Primacy of angiotensin converting enzyme in angiotensin-(1–12) metabolism

Norihito Moniwa, Jasmina Varagic, Stephen W. Simington, Sarfaraz Ahmad, Sayaka Nagata, Jessica L. VonCannon, Carlos M. Ferrario

Abstract

Angiotensin-(1–12) [ANG-(1–12)], a new member of the renin-angiotensin system, is recognized as a renin independent precursor for ANG II. However, the processing of ANG-(1–12) in the circulation in vivo is not fully established. We examined the effect of angiotensin converting enzyme (ACE) and chymase inhibition on angiotensin peptides formation during an intravenous infusion of ANG-(1–12) in normotensive Wistar-Kyoto rats (WKY) and spontaneously hypertensive rats (SHR). WKY and SHR were assigned to a short ANG-(1–12) infusion lasting 5, 15, 30, or 60 min (n = 4–10 each group). In another experiment WKY and SHR were assigned to a continuous 15-min ANG-(1–12) infusion with pretreatment of saline, lisinopril (10 mg/kg), or chymostatin (10 mg/kg) (n = 7–13 each group). Saline or lisinopril were infused intravenously 15 min before the administration of ANG-(1–12) (2 nmol·kg−1·min−1), whereas chymostatin was given by bolus intraperitoneal injection 30 min before ANG-(1–12). Infusion of ANG-(1–12) increased arterial pressure and plasma ANG-(1–12), ANG I, ANG II, and ANG-(1–7) levels in WKY and SHR. Pretreatment with lisinopril caused increase in ANG-(1–12) and ANG I and large decreases in ANG II compared with the other two groups in both strains. Pretreatment of chymostatin had no effect on ANG-(1–12), ANG I, and ANG II levels in both strains, whereas it increased ANG-(1–7) levels in WKY. We conclude that ACE acts as the primary enzyme for the conversion of ANG-(1–12) to smaller angiotensin peptides in the circulation of WKY and SHR and that chymase may be an ANG-(1–7) degrading enzyme.

  • angiotensin I
  • angiotensin II
  • angiotensin-(1–7)
  • angiotensin-(1–12)
  • angiotensin converting enzyme
  • blood pressure
  • chymase
  • hypertension

the renin-angiotensin system (RAS) is a key factor in the pathogenesis of hypertension, atherosclerosis, heart failure, and renal disease (33, 37). In the classical RAS, angiotensinogen (Aogen) is cleaved by renin into angiotensin I (ANG I). ANG I is subsequently cleaved by angiotensin converting enzyme (ACE) to angiotensin II (ANG II), the primary effector of the RAS inducing vasoconstriction, sodium retention, and proliferative effects upon binding to ANG II type 1 receptor (AT1R) (32). In the past few decades, a more diverse biochemical pathway for the generation of angiotensin peptides downstream from ANG I was revealed through the identification of angiotensin-(1–7) [ANG-(1–7)] and angiotensin converting enzyme 2 (ACE2) (8, 13, 14, 32). The complexity of the biotransformation processes leading to ANG II formation is further expanded by the recent discovery of ANG-(1–12), a COOH-terminally extended form of ANG I (26). ANG-(1–12) is present in plasma and peripheral tissues including aorta, heart, and kidneys. Proof that ANG-(1–12) can function as an endogenous substrate for ANG II production was derived from experiments in which the dodecapeptide elicited a systemic dose-dependent vasoconstrictor response in the aorta from isolated rats that was abrogated by prior treatment with either an ACE inhibitor or an ANG II receptor blocker (26). In exploring the physiological role of ANG-(1–12), we showed that central immunoneutralization of ANG-(1–12) lowers arterial pressure and improves baroreflex sensitivity and heart rate variability in (mRen2)27 transgenic hypertensive rats (18), whereas microinjections of ANG-(1–12) into the nucleus tractus solitarii (NTS) increased systemic blood pressure, an effect abrogated by ACE inhibition or AT1R blockade (5). Early studies of ANG-(1–12) metabolism showed that renin did not generate ANG II from ANG-(1–12) (31), whereas a novel contribution of chymase as an ANG-(1–12) degrading enzyme was found in neonatal cardiac myocytes from spontaneously hypertensive rats (SHR) but not Wistar-Kyoto rats (WKY) (2). Additional studies suggested a diversity of enzymatic mechanisms accounting for the production of ANG II from ANG-(1–12) as chymase rather than ACE accounted for ANG-(1–12) metabolism in human atrial diseased tissue (1) and normal left ventricular myocytes (3). The diversity of enzymatic pathways associated with ANG-(1–12) biotransformation suggested a need to ascertain whether metabolic pathways by which ANG-(1–12) generates ANG II differ in the circulation versus the tissue compartment and whether the nature of the enzymatic pathways involved in this processes are not the same in normal and disease states. With this in mind, we investigated the effect of systemic inhibition of either ACE or chymase in the production of ANG I, ANG II, and ANG-(1–7) from exogenously infused ANG-(1–12) in both WKY and SHR.

MATERIALS AND METHODS

Animals

Experiments were conducted in male age-matched (12 to 13 wk) WKY (n = 51) and SHR (n = 40) obtained from Charles River (Wilmington, MA). All animal procedures were performed in accordance with National Institutes of Health (NIH) guidelines and were approved by the Wake Forest University animal care and use committee. Rats were housed in individual cages under a 12-h:12-h light (06:00–18:00)/dark (18:00–06:00) cycle, at a constant humidity and temperature, with free access to standard laboratory rat chow and tap drinking water.

Treatment Protocols

Experiment 1.

Rats were anesthetized with an intraperitoneal injection of 100 mg/kg thiobutabarbital (Inactin) and instrumented with a polyethylene catheter (PE-10) inserted into a right jugular vein and a PE-50 catheter placed into the right carotid artery. After a 30-min recovery from the surgical procedure, WKY and SHR were given a short ANG-(1–12) (2 nmol·kg−1·min−1 iv) infusion lasting for 5, 15, 30, or 60 min, respectively (n = 4–10 each groups).

Experiment 2.

A second group of WKY and SHR instrumented as described above were randomly assigned to receive a 15-min infusion of ANG-(1–12) at a dose of 2 nmol·kg−1·min−1 co-infused with either saline, lisinopril (10 mg/kg iv), or chymostatin (10 mg/kg ip). To ensure sustained ACE blockade, the lisinopril infusion began 15 min before the administration of ANG-(1–12) and continued during the 15-min ANG-(1–12) infusion. Chymostatin (10 mg/kg) was administered by bolus intraperitoneal injection 30 min before starting ANG-(1–12). At the completion of the 15-min ANG-(1–12) infusion, blood was collected from the right carotid artery catheter for measurements of ANG-(1–12), ANG I, ANG II, and ANG-(1–7) levels by radioimmunoassay (RIA). Lisinopril and chymostatin were obtained from Sigma-Aldrich (St. Louis, MO). ANG-(1–12) (Asp1-Arg2-Val3-Tyr4-Ile5-His6-Pro7-Phe8-His9-Leu10-Leu11-Tyr12) was custom-synthesized by GenScript (Piscataway, NJ). The doses of the drugs used in these experiments were shown before to suppress the activity of either chymase or ACE in converting ANG I into ANG II (13, 7, 9, 35).

In all experiments, arterial pressure and heart rate were measured with a computerized MP100 data acquisition system (BIOPAC Systems, Goleta, CA) using a solid-state pressure transducer connected to the catheter placed into a right carotid artery. ΔMAP (Δmean arterial pressure) was defined as the difference between 5-s averages taken each 30 s and 5-min average of baseline period.

Biochemistry

ANG-(1–12), ANG I, ANG II, and ANG-(1–7) peptides were measured by RIA as described by us elsewhere (15, 19, 22, 23).

Statistical Analysis

All values are expressed as means ± SE. Comparisons between WKY and SHR in Table 1 were analyzed by the unpaired Student's t-test. Changes in ΔMAP were analyzed by two-way ANOVA followed by the Bonferroni's post hoc test. Other data were analyzed by one-way ANOVA followed by the Tukey's post hoc test. All data were analyzed using GraphPad PRISM Version 5 (GraphPad, San Diego, CA) with P < 0.05 considered statistically significant.

RESULTS

A pilot study was performed to determine the dose of ANG-(1–12) to be used in the experiments. For this purpose, rats were infused with saline or ANG-(1–12) at the dose of 0.2, 2, 20, or 200 nmol·kg−1·min−1 at a rate of 0.1 ml·100 g−1·min−1. Saline infusion had no effect on blood pressure, whereas a dose of 0.2 nmol·kg−1·min−1 of ANG-(1–12) increased arterial pressure by <8 mmHg. Infusions of ANG-(1–12) at doses of 2, 20, and 200 nmol·kg−1·min−1 were associated with significant rises in arterial pressure and progressive increases in plasma ANG-(1–12) content (Fig. 1). On the basis of this pilot study, the dose of 2 nmol·kg−1·min−1 of ANG-(1–12) was chosen for metabolism studies given that these amounts were associated with robust increases in plasma ANG-(1–12) concentrations and minimal changes in arterial pressure.

Fig. 1.

Peak plasma ANG-(1–12) concentrations produced by intravenous infusion of vehicle (saline) or the dodecapeptide at doses between 0.2 and 200 nmol·kg−1·min−1.

Baseline characteristics of the WKY and SHR used in these experiments are documented in Table 1. As expected, the arterial pressure and heart rate of SHR were significantly higher than those measured in anesthetized WKY, whereas there were no significant differences of body weight between WKY and SHR.

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Table 1.

Baseline characteristics of Wistar-Kyoto rats and spontaneously hypertensive rats in Experiments 1 and 2

Experiment 1.

In both strains the elevation in mean arterial pressure induced by ANG-(1–12) administration peaked at 5–10 min (151 ± 41 mmHg, vs. baseline: 104 ± 5 mmHg, P < 0.0001, n = 10 in WKY and 194 ± 14 mmHg, vs. baseline: 146 ± 17 mmHg, P = 0.001, n = 10 in SHR) and remained higher than the baseline pressure until the end of the experiment.

The effects of ANG-(1–12) infusion on plasma angiotensin peptides concentrations in both WKY and SHR are illustrated in Fig. 2. ANG-(1–12), ANG I, ANG II, and ANG-(1–7) peaked at 15–30 min of infusion in WKY. However, in SHR the increases in angiotensin peptides concentrations as a function of time tend to be more variable and associated with ANG I values lower than those found in WKY (Fig. 2).

Fig. 2.

Plasma ANG-(1–12), ANG I, ANG II, and ANG-(1–7) levels measured in Wistar-Kyoto rats (WKY) and spontaneously hypertensive rats (SHR) infused with ANG-(1–12) (2 nmol·kg−1·min−1) at the completion of 0, 5, 15, 30, and 60 min of peptide administration. Values are means ± SE.

Experiment 2.

Administration of lisinopril in either WKY or SHR prevented the development of the pressor response to ANG-(1–12) administration (Fig. 3), whereas chymostatin was without effect. Figure 4 shows the effects of systemic blockade of either ACE or chymase on plasma angiotensin peptides concentrations at the end of the ANG-(1–12) infusion period in both WKY and SHR. The peak ANG-(1–12) levels attained in the blood of SHR is almost one-fifth of that in WKY, whereas co-infusion of lisinopril increased ANG-(1–12) more in SHR than in WKY (987% and 322% increase compared with co-infusion of saline, respectively). Likewise, plasma ANG I levels in SHR is almost one-third of that in WKY, whereas co-infusion of lisinopril increased ANG I more in SHR than in WKY (368% and 200% increase compared with co-infusion of saline, respectively). Lisinopril suppressed the increases in plasma ANG II levels induced by ANG-(1–12) infusion, whereas plasma ANG-(1–7) concentrations did not change (Fig. 4). Administration of chymostatin is associated with plasma concentrations of ANG-(1–12), ANG I, and ANG II that are not different from saline-treated rats for both WKY and SHR (Fig. 4). On the other hand, exposure to chymostatin results in higher levels of ANG-(1–7) in WKY but not SHR.

Fig. 3.

Top: changes in Δmean arterial pressure (ΔMAP) induced by the continuous infusion of ANG-(1–12) (2 nmol·kg−1·min−1) in WKY (top left) and SHR (top right) pretreated with saline, lisinopril, and chymostatin. Bottom: area under the curve (AUC) of ΔMAP in ANG-(1–12)-infused WKY (bottom left) and SHR (bottom right) pretreated with saline, lisinopril, and chymostatin. Values are means ± SE.

Fig. 4.

Plasma ANG-(1–12), ANG I, ANG II, and ANG-(1–7) levels found at the completion of a 15-min infusion of ANG-(1–12) (2 nmol·kg−1·min−1) in WKY and SHR pretreated with saline, lisinopril, or chymostatin. Values are means ± SE. *P < 0.05 vs. corresponding saline; **P < 0.01 vs. corresponding saline; ***P < 0.001 vs. corresponding saline; †P < 0.05 vs. corresponding lisinopril; ††P < 0.01 vs. corresponding lisinopril; †††P < 0.001 vs. corresponding lisinopril.

Fig. 5 shows the ratio of plasma ANG I/ANG-(1–12), ANG II/ANG-(1–12), ANG II/ANG I, and ANG-(1–7)/ANG II found at the completion of the 15-min infusion of ANG-(1–12) in saline-, lisinopril-, or chymostatin-pretreated WKY and SHR. Lisinopril tended to reduce the ANG I/ANG-(1–12) ratio compared with saline in both WKY and SHR, although the differences were not significant. ANG I-to-ANG-(1–12) ratios were significantly higher than those of lisinopril-pretreated rats in chymostatin-pretreated WKY and SHR. Lisinopril significantly reduced the ratio of ANG II to ANG-(1–12) and ANG II to ANG I, and significantly increased ANG-(1–7)-to-ANG II ratio compared with saline and chymostatin in both WKY and SHR. Chymostatin increased ANG II-to-ANG-(1–12) ratio compared with saline only in WKY.

Fig. 5.

The ratio of ANG I to ANG-(1–12), ANG II to ANG-(1–12), ANG II to ANG I, and ANG-(1–7) to ANG II found at the completion of a 15-min infusion of ANG-(1–12) (2 nmol·kg−1·min−1) in WKY and SHR pretreated with saline, lisinopril, or chymostatin. Values are means ± SE. **P < 0.01 vs. corresponding saline; ***P < 0.001 vs. corresponding saline; ††P < 0.01 vs. corresponding lisinopril; †††P < 0.001 vs. corresponding lisinopril.

DISCUSSION

Accumulating evidence continues to support a functional role of ANG-(1–12) as a substrate giving rise to the generation of angiotensin peptides in both rodents and humans. The current experiments show that ACE hydrolyzes ANG-(1–12) in the circulation of both WKY and SHR. Although previous studies suggested a differential contribution of ACE and chymase to ANG-(1–12) hydrolysis in rodents (2) versus humans (1, 3), this study is the first to assess the nature of the enzymatic pathways by which ANG-(1–12) acts as an endogenous source for the generation of angiotensin peptides in the circulation of a normotensive and hypertensive rat strain.

ACE, acting as a dipeptidyl carboxypeptidase, cleaves the Leu10-Leu11 bond of ANG-(1–12) to produce ANG I. Our finding that lisinopril, while blocking ANG II production, caused a large rise in plasma ANG I concentrations may reflect an independent pool of ANG I production resulting from activation of a renal renin-dependent mechanism due to the removal of the ANG II-mediated negative feedback on renal AT1 receptors. The decrease in the ANG I-to-ANG-(1–12) ratio in lisinopril-treated rats suggests that the increase of ANG-(1–12) due to inhibition of its degradation surpasses the increase of ANG I produced from the action of renin on Aogen. Because previous studies showed that renin does not participate in ANG-(1–12) production and metabolism (16, 25, 31), the increases in plasma ANG-(1–12) concentrations solely reflect the elimination of the activity of ACE as an ANG-(1–12) protease. In agreement with this interpretation, ex vivo studies showed a direct role of ACE in ANG-(1–12) metabolism (38). Our data also confirm that ANG II formation from ANG-(1–12) may require the intermediate generation of ANG I by truncation of the COOH-terminal amino acid sequence at positions 10–12 (38).

The absence of a hydrolytic action of chymase demonstrates a significant difference in the metabolism of ANG-(1–12) in tissues and humans. Chymase contributed to the processing of ANG-(1–12) into ANG II in neonatal myocytes obtained from SHR, whereas chymase was solely responsible for the direct generation of ANG II from ANG-(1–12) in human plasma membranes of atrial (1) and left ventricular myocytes (3). These data demonstrate the existence of different processing pathways for blood and tissues as well as the possibility of significant differences between rodents and humans. This interpretation agrees with the findings from numerous studies, which implicated chymase as the primary enzyme accounting for the production of ANG II in human cardiac and vascular tissues (6, 7, 11, 12, 20, 27).

Chymase is a chymotrypsin-like enzyme that is expressed in the secretory granule of mast cells (21). Chymase has a pathological role in cardiovascular disease through the conversion of ANG I to ANG II in the tissue including heart and vasculature in human, monkey, hamster, and mice (6, 7, 11, 12, 17, 20, 27, 30, 36). Elevated plasma chymotrypsin-like protease activity has been documented in patients with essential hypertension (4) and women with preeclampsia (34). Although the current study clearly demonstrates the absence of a chymase metabolizing activity in the circulation of WKY and SHR, we showed previously a chymase contribution to ANG-(1–12) metabolism in neonatal cardiomyocytes obtained from SHR but not WKY controls (2). These findings provide further evidence for differential enzymatic pathways in blood and tissues, as further documented by the studies enclosed in Table 2. The current findings also showed that the generation of ANG-(1–7) from infused ANG-(1–12) was significantly augmented in the presence of chymostatin in WKY but not SHR. This new finding indicates that chymase is associated with the degradation of ANG-(1–7) in WKY.

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Table 2.

The enzymes responsible for the metabolism of ANG-(1–12)

Our metabolism studies also revealed a significant difference in the amount of peptides found in the blood of SHR compared with WKY. These results indicate that ANG-(1–12) and ANG I are more rapidly metabolized to ANG II in SHR than in WKY probably because SHR have higher ACE activity than WKY. In agreement with this interpretation, we reported previously higher ACE activity levels in neonatal cardiac myocytes from SHR (2) as well as increased ANG-(1–12) uptake in cardiac myocytes from the same hypertensive strain (2). This interpretation does not exclude the additional possibility of increased hydrolytic activity in SHR as compared with WKY, since the differences in peptide concentrations in SHR were also shown for ANG I and ANG II.

We previously reported that in the bilateral nephrectomized rats plasma ANG-(1–12) was only moderately reduced, indicating that the production of ANG-(1–12) from Aogen in plasma is independent of renin (16). In agreement, in rats fed low-salt diet plasma ANG-(1–12) levels remained unchanged despite the increase in PRA (25). Furthermore, our previous study in isolated perfused hearts from normotensive and hypertensive rats demonstrated that exogenous ANG-(1–12) was metabolized to both ANG II and ANG-(1–7) independently of renin (31). Thus it seems that renin is involved neither in production of ANG-(1–12) from Aogen nor production of smaller angiotensin peptides from ANG-(1–12) in the systemic circulation. Further work will be necessary to identify the enzyme(s) accounting for the processing of Aogen into ANG-(1–12).

Although the data reported here are in keeping with previous ex vivo findings implicating ACE as an ANG-(1–12) convertase in rodents (2, 24, 26, 29, 38), the conclusions obtained from this study were derived from exogenous ANG-(1–12) administration of doses that resulted in plasma peptide levels much higher than expected in either physiological or pathological conditions. Lower doses of ANG-(1–12) may result in unmasking of alternate pathways since the ratio of substrate to enzyme activity may influence results.

In conclusion, we showed that ACE acts as the primary enzyme for the conversion of ANG-(1–12) into ANG II in the circulation of WKY and SHR, whereas chymase might contribute to the degradation of ANG-(1–7) in WKY.

GRANTS

This study was supported by the National Heart, Blood, and Lung Institute Grant HL-051952. N. Moniwa was supported in part by the Consortium for Southeastern Hypertension Control (COSEHC) 2012 Cardiovascular Postdoctoral Fellowship during these studies.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: N.M. and C.M.F. conception and design of research; N.M., S.W.S., S.A., S.N., and J.L.V. performed experiments; N.M., J.V., and C.M.F. analyzed data; N.M., J.V., S.W.S., S.A., S.N., J.L.V., and C.M.F. interpreted results of experiments; N.M. and C.M.F. prepared figures; N.M. and C.M.F. drafted manuscript; N.M., J.V., S.W.S., S.A., S.N., J.L.V., and C.M.F. approved final version of manuscript; J.V., S.W.S., S.A., S.N., J.L.V., and C.M.F. edited and revised manuscript.

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