| | Endogenous ghrelin released in response to endothelin stimulates growth hormone secretion in cattleReceived 25 April 2009; received in revised form 25 July 2009; accepted 25 July 2009. published online 07 September 2009. Abstract The purpose of this study was to evaluate whether circulating ghrelin and growth hormone (GH) concentrations in cattle are regulated by endothelin-1 (ET-1), endothelin-3 (ET-3), and secretin. Six Holstein steers (242 ±1 d old, 280.5±4.4kg body weight [BW]; mean±SEM) were allocated randomly in an incomplete Latin square design to receive each of 4 treatment compounds (vehicle, ET-1, ET-3, and secretin) with 1-d intervals between successive treatments. The treatment compounds were injected intravenously via a catheter inserted into the external jugular vein of each steer. Blood was sampled from the indwelling catheter at -30, -15, 0, 5, 10, 15, 20, 30, 45, 60, 90, 120, 150, and 180min. Plasma ghrelin and GH responses to the treatment compounds were measured by a double-antibody radioimmunoassay system. Data were analyzed by using a MIXED procedure of SAS, version 9.1. Plasma acyl ghrelin, total ghrelin, and GH concentrations were increased by both ET-1 and ET-3 injection (ET-1 injection: 311±15pg/mL vs 245±15pg/mL, 2.4±0.2ng/mL vs 1.61±0.05ng/mL, 4.73±0.92ng/mL vs 1.17±0.09ng/mL for acyl ghrelin, total ghrelin, and GH, respectively; ET-3 injection: 337±27pg/mL vs 245±15pg/mL, 2.6±0.1ng/mL vs 1.61±0.05ng/mL, 5.56±0.97ng/mL vs 1.17±0.09ng/mL for acyl ghrelin, total ghrelin, and GH, respectively; P<0.01). Ghrelin and GH concentrations were not changed by secretin injection throughout the experimental periods. These results indicate that ET-1 and ET-3 stimulate ghrelin and GH secretion in cattle and demonstrate for the first time that endogenous ghrelin released in response to endothelin injection stimulates GH secretion in vivo in cattle. Article Outline• Abstract • 1. Introduction • 2. Materials and methods • 2.1. Allocation of animals • 2.2. Peptides • 2.3. Administration of peptides • 2.4. Measurements of plasma hormones and metabolites • 2.4.1. Radioimmunoassay for endothelins • 2.4.2. Radioimmunoassay for secretin • 2.4.3. Radioimmunoassay for ghrelin, GH, and insulin • 2.4.4. Radioimmunoassay for glucagon • 2.5. Statistical analysis • 3. Results • 3.1. Effect of ET-1 injection on ghrelin, GH, insulin, glucagon, and metabolites • 3.2. Effect of ET-3 injection on ghrelin, GH, insulin, glucagon, and metabolites • 3.3. Effect of secretin injection on ghrelin, GH, and metabolites • 4. Discussion • Source of funding • References • Copyright 1. Introduction  Growth hormone (GH) is of great importance in controlling growth processes in ruminants. Its secretion from the anterior pituitary gland is preferable, as it stimulates the anabolic processes leading to skeletal and muscle tissue growth in these animals. To improve growth efficiency, it is crucial to understand not only the regulators of GH secretion, but also the relationship between these regulators and other peptide hormones. Ghrelin is one of the growth-hormone–releasing peptides originally isolated from the rat stomach as an endogenous ligand for the growth hormone secretagogue receptor (GHS-R) [1]. Ghrelin is predominantly produced by the stomach, whereas substantially lower amounts are derived from the bowel, pancreas, pituitary, kidney, and placenta [2]. Ghrelin causes a positive energy balance not only by stimulating GH release [1], [2], [3], [4], [5], but also by stimulating food intake and decreasing fat utilization [6], [7]. Circulating levels of ghrelin were stimulated by adrenaline, noradrenaline, endothelin, and secretin [8] and were suppressed by food intake [9], somatostatin [8], [10], urocortin-1 [11], bombesin, gastrin-releasing peptide [8], exendin-4 [12], and GH [13]. The regulation of ghrelin secretion by insulin, leptin, glucagon, glucagon-like peptide-1 (GLP-1), gastrin, and cholecystokinin (CCK) is under debate [8], [14], [15], [16], [17], [18], [19], [20]. It can be postulated that the regulatory mechanism of ghrelin secretion is conclusively complex and that there is relatively little information on the regulation of ghrelin and GH secretion, especially in ruminants. Endothelins (ETs) are a peptide family first isolated from porcine endothelial cells as a potent vasoconstriction peptide with 2 sets of intrachain disulfide bonds [21] and consisting of 3 isoforms, ET-1, ET-2, and ET-3 [21], [22]. Endothelin-1 and ET-3 are widely expressed in mammalian tissues, including the gastrointestinal tract, pituitary gland [23], [24], and adrenal gland [25]. There are 2 subtypes of receptors for ET: endothelin A receptor (ETA) and endothelin B receptor (ETB). Endothelin A receptor has the highest selectivity to ET-1, whereas ETB accepts the 3 isopeptides of ETs equally [26], [27]. Both receptors are widely distributed in a wide variety of tissues, including the anterior pituitary gland [28], adrenal gland [29], stomach, intestine [23], [29], liver, and fat cells [30], [31]. Several lines of evidence suggest that ETs may function as neuromodulators or neurotransmitters within the central nervous system (CNS). These evidence suggest that ETs participate in the regulation of diverse biological functions. Secretin is a hormonal substance that was first discovered in 1902 [32]. It is produced by the intestinal endocrine cells and neurons of different brain regions [32], [33], and it is considered as part of the brain-gut axis, as it takes part in the pancreatic and gastrointestinal system to control appetite [34], [35]. Results from a more recent microdialysis study illustrate the effects of ET-1 and secretin on ghrelin secretion from rat stomach ghrelin cells [8]. It remains to be investigated whether or not these peptides regulate ghrelin and GH secretion in ruminants. The present study was designed to evaluate the effects of exogenous ET-1, ET-3, and secretin injection on ghrelin and GH secretion in cattle. 2. Materials and methods All animal procedures undertaken were approved by the Animal Care and Use Committee of Obihiro University of Agriculture and Veterinary Medicine. 2.2. Peptides Human ET-2 (Code: 4209-s, Lot: 420604), human ET-3 (Code: 4199-s, Lot: 530726), and human glucagon (Code: 4098-s, Lot: 570419) were purchased from Peptide Institute, Inc., Osaka, Japan. Human oxyntomodulin (Code: 028-22, Lot: 425854) and human secretin (S-7147, Lot: 77H03851) were purchased from Phoenix Pharmaceuticals, Inc. and Sigma Chemical Company, respectively. Endothelin-1 (CSCSSLMDKECVYFCHLDIIW), bovine ET-3 (CTCFTYKDRECVYYCHLDIIW, accession XM 591496), bovine secretin (HSDGTFTSELSRLRDSARLQRLLQGLV-NH2), [Cys-3]-bovine secretin (4-26) amide, [Cys-18]-bovine glucagon (19-29), bovine glucagon antagonist (SQGTTSEYSKYLDSRRAQDFVQWLMNT-NH2), bovine GLP-1 (7-36) amide (HAEGTFTSDVSSYLEGQAAKEFIAWLVKGR-NH2), and bovine glucagon-like peptide-2 (GLP-2) (HADGSFSDEMNTVLDSLATRDFINWLLQTKITDRK) were synthesized by Fmoc (9-fluorenylmethoxycarbonyl) solid-phase peptide synthesis procedures and purified by reverse-phase HPLC (TSKgel ODS-120A column; TOSOH, linear gradient of 0%-60% CH3CN). Purified peptides were lyophilized and stored at -30°C. These peptides were used for injection or as the cold standard, or they were to be labeled for radioimmunoassay (RIA). Synthesized [Cys-3]-bovine secretin (4-26) amide was conjugated with maleimide activated mariculture keyhole limpet hemocyanin (mcKLH) and used for raising antibody in rabbits. The biological activity of secretin was checked in 7-wk-old male rats. Secretin was administered intravenously at the dosage of 50μg/kg BW, and the amount of pancreatic juice was measured. The secretion of pancreatic juice was more than 10 times greater than the baseline level for 1h after the administration of secretin. 2.3. Administration of peptides The dosage of endothelin, 1μg/kg, was chosen based upon the results of Rossi and Scharrer (1992), in which intraperitoneal injection of ET-1 increased plasma glucose and lipid concentrations in Pygmy goats [36]. It has been reported that secretin depressed food intake in sheep at 8 clinical units (CU)/kg (equivalent to 0.7μg/kg) [35]. In our experiment, secretin 50μg/kg BW was used as a pharmacological dose. This dosage, about 10,000 times the basal plasma concentration in sheep [37], was calculated according to the BW of steers. The blood volume of steers was estimated to be 8% of BW [38]. Peptides were administered in random order. The day before the experiment, a polyethylene catheter was inserted into the external jugular vein of each steer, and patency was maintained with heparinized saline (10 IU heparin/mL saline). This catheter was used for both injection of peptide and sampling of blood. Body weight was measured 1 d before each experiment. Throughout the experimental period, steers were loosely chained and allowed ad libitum access to hay and water. On the experiment days, lyophilized ET-1, ET-3, and secretin powder were dissolved in sterilized, double-distilled water (at a concentration of 1mg/mL for ET-1 or ET-3, and 20mg/mL for secretin, respectively), and the desired concentration of each peptide was prepared using 0.1% bovine serum albumin (BSA) in saline to obtain 5mL of total injection volume. After the baseline samples (-30,-15, and 0min) had been taken, treatment compounds were administered intravenously and flushed with 5mL of heparinized saline. Blood was sampled at 5, 10, 15, 20, 30, 45, 60, 90, 120, 150, and 180min after administration of the treatment compounds. For all blood sampling, approximately 4mL of blood was collected into a syringe and immediately transferred into the pre-chilled polystyrene tube containing 40 IU of heparin sodium. Blood tubes were immediately centrifuged at 4°C for 30min at 1870×g, and plasma was kept at -30°C until analysis. Plasma samples for ghrelin measurement were acidified with 1N HCl with the ratio of 1:20 (HCl:plasma) to hinder the breakdown of acyl ghrelin before freezing. 2.4. Measurements of plasma hormones and metabolites Endothelin, secretin, ghrelin, GH, insulin, and glucagon concentrations were measured using a double-antibody RIA method. All assays were performed with triplicate standards and duplicate samples. All 125I-labeled antigens used to measure plasma concentrations of peptides were radioiodinated using the chloramine-T method [39] and purified by HPLC. Plasma glucose and nonesterified fatty acid (NEFA) concentrations were measured by commercially available kits (Code No. 43990901 and 279-75401, respectively, Wako, Japan). 2.4.1. Radioimmunoassay for endothelins Plasma endothelins were measured using relevant 125I-labeled ET as the tracer and anti-endothelin-1 antiserum (catalog number E1645, Lot: 126K4848, Sigma Chemical Company) at a final dilution of 1:15,000 for ET-1 RIA, or 1:5000 for the ET-3 RIA system. This antiserum cross-reacted 100%, <50%, <50%, and <50% with ET-1, human ET-2, human ET-3, and bovine ET-3, respectively, with the ET-1 RIA system (Fig. 1A), and 100% with these 3 peptides with the ET-3 RIA system (Fig. 1B). The ET-1 RIA system showed that anti-endothelin-1 antiserum had the greatest affinity for ET-1, as shown by the greatest reduction in the amount of labeled ET-1 bound to antibody. Therefore, the concentrations measured with the ET-1 RIA system were termed ET-1 concentrations. The ET-3 RIA system showed that anti-ET-1 antiserum had almost equal affinity for ET-1, ET-2, and bovine ET-3. Therefore, the concentrations measured with the ET-3 RIA system were termed ET concentrations. Standard concentrations of ET-1 and ET-3 ranged from 0.06ng/mL to 1μg/mL. Standards and plasma samples were incubated with antibodies diluted in assay buffer (0.05M phosphate, 0.9% saline, 0.025M EDTA, 0.08% sodium azide, 1% BSA, pH 7.4) and relevant 125I-labeled ET (10,000 counts per minute/100μL assay buffer containing 1% carrier serum). The tubes were incubated for 24h at 4°C. The next day, the second antibody was added to the tubes, and the tubes were incubated for 30min at 4°C. After incubation, bound and free antigens were separated by centrifugation for 30min at 1870×g and 4°C, and radioactivity in the pellets was determined using a gamma counter (ARC-1000, Aloka, Japan). Sensitivities, average intra-assay CVs, and average recovery rates of 3 known amounts (0.97, 1.95, and 3.9ng/mL for ET-1 RIA and 1.95, 3.9, and 7.8ng/mL for ET-3 RIA) of added hormones were 6pg/tube, 2.9%, and 105%, and 6pg/tube, 7.9%, and 95% for the ET-1 and the ET-3 RIA system, respectively. Interassay CVs for the ET-1 and ET-3 assays are not available, as these assays were carried out only once. 2.4.2. Radioimmunoassay for secretin Plasma secretin was measured by the in-house RIA system using rabbit antiserum against mcKLH-[Cys-3]-bovine secretin (4-26) amide at a final dilution of 1:60,000. This antiserum cross-reacted 100% with bovine and human secretin, but did not cross-react with glucagon, oxyntomodulin, GLP-1, and GLP-2 (Fig. 2). Standard concentrations of secretin ranged from 0.015ng/mL to 1μg/mL. Standards and plasma samples were incubated with antibodies diluted in assay buffer (0.2M glycine, 0.03M EDTA, 0.08% sodium azide, and 1% BSA, pH 8.8) and 125I-labeled-secretin (10,000 counts per minute/100μL assay buffer containing 1% carrier serum). The tubes were incubated for 24h at 4°C. The procedures being performed on the next day were the same as in the ET RIA system, as described above. Sensitivity, average intra-assay CV, and average recovery rate of 3 known amounts (1.95, 3.9, 7.8ng/mL) of added hormones were 80pg/mL, 8.9%, and 105%, respectively. The interassay CV for the secretin assay is not available, as this assay was carried out only once. Plasma samples taken at 5min after the secretin injection were measured by diluting the samples to demonstrate that purified bovine secretin, that is, the standard, and secretin in bovine plasma have similar immunological properties. The shapes of the displacement curves of diluted plasma and the standard were similar (Fig. 2). It has been reported that the elimination of plasma interference using extraction procedures before the assay results in the most sensitive secretin assay [40]. In our secretin RIA system, plasma secretin can be measured without extraction with a sensitivity of 80pg/mL. It is worth mentioning that the methods of antibody production and preparation of radioactive tracers are important for a sensitive RIA, as reported by Trenkle [41]. 2.4.3. Radioimmunoassay for ghrelin, GH, and insulin Plasma acyl ghrelin, total ghrelin, GH, and insulin concentrations were measured as previously described [4]. Sensitivities, average intra-assay CVs, and average recovery rates of 3 known amounts of added hormones were 42.08pg/mL, 8.42%, and 94.8% for acyl ghrelin RIA; 0.17ng/mL, 4.48%, and 106% for total ghrelin RIA; 0.03ng/mL, 7.8%, and 112% for GH RIA; and 0.017ng/mL, 7.3%, and 107% for insulin RIA, respectively. Average interassay CVs of ghrelin assays were 9.53% and 4.54% for acyl ghrelin and total ghrelin RIA, respectively. Interassay CVs for GH and insulin assays are not available, as both these assays were carried out only once. 2.4.4. Radioimmunoassay for glucagon The procedures for the glucagon assay were the same as for the secretin RIA system mentioned above, except for using human glucagon as the cold standard, 125I-labeled human glucagon as the tracer, and guinea pig antiserum against mcKLH-[Cys-18]-bovine glucagon (19-29) at a final dilution of 1:5000. This antiserum cross-reacted 100% with human glucagon and bovine glucagon antagonist and <0.4% with bovine oxyntomodulin and GLP-1 when these 2 hormones were at very high concentrations (Fig. 3). Sensitivity, average intra-assay CV, and average recovery rate of 3 known amounts (1.95, 3.9, and 7.8ng/mL) of added hormones were 80pg/mL, 8.9%, and 105%, respectively. Interassay CV for the glucagon assay is not available, as this assay was carried out only once. 2.5. Statistical analysis Data were analyzed by using a MIXED procedure of SAS, version 9.1 (SAS Institute, Inc., Cary, NC, USA), with steer as the error term, hormone or metabolite concentrations as the repeated variable, and day and time of sampling within treatment as the fixed effects. A P value of <0.05 was regarded as statistically significant. Data are presented as mean±SEM of 6 animals. 3. Results Alterations in concentrations of endothelin after ET-1 and ET-3 injection, and secretin after secretin injection are shown in Fig. 4. The basal ET-1 concentration was 0.42±0.03ng/mL. Concentrations of ET-1 increased to 3.8±0.56ng/mL at 5min after ET-1 injection (P<0.01). The concentrations declined at 10min and returned nearly to the basal concentration 10min after ET-1 injection (Fig. 4A). The basal concentration of ET before ET-3 injection was 2.49±0.13ng/mL. Concentrations of ET increased to 11.31±2.75ng/mL at 5min after ET-3 injection (P<0.01). The concentrations declined at 10min and returned nearly to the baseline value 10min after ET-3 injection (Fig. 4B). Secretin concentrations before injection of secretin or vehicle were not different (0.25±0.01ng/mL; Fig. 4C). Concentrations increased to 160.00±5.56ng/mL at 5min after secretin injection (P<0.01). Concentrations declined at 10min and returned to the baseline value 20min after injection. Plasma ghrelin, GH, ETs, secretin, insulin, glucagon, glucose, and NEFA concentrations were not affected by vehicle injection throughout the experimental periods. 3.1. Effect of ET-1 injection on ghrelin, GH, insulin, glucagon, and metabolites The average pre-injection concentrations of acyl ghrelin and total ghrelin among ET-1–, ET-3–, secretin-, and vehicle-injected groups were not different (acyl ghrelin: 245±15pg/mL; total ghrelin: 1.61±0.05ng/mL). Concentrations of acyl ghrelin increased to the peak concentrations of 311±15pg/mL at 10min after ET-1 injection (P<0.01) and remained elevated (271±9pg/mL) at 15min after ET-1 injection (P<0.01, Fig. 5A). Acyl ghrelin concentrations in the ET-1–injected and vehicle-injected groups were different at 10 (P<0.01) and 15min (P<0.05). Plasma total ghrelin concentrations increased to 1.91±0.1ng/mL at 5min after ET-1 injection (P<0.01), and peak concentrations (2.4±0.2ng/mL) occured at 10min (P<0.01). The concentrations remained elevated (1.9±0.1ng/mL) at 15min after ET-1 injection (P<0.05, Fig. 5B). Total ghrelin concentrations in the ET-1–injected and vehicle-injected groups were different at 5, 10 (P<0.01), and 15min (P<0.05). The average pre-injection concentrations of GH among the ET-1–, ET-3–, and vehicle-injected groups were not different (1.06±0.07ng/mL). Plasma GH concentrations increased to 4.73±0.92ng/mL (P<0.01) at 30min after ET-1 injection. The concentrations remained elevated (3.58±0.63ng/mL) at 45min after ET-1 injection (P<0.01; Fig. 5C). Concentrations of GH in the ET-1–injected and vehicle-injected groups were different at 30, 45 (P<0.01), and 60min (P<0.05). Injection of ET-1 did not change plasma insulin and glucagon concentrations throughout the experimental periods (insulin: 0.48±0.05ng/mL (Fig. 5D); glucagon: 2.17±0.16ng/mL; Fig. 5E). Glucose concentrations increased from the basal concentration (98.18±1.79mg/dL) to 120.62±5.07mg/dL at 10min after ET-1 injection, and the increased concentrations lasted for about 2h (P<0.01; Fig. 5F). Glucose concentrations in the ET-1–injected and vehicle-injected groups were different from 15-90min (P<0.05). Concentrations of NEFA increased from the basal concentration (64.47±3.69μEq/L) to 108.26±13.86μEq/L at 15min after ET-1 injection, and the increased concentrations lasted until 45min after ET-1 injection (P<0.05; Fig. 5G). Concentrations of NEFA in the ET-1–injected and vehicle-injected groups were different from 15-45min (P<0.05). 3.2. Effect of ET-3 injection on ghrelin, GH, insulin, glucagon, and metabolites Concentrations of acyl ghrelin increased to the peak concentrations of 337±27pg/mL at 10min after ET-3 injection (P<0.01). The concentrations were still elevated (325±28pg/mL) by 15min after ET-3 injection (P<0.05; Fig. 6A). Acyl ghrelin concentrations in the ET-3–injected and vehicle-injected groups were different at 5, 10, and 15min (P<0.01). Plasma total ghrelin concentrations increased to 2.6±0.1ng/mL at 10min after ET-3 injection (P<0.01). The concentrations remained elevated (2.41±0.27ng/mL) at 15min after ET-3 injection (P<0.01; Fig. 6B). Total ghrelin concentrations in the ET-3–injected and vehicle-injected groups were different at 5 (P<0.05), 10, and 15min (P<0.01). Plasma GH concentrations increased to 2.71±0.30ng/mL at 15min (P<0.05), to 5.42±0.85ng/mL at 20min (P<0.01), and to the peak concentration of 5.56±0.97ng/mL at 30min after ET-3 injection (P<0.01; Fig. 6C). Growth hormone concentrations in the ET-3–injected and vehicle-injected groups were different at 15, 20, and 30min (P<0.01). Injection of ET-3 did not change plasma insulin and glucagon concentrations throughout the experimental periods (insulin: 0.52±0.02ng/mL [Fig. 6D]; glucagon: 2.00±0.14ng/mL [Fig. 6E]). Glucose concentrations initially increased to 131.16±4.05mg/dL at 10min after ET-3 injection, and the increased concentrations lasted until 30min after ET-3 injection (P<0.05; Fig. 6F). Glucose concentrations in the ET-3–injected and vehicle-injected groups were different from 10-45min (P<0.01). Concentrations of NEFA increased to 136.82±11.47μEq/L at 15min after ET-3 injection, and the increased concentrations lasted until 30min after ET-3 injection (P<0.05; Fig. 6G). Concentrations of NEFA in the ET-3–injected and vehicle-injected groups were different from 5-30min (P<0.05). 4. Discussion This study has clearly shown that injection of both ET-1 and ET-3 increased the concentrations of both acyl and total ghrelin at 10min after injection of each peptide. Ghrelin concentrations remained increased until 15min after injection. Our observations that exogenous ET-1 and ET-3 stimulate ghrelin secretion are in agreement with the previous report of de la Cour et al, in which local microinfusion of human ET-1 raised the concentration of rat microdialysate ghrelin 3-fold [8]. In this study, we observed that secretin injection did not change plasma ghrelin and GH concentrations. Our observation that secretin has no effect on ghrelin secretion in cattle does not support their report. It could be considered that the different effects of secretin on ghrelin secretion might be owing to differences in experimental animals and conditions; physiological stages; or route of administration, such as local or peripheral, or acute or chronic administration. According to our results, GH concentrations had significantly increased at 15, 20, and 30min after ET-3 injection, and at 30min after ET-1 injection. It can be considered that the endogenous ghrelin released in response to ET-1 or ET-3 injection stimulates the anterior pituitary gland to secrete GH. From our results, plasma GH release after ET-1 injection, when compared with ET-3 injection, was relatively slower and weaker. It has been described that in rat glomerulus and mesangial cells, ET-3 stimulates cyclic guanosine monophosphate (cGMP) production via the ETB receptor [42]. In accordance with the GH releasing activity, ghrelin stimulates GH release via activation of the nitric oxide/cGMP signaling pathway [43]. From these reports, it can be considered that the relatively stronger response of GH to ET-3 injection may be owing to the activation of cGMP generation in the same cell by ET-3 and ghrelin. Moreover, it is generally accepted that if 2 endothelin receptors, namely, ETA and ETB, coexist in the same cell, one ET receptor competes with another ET receptor to bind with ET-1 or ET-3, leading to the weakened effect of ligand–receptor binding. This general concept could be considered as a possible factor for weakened time-course response of GH to ET-1 injection. Our present data showed that both ET-1 and ET-3 increased plasma glucose and NEFA concentrations without changing insulin and glucagon concentrations. Several studies showed the stimulatory effect of ETs on glucose concentration in humans, goats, and rats [30], [36], [44], [45], [46], [47]. In accordance with the effects of ETs to increase glucose release, some studies have shown that ETs reduced insulin-dependent glucose uptake in skeletal muscle [44] or stimulated glycogenolysis [30], [45], [46], or possibly interfered with normal insulin activity, thereby producing an insulin-resistant condition [47], or ETs inhibited glucokinase activity in isolated rat hepatocytes [48]. In addition, Juan et al. [47] reported that the hyperglycemic reaction induced by intraperitoneal injection of ET-1 in rats lasted for at least 3h, and plasma insulin concentrations were not affected by a low dose (5μg/kg) of ET-1, although these concentrations had decreased at 2 and 3h after a high dose (10μg/kg) of ET-1. From these results and ours, it can be concluded that ET-1 and ET-3 stimulated glucose release by inducing a normal insulin-resistant condition and inhibiting glucokinase activity in cattle. The previous in vitro study in which ET-1 elicited adrenaline and noradrenaline release by bovine adrenal chromaffin cells [49] and the in vivo study in which adrenaline stimulated glucose production and plasma FFA concentrations [50] seem to support the hypothesis that our results of relatively long-lasting responses of plasma glucose and NEFA to ET-1 and ET-3 injection may be owing to both direct and indirect effects on the liver cells. In conclusion, our study has shown that ET-1 and ET-3 stimulate ghrelin and GH secretion in cattle and demonstrated for the first time that endogenous ghrelin released in response to endothelin injection stimulates GH secretion in vivo in cattle. As endothelin stimulates ghrelin and GH secretion, it is suggested that endothelin can contribute, as an alternative for exogenous ghrelin administration, to enhancing GH concentrations in animals. 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Department of Life Science and Agriculture, Obihiro University of Agriculture and Veterinary Medicine, Inada, Obihiro 080-8555, Japan  Corresponding author at: Department of Life Science and Agriculture, Obihiro University of Agriculture and Veterinary Medicine, Nishi 2-11, Inada, Obihiro 080-8555, Japan. Tel.: +81 155 49 5434; fax: +81 155 49 5434.
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