1 (1–31) after administration of Big ET-1 (Fig. 7A). In addition, chymostatin did not interfere with the increased plasma levels of ET-1 (1–31) after systemic administration of the same peptide (Fig. 7B). Finally, Suc-Val-Pro-PheP(OPh)2 administered before Big ET-1 abolished the increases in plasma levels of immunoreactive ET-1 (1–31) (Fig. 7C) and ET-1 (Fig. 7D).
Discussion
The two main results of the present study are, firstly, that chymase is significantly involved in the pressor response to Big ET-1 and that ET-1 (1–31) requires the activity of the neutral endopeptidase to increase blood pressure in the mouse model. Furthermore, our data suggest that chymase is also pivotal in the increased plasma levels of ET-1 (1–31) and ET-1 after systemically administering Big ET-1.
Unlike the ECE or the NEP, chymase is not a membrane-bound serine protease, albeit the activity of the latter enzyme is limited in the circulation by blood-borne endogenous inhibitors (Lindstedt et al., 2001). It is interesting that in the present study, we were able to detect chymase-like activity in the supernatant of cardiac, pulmonary, and aorta homogenates of the mouse.
It is noteworthy that chymase is also located in the adventia and media of human coronary arteries (Borland et al., 2005); therefore, a significant contribution of that particular vasculature should also be taken into account in the overall chymase activity detected in the present study in the whole-heart homogenates of the mouse. Finally, a marked increase in cardiac mMCP-4 and mMCP-5 mRNA levels has been shown in a mouse model of heart failure (Kitaura-Inenaga et al., 2003), thus confirming the present results as far as the presence of both chymase isoforms are concerned in cardiac homogenates.
We suggest that chymostatin is more efficient than Suc-Val-Pro-PheP(OPh)2 to reduce the hydrolysis of the fluorogenic peptide Suc-Leu-Leu-Val-Tyr-MCA in all tissues tested because of the nonspecific characteristics of the former inhibitor (Suga et al., 1993; Miyazaki et al., 2006) and the nonexclusive cleavage of the fluorogenic substrate by chymase (Nakano et al., 1997). It is also of interest that the enhanced inhibitory properties of the chymase inhibitor Suc-Val-Pro-PheP(OPh)2 in aortic homogenates compared with the lungs and cardiac tissue-derived homogenates is correlated with the marked mMCP-4 and -5 mRNA levels measured in the conductance vessel.
We further show that the NEP inhibitor thiorphan partially blunts or abolishes the pressor response to Big ET-1 or ET-1 (1–31), respectively, in the mouse model. Furthermore, a selective ECE inhibitor, CGS 35066, at low doses, only partially abrogates the pressor response to Big ET-1 without affecting that afforded by ET-1 (1–31). Doses of CGS 35066 (i.e., 0.25 nmol/kg) induced a nonspecific inhibition of the NEP in the mouse model (results not shown) as previously reported by Jeng et al. (2002), thus explaining why the ECE inhibitor at 1 but not 0.1 nmol/kg abolished the pressor response to Big ET-1 in the present study.
Finally, the dual inhibition of ECE and chymase promotes an additional reduction in the pressor response to Big ET-1. Thus, we have shown that the pressor responses to Big ET-1 and ET-1 (1–31) are predominantly amenable by combined chymase/ECE for the former and fully by NEP processes for the latter peptide.
In the present in vivo model, ET-1 (1–31) requires the obligatory contribution of NEP to induce its pressor response. Albeit not demonstrated in the present study, we suggest that first pass of intravascularly administered ET-1 (1–31) in the lungs, which is an NEP-rich organ (Baraniuk et al., 1995), may be involved in this process. In support of our hypothesis, high mRNA levels of NEP 24.11 were measured in murine pulmonary and, interestingly, in aorta-derived homogenates. The mRNA monitoring and MCA studies presented here point out the presence of marked chymase and NEP activity in the aorta, thus providing evidence toward a contribution of this alternate pathway in the processing of Big ET-1 in the conductance vessel as also previously shown by Watts et al. (2007) in isolated rat aortic rings. We also suggest that large conductance vessels such as the aorta are the main targets for the inhibition of chymase-dependent interstitial conversion of Big ET-1 to ET-1 in the cardiovascular system.
We also show that both Suc-Val-Pro-PheP(OPh)2 and chymostatin abolished the increases in ET-1 (1–31) plasma levels afforded by systemic administration of Big ET-1. In addition, specific inhibition of chymase markedly blunted the increase plasma levels of ET-1 after administration of the 38-amino acid precursor. Our data also show that chymostatin had no effect on ET-1 (1–31) plasma levels after systemic administration of the same peptide. Therefore, we demonstrate for the first time that chymase is the enzyme predominantly involved in the processing of intravascularly administered Big ET-1 to ET-1 in the mouse model. It is noteworthy that we did not assess the contribution of the ECE on the increased ET-1 plasma levels after systemic administration of Big ET-1 in the mouse. Firstly, specific chymase inhibition virtually abolished the increased plasma levels of both ET-1 (1–31) and ET-1 in the present study. Furthermore, Okumura et al. (1993) had shown previously that the NEP/ECE inhibitor, phosphoramidon (10 mg/kg), does not alter the increase in ET-1 plasma levels after systemic administration of Big ET-1 in the mouse model, unlike what we have reported previously in the rabbit (Gratton et al., 1997). This discrepancy points out interspecies variations and also supports the concept that chymase plays a more important role in the systemic conversion of Big ET-1 than the ECE in the mouse model.
On the other hand, it is also of importance to point out that the increased levels of ET peptides after administration of the precursor should be considered as a spillover (i.e., representing a fraction) of interstitially preformed ET-1 (1–31) and ET-1, thus explaining why chymase inhibition abrogates the increased plasma levels of both peptide while only reducing the pressor responses after administration of Big ET-1.
Based on the above considerations, we also suggest that the specificity of ECE inhibitors in the cardiovascular system must be assessed pharmacologically and biochemically by monitoring circulating metabolites. By not correlating these two parameters concomitantly, the contribution of chymase/NEP-dependent production of ET-1 from Big ET-1 in vivo is likely to be underestimated.
Maurer et al. (2004), on the other hand, have suggested that mast cell-derived chymase degrades in a chymostatin-sensitive fashion intraperitoneal administered ET-1 in the mouse model. To support their hypothesis, Maurer et al. (2004) used mast cell-deficient mice. However, it is well established that peritoneal mast cells contain not only chymase but also several other proteases, such as serine proteases and carboxypeptidase A (Pejler et al., 2007). It is noteworthy that Schneider et al. (2007) have recently identified carboxypeptidase A as the predominant enzyme involved in murine mast cell-dependent degradation of ET-1. Therefore, the present study supports the concept that proteases other than chymase may inactivate ET-1 in the mouse model.
Onset and maximal pressor responses for the three peptides also support the concept of differential maturation of Big ET-1 in vivo. The onset, the time required to obtain a maximal pressor response, and the relative affinity correlate with the length of the peptide administered with ET-1 being the more potent and effective and Big ET-1 being the lesser. These results are in accordance with those of Nakano et al. (1997) in rat tracheal rings.
Finally, some groups have suggested that ET-1 (1–31) acts as a selective ETA agonist, albeit in in vitro models (Ishizawa et al., 2004; Zhou et al., 2006). In vivo, in the mouse model, however, ETA or ETB receptor blockade reduce or amplify, respectively, the pressor responses to Big ET-1, ET-1 (1–31), and ET-1. These results suggest that irrespective of the source (i.e., chymase/NEP dependent or ECE dependent), the ET-1 produced from converted Big ET-1 or ET-1 (1–31) activates the same receptors types to induce its hemodynamic effect in C57BL/6J mice. ETB receptors, most probably located on the endothelium (to release nitric oxide), seem to limit the pressor responses to Big ET-1, its 31 amino acids intermediate, and ET-1 in this mouse model. The concept that ET-1 (1–31), similarly to ET-1, activates both ETA and ETB receptors in the cardiovascular system suggests that the 31-amino acid peptide must be converted to ET-1, via an NEP-dependent process in vivo in the mouse as previously demonstrated in the rabbit model by our group (Fecteau et al., 2005).
Therefore, our study supports the concept of an alternative chymase/NEP-dependent pathway in the in vivo processing of exogenous Big ET-1 and confirms the important role of NEP in the cardiovascular effects of ET-1 (1–31) in the mouse model. Thus, not only the ECE but also chymase matures ET-1 from its 38-amino acid precursor in vivo.
To our knowledge, this is the first report demonstrating the chymase-dependent processing of Big ET-1 in vivo. Moreover, the present results obtained with the specific inhibitor, Suc-Val-Pro-PheP(OPh)2, support previous in vitro studies reporting chymase-dependent mechanisms involved in the conversion of Big ET-1 in terms of production of the intermediate ET-1 (1–31) by human mast cell-extracted chymase (Nakano et al., 1997) and vasoconstriction in rat arterial vessels (Watts et al., 2007).
In conclusion, we suggest that interfering specifically with the ECE pathway with selective inhibitors would leave unaltered the production of ET-1 (1–31) and the subsequent NEP-dependent production of ET-1. However, whether the chymase/NEP pathway constitutes a therapeutically valid target toward the endothelin system in vascular diseases will require the full assessment of this paradigm with potent and highly specific chymase inhibitors.
Acknowledgments
We thank Dr. Robert Day for logistical support with the fluorogenic assay experiments and Helen Morin for secretarial support.
Footnotes
-
This work was supported by the Canadian Institutes for Health Research and by the Etienne Lebel Clinical Research Center.
-
The authors of the present manuscript declare that there are no financial links, including consultancies with manufacturers of material or devices described in the article, and no links to the pharmaceutical industry or regulatory agencies or any other potential conflicts of interest.
-
Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
-
doi:10.1124/jpet.108.142992.
-
ABBREVIATIONS: ECE, endothelin-converting enzyme; ET, endothelin; phosphoramidon, N-(α-rhamno-pyranosyl-oxy-hydroxy-phosphinyl)-Leu-Trp disodium salt; NEP, neutral endopeptidase; Big ET-1 (1–38), big endothelin-1 (1–38); chymostatin, [(S)-1-carboxy-2-phenylethyl]-carbamoyl-α-[2-iminohexahydro-4(S)-pyrimidyl]-(S)-Gly-X-Phe-al, where X can be the amino acid Leu, Val, or Ile; mMCP, murine mast cell protease; PCR, polymerase chain reaction; F, forward; R, reverse; MCA, 7-amino-4-methylcoumarin; MAP, mean arterial pressure; HR, heart rate; BQ-123, cyclo(d-Asp-Pro-d-Val-Leu-d-Trp; BQ-788, N-[N-[N-[(2,6-dimethyl-1-piperidinyl)carbonyl]-4methyl-l-leucyl]-1-(methoxycarbonyl)-d-tryptophyl]-d-norleucine sodium salt; thiorphan, dl-3-mercapto-2-benzylpropanoylglycine; CGS 35066, α-[(S)-(phosphonomethyl)amino]-3-dibenzofuranopropanoic acid; HPLC, high-pressure liquid chromatography; RIA, radioimmunoassay; EIA, enzymatic immunoassay; IR, immunoreactive; PBS, phosphate-buffered saline; NK-2301, 2-(5-formylamino-6-oxo-2-phenyl-1,6-dihydropyrimidine-1-yl)-N-[[3,4-dioxo-1-phenyl-7-(2-pyridyloxy)]-2-heptyl]acetamide; SUN-C8257, 3-[(3-amino-4-carboxy)phenylsulfonyl]-7-chloroquinazoline-2,4(1H,3H)-dione; TY-51184, 2-[4-(5-fluoro-3-methylbenzo-[b]thiophen-2-yl)sulfonamide-3-methanesulfonylphenyl] oxazole-4-carboxylic acid; IRL 1620, Suc-[Glu9,Ala11,15]-endothelin-1(8-21).
- Received July 5, 2008.
- Accepted November 4, 2008.
- The American Society for Pharmacology and Experimental Therapeutics