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Department of Pharmacology (M.H., T.A., Y.F., J.B., A.T.), National Research Institute for Child Health and Development, Setagaya-ku, Tokyo 157-8535, Japan; The Chemo-Sero-Therapeutic Research Institute (Kaketsuken) (R.T.), Kumamoto-shi, Kumamoto 860-8568, Japan; Department of Pediatrics (K.K., F.E.), Graduate School of Medical Sciences, Kumamoto University, Kumamoto-shi, Kumamoto 860-8556, Japan; and Department of Health Science (Y.S.), Faculty of Medical Sciences, University of Fukui, Fukui, 910-1193, Japan
Address all correspondence and requests for reprints to: Akito Tanoue, Department of Pharmacology, National Research Institute for Child Health and Development, 2-10-1 Okura, Setagaya-ku, Tokyo 157-8535, Japan. E-mail: atanoue{at}nch.go.jp.
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ABSTRACT |
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INTRODUCTION |
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There is increasing evidence to suggest that AVP plays a crucial role in the regulation of the blood levels of glucose and other energy substrates. Previous work has implicated the metabolic effects of AVP (2). AVP infusions are associated with increases in circulating glucose levels (3, 4). This effect is supposed to be due to two distinct actions of this hormone. AVP stimulates the glucagon release from pancreatic islet cells incubated in vitro (5), especially via the V1b receptor (6). The effect of AVP on glucagon secretion occurs without apparent changes in insulin secretion. In addition, AVP can act directly in the liver to stimulate glucose production. In hepatocytes, AVP interacts with specific V1 receptor sites (7) and promotes glycogenolysis and gluconeogenesis (8). These actions of AVP in the liver are distinct from those of glucagon and are mediated by a calcium-dependent pathway (9), possibly via the V1a receptor.
Further involvement of AVP in regulating the supply of energy substrates has been suggested by the finding that AVP increases the taurocholate efflux from hepatocytes (10). Thus, AVP is involved in the lipid metabolism through the regulation of hepatic bile salt secretion. Rofe and Williamson (3) found that AVP infusions decreased circulating ketone bodies in starved rats, suggesting that AVP has an antilipolytic action on adipocytes. The understanding of the physiological importance, as well as the complex nature, of AVP involvement in regulating blood glucose levels and supplying energy substrates requires further study.
Recently, we generated V1a receptor-deficient (V1aR–/–) mice, which are not lethal and have no apparent anatomical anomaly but exhibit an impairment of the spatial memory in an eight-arm radial maze (11). Further research demonstrated that V1aR–/– mice have a significantly lower basal blood pressure with a 9% reduced circulating body blood volume but not a significant alteration in cardiac functions (12). Because this V1aR–/– mouse was considered to be a good model for the analysis of the AVP-inducible lipid metabolism, we investigated the effects of V1a receptor deficiency on the lipid metabolism.
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RESULTS |
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), whereas the glycerol level of V1aR–/– mice was slightly higher than that of V1aR+/+ mice under the fasting condition (173.6 ± 15.15 µM in V1aR+/+, n = 4; and 228.3 ± 39.2 µM in V1aR–/– mice, n = 6) (Fig. 1B). Thus, the increased level of blood glycerol in V1aR–/– mice was more evident under the feeding condition, indicating that lipolysis is enhanced in V1aR+/+ mice as well as in V1aR–/– mice under the fasting condition.
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, A–F). Under the feeding condition, the serum levels of acetoacetic acid, 3-hydroxybutyric acid, and total ketone bodies were not significantly different in V1aR+/+ and V1aR–/– mice (acetoacetic acid: 13.1 ± 1.05 µmol/liter in V1aR+/+, n = 11, and 10.0 ± 1.22 µmol/liter in V1aR–/– mice, n = 11; 3-hydroxybutyric acid: 128.6 ± 12.21 µmol/liter in V1aR+/+, n = 12, and 99.8 ± 20.39 µmol/liter in V1aR–/– mice, n = 11; total ketone bodies: 142.4 ± 10.86 µmol/liter in V1aR+/+, n = 12, and 109.8 ± 21.38 µmol/liter in V1aR–/– mice, n = 11) (Fig. 2, A–C). On the other hand, the serum levels of acetoacetic acid, 3-hydroxybutyric acid, and total ketone bodies were significantly higher in V1aR–/– mice than in V1aR+/+ mice under the fasting condition (acetoacetic acid: 85.3 ± 16.54 µmol/liter in V1aR+/+, n = 4, and 272.0 ± 46.11 µmol/liter in V1aR–/– mice, n = 5, P = 0.0108; 3-hydroxybutyric acid: 652.5 ± 115.0 µmol/liter in V1aR+/+, n = 4, and 2256.0 ± 430.11 µmol/liter in V1aR–/– mice, n = 5, P = 0.0147; total ketone bodies: 737.7 ± 129.72 µmol/liter in V1aR+/+, n = 4, and 2528 ± 473.65 µmol/liter in V1aR–/– mice, n = 5) (Fig. 2, D–F).
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) because glycerol and ketone bodies were produced by the TG or FFA catabolism. The levels of TG and FFA of V1aR–/– mice were significantly lower than those of V1aR+/+ mice (TG: 57.3 ± 2.17 mg/dl in V1aR–/–, n = 8, and 82.1 ± 7.41 mg/dl in V1aR+/+, n = 8, P = 0.0074; FFA: 0.42 ± 0.027 mEq/liter in V1aR–/–, n = 12, and 0.57 ± 0.03 mEq/liter in V1aR+/+, n = 12, P = 0.00088), indicating that those catabolisms were enhanced in V1aR–/– mice.
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). The tissue TG content in V1aR–/– mice was slightly, but not significantly, reduced in the heart and kidney under the fasting condition and in liver and kidney under the feeding condition when compared with TG content in V1aR+/+ mice. This may be because TG is catabolized mostly in adipose tissue, and other tissues, such as the liver, heart, kidney, and muscle have minor involvement in the TG catabolism.
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). The analysis with tandem mass spectrometry revealed that the level of free carnitine in V1aR–/– mice was significantly higher than that of V1aR+/+ mice under feeding and fasting conditions and that the free carnitine levels were constant under both feeding and fasting conditions. 5-Carbon acylcarnitine (C5) and hydroxy-5-carbon acylcarnitine (C5-OH) were lower in V1aR–/– mice than in V1aR+/+ mice under both feeding and fasting conditions, in correlation with the decreased level of the branched-chain amino acids (BCAAs) isoleucine, leucine, and valine. The C4 level was not at all different in V1aR+/+ and V1aR–/– mice and was reduced in both mice by fasting. Other acylcarnitine levels of V1aR–/– mice were higher than those of V1aR+/+, especially in the case of C2, C10:1, C10, C14:1, C16, C18:1, and OH-C18:1, under feeding conditions. After 24 h of fasting, the variations of those acylcarnitines in V1aR+/+ and V1aR–/– mice were smaller, suggesting that the lipid metabolism in V1aR+/+ mice was also enhanced by fasting.
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). Tissue carnitine and acetylcarnitine were detected by tandem mass spectrometry. Acetylcarnitine was shown as a representative, since the levels of other acylcarnitines were greatly lower than acetylcarnitine. The carnitine level in the liver of V1aR–/– mice, which is a unique tissue for the production of carnitine, was largely increased under both feeding and fasting conditions, indicating that carnitine production was increased in V1aR–/– mice.
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Expressions of AVP Receptors
To assess the role of AVP receptors, including the V1a receptor, in the lipid metabolism, we examined those expressions by the RT-PCR method with RNA from tissues [heart, liver, kidney, brown adipose tissue (BAT), and white adipose tissue (WAT)] that are related with the lipid metabolism. The V1a receptor was expressed in all tissues examined, whereas the V1b and V2 receptors were only detected in WAT and in the kidney, respectively (Fig. 4).
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). Under this condition, more than 0.5 µM of isoproterenol stimulated lipolysis in a dose-dependent manner. Furthermore, isoproterenol-induced lipolysis was greatly promoted in the adipocytes of V1aR–/– mice (approximately 3-fold) (Fig. 5).
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). This result is correlated with an observation in the lipolysis reaction in response to isoproterenol (Fig. 5
). Next, we investigated the phosphorylation state of Akt, which is a key molecule in insulin signaling, because insulin signaling suppressed isoproterenol-induced lipolysis via the reduction of cAMP by the activation of phosphodiesterase 3B (PDE) (14). The differentiated adipocytes were stimulated by 50 nM insulin for 5 min. Akt was phosphorylated in response to insulin, but the phosphorylation level of Akt in adipocytes of V1aR–/– mice was largely reduced (Fig. 7).
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). The liver weight of V1aR–/– mice was significantly smaller than that of V1aR+/+ mice, whereas the cholecyst weight of V1aR–/– was significantly bigger than that of V1aR+/+ mice (liver weight: 1293.3 ± 70.85 mg in V1aR–/–, n = 8, and 1016.4 ± 62.8 mg in V1aR+/+, n = 7, P = 0.0127; cholecyst weight: 20.67 ± 2.82 mg in V1aR–/–, n = 4, and 38.3 ± 3.49 mg in V1aR+/+, n = 6, P = 0.007). The cholesterol concentration of V1aR–/– mice was higher than that of V1aR+/+ mice (99.64 ± 7.57 mg/dl in V1aR–/–, n = 7, and 81.77 ± 2.69 mg/dl in V1aR+/+, n = 6, P = 0.061). The taurine concentration of V1aR–/– mice was also higher than that of V1aR+/+ mice (768.8 ± 52.8 nmol/ml in V1aR–/–, n = 5; and 608.6 ± 28.2 nmol/ml in V1aR+/+, n = 5, P = 0.028) under feeding conditions. The total bile acid level of V1aR–/– mice was 10.25 ± 6.59 µmol/liter (n = 4), but in V1aR+/+ mice it was undetectable (less than 3 µmol/liter, n = 3).
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DISCUSSION |
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). Ketone bodies were increased in V1aR–/– mice under the fasting condition (Fig. 2), corresponding to the finding that AVP decreased the circulating ketone bodies (3). The increase of glycerol and ketone bodies in V1aR–/– mice indicates that lipolysis is promoted under a condition of abolished AVP signaling via the V1a receptor. The increased circulating glycerol could be due to increased production, which results from the increased activity of tissue lipase. Lipase mediates a breakdown of TG into glycerol and FFA. The FFA then serves as a precursor of ketoacids (19, 20). Therefore, we investigated whether the catabolisms of TG and FFA were promoted in V1aR–/– mice. The levels of TG and FFA in blood were significantly decreased in V1aR–/– mice, suggesting that the catabolisms of TG and FFA in V1aR–/– mice were enhanced, in good correspondence with the increased levels of glycerol. The tissue TG was slightly reduced in the heart and kidney under the fasting condition and in the liver and kidney under the feeding condition (Table 1), suggesting that the catabolisms of TG and FFA in V1aR–/– mice were enhanced.
Next, we examined the levels of carnitine and acylcarnitines in the blood of V1aR+/+ and V1aR–/– mice to assess the status of ß-oxidation. Our results revealed that carnitine and most acylcarnitines were increased in V1aR–/– mice under both feeding and fasting conditions, indicating that the ß-oxidation of lipids was remarkably enhanced under the feeding condition (Table 2). However, the levels of the ketone bodies of V1aR–/– mice were not increased under the feeding condition (Fig. 2). To elucidate the discrepancy, acylcarnitine in tissues was assessed. In the liver, tissue acylcarnitine was decreased under the feeding condition but increased under the fasting condition. Furthermore, acylcarnitine in muscle was increased under both conditions, but particularly under the feeding condition. Based on these results, although the change in circulating lipids could be due to clearance by the heart and muscle, ß-oxidation could be promoted in muscles under both conditions, but particularly under the feeding condition, and promoted in the liver under the fasting condition because ketone bodies were produced in the liver but not in muscle (21).
Furthermore, the decreases in C5 and C5-OH levels were well correlated with the low levels of the BCAAs, valine, leucine, and isoleucine. The decrease of those BCAAs, which metabolize in muscle, could imply the hypermetabolism of amino acids in muscle (data not shown). Thus, the level of odd-chain acylcarnitine (representing intermediates in the BCAA catabolism), such as C5, is lower because it reflects a switch in substrate preference in favor of fatty acid over BCAA rather than hypermetabolism. An altered acylcarnitine metabolism has been reported in the inherited deficiency of enzyme activities in ß-oxidation, such as medium-chain acyl-CoA dehydrogenase deficiency (22, 23, 24, 25, 26, 27), very-long-chain acyl-CoA dehydrogenase deficiency (28), long-chain hydroxyacyl-CoA dehydrogenase deficiency (29), and systemic carnitine deficiency (28, 30, 31, 32). However, the acylcarnitine profile of V1aR–/– mice was different from the profiles of disorders including deficiencies in these enzymes. Instead, the promotion of lipolysis and ß-oxidation is observed in diabetes because insulin suppresses the catabolism of fat in fat tissue by inhibiting hormone-sensitive lipase in health.
To consider the potential roles of AVP receptors (V1b and V2) other than the V1a receptor in the lipid metabolism, we examined those expressions in the heart, liver, kidney, skeletal muscle, BAT, and WAT. The V1a receptor was expressed in all tissues examined, but the V1b receptor was expressed in WAT, and the V2 receptor was expressed only in the kidney. These results indicate that the V1a receptor is likely to work in the lipid catabolism because the V1b receptor is hardly expressed in tissues in which it carries out ß-oxidation. The V2 receptor is involved in antidiuresis in the kidney. Using the precursor cells of BAT, we investigated whether isoproterenol-induced lipolysis was altered in V1aR–/– mice because isoproterenol stimulated lipolysis and AVP inhibited isoproterenol-induced lipolysis in vitro (13). Isoproterenol-induced lipolysis was greatly promoted in the adipocytes of V1aR–/– mice (approximately 3-fold) (Fig. 5). It is known that isoproterenol induces lipolysis via Gs-signaling involving adenylate cyclase, cAMP, protein kinase A, and hormone-sensitive lipase (13). Therefore, it was investigated whether the increase of cAMP was enhanced in the adipocytes of V1aR–/– mice in response to isoproterenol. The cAMP was significantly increased in the adipocytes of V1aR–/– mice, suggesting that Gs-signaling was promoted in the adipocytes of V1aR–/– mice. Insulin is the most potent antilipolytic hormone (33), and it suppresses lipolytic signaling via the reduction of cAMP by PDE 3B activation (14). Furthermore, AVP inhibited forskolin- and isoproterenol-induced lipolysis (13, 18) and activated insulin signaling, such as Akt and p70S6kinase, via the EGF receptor-phosphatidylinositol 3-kinase pathway (34), suggesting that AVP can modulate the lipolysis by affecting insulin signaling. To investigate whether insulin signaling was suppressed in the adipocytes of V1aR–/– mice, Akt phosphorylation after insulin stimulation was examined. The phosphorylation of Akt was significantly reduced in the adipocytes of V1aR–/– mice, suggesting that AVP modulated lipolysis via the AVP V1a receptor even in the absence of AVP stimulation.
AVP has been determined to regulate the bile flow in the rat on the basis of its capacity to generate intercellular calcium waves, which are spatially oriented along the acinus axis by a V1a receptor gradient (35, 36). We analyzed the blood bile acid concentrations in V1aR+/+ and V1aR–/– mice. The total bile acid was detected in V1aR–/– mice but not in V1aR+/+ mice (Table 4). This suggested that reabsorption, secretion, or production of bile acid or cholestasis, which is usually accompanied with a high level of bilirubin, could be enhanced in V1aR–/– mice. Based on the finding that AVP promotes the secretion of bile acid and that the bilirubin level is not different in V1aR+/+ and V1aR–/– mice (data not shown), the enhanced secretion of bile acid and cholestasis should be excluded as a cause of the high level of bile acid. Thus, the reabsorption or production of bile acid could be enhanced, resulting in a high level of circulating bile acid in V1aR–/– mice. Furthermore, the circulating bile acid levels were correlated with the size of the cholecyst of V1aR–/– mice, which was significantly larger than that of V1aR+/+ mice (Table 4).
To investigate the reasons for the increased circulating bile acid and the enlargement of the cholecyst in V1aR–/– mice, the levels of cholesterol and taurine were compared between V1aR+/+ and V1aR–/– mice because cholesterol is conjugated with either glycine or taurine to give glycocholate or taurocholate (bile acids), respectively. The levels of cholesterol and taurine in V1aR–/– mice were higher than those in V1aR+/+ mice (Table 4). Thus, the increased level of cholesterol and taurine could explain the reason for the increased level of bile acid. However, it is unclear why the level of taurine was increased in V1aR–/– mice. On the other hand, it has been reported that taurine has a hypocholesterolemic action in rats fed a high-cholesterol diet (37). This raises the question of why blood cholesterol increased despite the increased taurine level in V1aR–/– mice. Acetyl-coenzyme A (CoA) is synthesized in mitochondria by fatty acid oxidation. Acetylcarnitine is synthesized by carnitine O-acetyltransferase reaction (EC 2.3.1.7), which reversibly converts acetyl-CoA and free carnitine to acetylcarnitine and sulfhydryl-CoA without ATP utilization and is then transported to the cytosolic compartment as acetylcarnitine in both liver (38, 39) and heart myocytes (40). Cholesterol can be synthesized from cytosolic acetylcarnitine (41). Thus, a possible reason is that cholesterol production from acetylcarnitine was also increased by the promoted ß-oxidation of fatty acid.
In the mechanisms of AVP-induced antilipolysis, two processes are likely to be inhibited. The administration of (lysine-) AVP to healthy subjects was followed by a significant decrease in the plasma nonesterified fatty acid level by inhibiting hormone-sensitive lipase in the adipose tissue (42). Furthermore, the administration of AVP in starved rats decreased in plasma nonesterified fatty acids (3). In vitro, AVP inhibits forskolin-induced lipolysis in human adipocytes by increasing intracellular [Ca2+] in a dose-dependent manner (43), as measured by the glycerol release into the culture medium (18). These findings indicate that AVP first inhibits the process of lipid metabolism by hormone-sensitive lipase. Secondly, the findings that AVP inhibits ketogenesis from oleate and stimulates the esterification and oxidation of this fatty acid (44) demonstrate the inhibition of ß-oxidation of fatty acid by AVP. Thus, AVP is likely to act negatively on two processes in lipid metabolism.
AVP plays a crucial role in the regulation of the blood levels of glucose and other energy substrates. For example, AVP infusions are associated with increases in circulating glucose levels (3, 4). AVP stimulates glucagon release from pancreatic islet cells incubated in vitro (5), especially, via the V1b receptor (6). AVP promotes glycogenolysis and gluconeogenesis (8). Furthermore, we showed that AVP had an antiproteolytic action by showing that proteolysis was promoted in V1aR–/– mice with hyperammonemia (Hiroyama, M., manuscript in preparation).
In conclusion, our results showed that the lipid metabolism producing ketone bodies via the metabolisms of TG and FFAs and ß-oxidation was enhanced in V1aR–/– mice. Furthermore, it was verified that AVP could modulate the lipid metabolism by the antilipolytic action and the synthesis of bile acid via the V1a receptor using V1aR–/– mice. The elucidation of the regulation mechanism of AVP-induced lipolysis inhibition may contribute to the establishment of a therapy for obesity. The involvement of this receptor in the glucose and protein metabolism and weight gain is now under investigation. Our findings from this study suggest that specific drugs for the V1a receptor may represent an effective therapy for the metabolic syndrome.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Blood Glycerol Measurement
To measure the glycerol concentration of blood, the serum was prepared from the inferior vena cava. Twenty microliters of serum were diluted with 30 µl H2O, and then 50 µl glycerol reagent (Zen-Bio, Inc., Research Triangle Park, NC) was added. Fifteen minutes later, the absorbances were measured at 540 nm.
Tissue TG Measurement
Tissues were perfused with PBS (–) from the left ventricle before dissection to reduce TG contamination from blood. The liver, heart, skeletal muscle, and kidney were dissected, and the exact mass of the sample was then determined using a microbalance. The tissues were homogenated in 100 µl H2O, and 600 µl chloroform:methanol (1:2) was then added to the homogenates. The homogenates were mixed and placed overnight at room temperature to extract the total lipids. Two hundred microliters of chloroform were added, and then 200 µl 1 N HCl was added to separate the layers by centrifugation. The lower phase was dried and resolved in the appropriate volume of isopropanol. The solutions were used for TG measurement using triglyceride E-test Wako (Wako, Osaka, Japan).
Electrospray Tandem Mass Spectrometry
Electrospray tandem mass spectrometry was carried out for the analysis of acylcarnitines, valine, leucine, and isoleucine in dried blood specimens collected from the tail vein. Three-millimeter discs were punched from the filter paper into a 96-well plate. The internal standards were added to each sample. The extracts were eluted with 120 µl methanol for 30 min and transferred to another 96-well plate for drying. The dried sample was dissolved in butanolic HCl and heated to 65 C for 15 min. The butanolic HCl was removed with a stream of oxygen-free nitrogen, and the dried sample was reconstituted in 150 µl 80% acetonitrile. Ten microliters of the solution were added to the electrospray ionization source of Alliance HT-2795 (Waters Corp., Milford, MA) and Quattro Micro API (Waters Corp.). For the analysis of tissue acylcarnitine and carnitine, approximately 30 mg of the cardiac left ventricle and 60 mg of other tissues (liver, kidney, and muscle) were placed in 1.5 ml Eppendorf vials (Eppendorf, Hamburg, Germany), and the exact mass of the sample was determined using a microbalance. The samples were freeze-dried overnight. The freeze-dried tissues were ground to powder using an Eppendorf micropestle. One milliliter of 80% acetonitrile was added to the extract, and the solution was then incubated for 1 h at room temperature. The solution was centrifuged at 12,000 x g for 5 min, and the supernatant was used for the analysis.
Biochemical Analysis
Free fatty acid, triglyceride, and cholesterol of the serum from the inferior vena cava were measured using NEFA C-test Wako, triglyceride E-test Wako, and cholesterol E-test Wako (Wako), respectively. For measuring the ketone bodies, including acetoacetic acid and 3-hydroxybutyric acid, 300 µl serum was prepared from the abdominal aorta. The total bile acid concentration in serum from the inferior vena cava was measured by the 3
-HSD /
4-DH method. The taurine concentration in serum from the inferior vena cava was measured by the HPLC method.
RT-PCR
The RNAs were purified from the heart, liver, kidney, muscle, BAT, and WAT using ISOGENE (Nippon Gene, Tokyo, Japan). The RNAs were reverse-transcribed using Superscript III (Invitrogen, Carlsbad, CA). The amplification reactions were conducted on a PCR thermal cycler (Takara, Shiga, Japan) at 95 C for 30 sec, 58 C for 1 min, and 72 C for 2 min for 25 cycles using specific primer sets: glyceraldehyde 3-phosphate dehydrogenase, 5'-GGTCATCATCTCCGCCCCTTC-3' and 5'-CCACCACCCTGTTGCTGTAG-3'; V1a receptor, 5'-TACATGCTGGTGGTGATGACGGCTG-3' and 5'-ATTGCGCCAGATGTGGTAACAGATG-3'; V1b receptor, 5'-CCGGAATTCCACCTGTTGCCACGTTCCC-3' and 5'-CCGCTCGAGAGAGATGCTGGTCTCCATAG-3'; V2 receptor, 5'- CAGCGAGGGAGCCCATGTAT-3' and 5'-GCTAAGCTACAATGGGTTT-3'. The products were detected under UV illumination.
Brown Fat Precursor Cell Isolation and Differentiation and Lipolysis Assay
Brown fat precursor cells were isolated from newborn V1aR+/+ and V1aR–/– mice (d 4) by collagenase digestion. Briefly, the interscapular BAT depots were dissected under sterile conditions, minced, and transferred to a Krebs-Ringer buffer (120 mM NaCl, 1.27 mM CaCl2, 1.2 mM MgCl2, 4.75 mM KCl, and 1.2 mM KH2PO4) containing 15 mM sodium bicarbonate, 10 mM HEPES, 2 mM sodium pyruvate, and 1% BSA (pH 7.4). Collagenase S-I (Nitta Gelatin, Japan) was added at 2 mg/ml. The tissues were shaken at 37OC for 60 min, and the digest was filtered through a 70-µm nylon filter (a BD Falcon cell strainer). The precursor cells were pelleted by centrifugation (80 x g, 5 min) and resuspended in the same buffer without collagenase. The cells were recentrifuged and finally resuspended in 10% fetal bovine serum (FBS)-DMEM. The precursor cells were seeded on a 96-well plate at 1.2–2 x 104/well, and the medium was changed to a fresh medium 20 h later. Adipocyte differentiation was induced by treating confluent cells in an induction medium (10% FBS-DMEM containing 0.5 mM isobutylmethylxanthine, 0.5 µM dexamethazone, 0.125 mM indomethacine, 20 nM insulin, and 1 nM T3) for 6 d, and the lipolysis assay was then carried out in a phenol red free-DMEM. Isoproterenol was added at the indicated concentrations for 5 h, and the glycerol content in the culture medium was measured with a lipolysis assay kit (Zen-Bio, Inc., Research Triangle Park, NC). TG in the cells was extracted using 200 µl chloroform:methanol (1:2) and separated by centrifugation after the addition of 80 µl chloroform and 80 µl 1N-hydrochloric acid. Dried TG was solved in isopropanol, and the TG concentration was then measured using the triglyceride E-test Wako. For oil-red O staining, cells fixed using 10% formalin for 1 h were soaked in 3 mg/ml oil-red O in 60% isopropanol for 1 h and then washed with 60% isopropanol for 2 min. To determine the protein concentration, separate wells were prepared, and the protein concentration was quantified using the BCA protein assay (Pierce).
cAMP Determination in Primary Brown Adipocytes
For the cAMP assay, 3-wk-old mice were used to collect a large number of precursor cells of adipocytes. The cells were isolated from interscapular, axillary, and cervical BAT depots as described above. The precursor cells were seeded on a 12-well plate with 10% FBS-DMEM. After the cells reached confluence, the differentiation was induced with an induction medium for 3 d. The cells were then starved over night with 0.3% BSA-DMEM. The cells were stimulated by isoproterenol at the indicated concentrations for 10 min. To terminate the stimulation, the media was aspirated, and 150 µl of ice-cold 0.1 N HCl was added to each well; then, the plate was placed at room temperature for 20 min. The cells were collected and centrifuged, and the supernatant was used for cAMP determination using a cAMP EIA kit (Cayman Chemical, Ann Arbor, MI). The protein concentration was quantified using a BCA protein assay (Pierce).
Phospho-Akt Assay
The precursor adipocytes were seeded on a 6-well plate and differentiated as described in the cAMP assay. The starved cells were stimulated with 50 nM insulin for 5 min. To terminate the stimulation, the cells were placed on ice and washed with ice-cold PBS (-). The cells were then lysed with 100 µl of a lysis buffer (50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1% Nonidet P-40, 2 mM Na3VO4, 10 mM NaF, 10 mM Na4P2O7, 1 mM EDTA, 1 mM EGTA, 5 µg/ml leupeptin, 5 µg/ml aprotinin, 1 mM phenylmethylsulfonylfluoride). The cell lysates were sonicated for 10 sec, and the protein concentration was quantified using a BCA protein assay (Pierce, Rockford, IL). Twenty micrograms of the cell lysates were applied on 7.5% SDS-PAGE gel and transferred to a polyvinylidene fluoride membrane. Phospho-Akt was detected using a 1:1000 dilution of a rabbit polyclonal antiphospho Akt (The308) and a mouse monoclonal antiphospho Akt (Ser473) (Cell Signaling Technology, Beverly, MA). Total Akt expression was detected using a mouse monoclonal anti-Akt1 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA).
Tissue Weight Measurements
The weights of livers were measured after perfusion of PBS (–) and removal of the cholecyst.
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FOOTNOTES |
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Disclosure Statement: The authors have nothing to disclose.
First Published Online October 4, 2006
Abbreviations: AVP, [Arg8]vasopressin; BAT, brown adipose tissue; BCAA, branched-chain amino acid; FFA, free fatty acid; TG, triacylglycerol; V1aR–/– mice, V1a AVP receptor knockout mice; WAT, white adipose tissue.
Received for publication February 9, 2006. Accepted for publication September 26, 2006.
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-cell line In-R1–G9: involvement of V1b receptors. Life Sci 63:1871–1878[CrossRef][Medline]This article has been cited by other articles:
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 T. Tsunematsu, L.-Y. Fu, A. Yamanaka, K. Ichiki, A. Tanoue, T. Sakurai, and A. N. van den Pol Vasopressin Increases Locomotion through a V1a Receptor in Orexin/Hypocretin Neurons: Implications for Water Homeostasis J. Neurosci., January 2, 2008; 28(1): 228 - 238. [Abstract] [Full Text] [PDF]
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 Y. Fujiwara, M. Hiroyama, A. Sanbe, T. Aoyagi, J.-i. Birumachi, J. Yamauchi, G. Tsujimoto, and A. Tanoue Insulin hypersensitivity in mice lacking the V1b vasopressin receptor J. Physiol., October 1, 2007; 584(1): 235 - 244. [Abstract] [Full Text] [PDF]
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M. Hiroyama, T. Aoyagi, Y. Fujiwara, S. Oshikawa, A. Sanbe, F. Endo, and A. Tanoue Hyperammonaemia in V1a vasopressin receptor knockout mice caused by the promoted proteolysis and reduced intrahepatic blood volume J. Physiol., June 15, 2007; 581(3): 1183 - 1192. [Abstract] [Full Text] [PDF]
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T. Aoyagi, J.-i. Birumachi, M. Hiroyama, Y. Fujiwara, A. Sanbe, J. Yamauchi, and A. Tanoue Alteration of Glucose Homeostasis in V1a Vasopressin Receptor-Deficient Mice Endocrinology, May 1, 2007; 148(5): 2075 - 2084. [Abstract] [Full Text] [PDF]
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