Leptin Improves Insulin Resistance and Hyperglycemia in a Mouse Model of Type 2 Diabetes
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《内分泌学杂志》
Diabetes Branch (Y.T., S.Y., D.L.) and Mouse Metabolic Core Facility (O.G., W.J., S.P.), National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892
Department of Physiology (Z.A., M.W.), University of Toronto, Toronto, Canada M5S 1A8
Abstract
Leptin has metabolic effects on peripheral tissues including muscle, liver, and pancreas, and it has been successfully used to treat lipodystrophic diabetes, a leptin-deficient state. To study whether leptin therapy can be used for treatment of more common cases of type 2 diabetes, we used a mouse model of type 2 diabetes (MKR mice) that show normal leptin levels and are diabetic due to a primary defect in both IGF-I and insulin receptors signaling in skeletal muscle. Here we show that leptin administration to the MKR mice resulted in improvement of diabetes, an effect that was independent of the reduced food intake. The main effect of leptin therapy was enhanced hepatic insulin responsiveness possibly through decreasing gluconeogenesis. In addition, the reduction of lipid stores in liver and muscle induced by enhancing fatty acid oxidation and inhibiting lipogenesis led to an improvement of the lipotoxic condition. Our data suggest that leptin could be a potent antidiabetic drug in cases of type 2 diabetes that are not leptin resistant.
Introduction
LEPTIN IS AN adipocyte-derived hormone that plays a key role in the regulation of food intake and energy expenditure (1, 2). Generally, leptin regulates energy homeostasis via the long isoform of the leptin receptor, which is highly expressed in hypothalamus (3, 4). It also regulates lipid and glucose metabolism and insulin action independently of its effects on food intake (5, 6). Leptin stimulates fatty acid oxidation and glucose uptake in skeletal muscle (7, 8, 9, 10), inhibits glucose output and lipogenesis in liver (11, 12), and inhibits insulin secretion (13). These metabolic effects of leptin are thought to be mainly exerted via the hypothalamus. However, the long isoform of the leptin receptor is also expressed in peripheral tissues (3, 4, 14). Thus, it is possible that leptin could regulate metabolism acting directly on peripheral tissues. However, the underlying mechanisms by which leptin regulates on metabolism and insulin action remain unclear.
Leptin deficiency contributes to the insulin resistance of the ob/ob mice bearing a mutation in the leptin gene (15, 16) and leptin administration to ob/ob mice ameliorates the hyperglycemia, hypertriglyceridemia, and hyperinsulinemia (6, 17, 18). Leptin deficiency is also a primary cause of insulin resistance in mice and humans with lipodystrophy who have reduced amount of adipose tissue. Leptin replacement therapy improves most of the metabolic abnormalities associated with lipodystrophy (19, 20, 21) and transgenic overexpression of leptin gene also improved insulin resistance and diabetes in a mouse model of lipoatrophic diabetes (22). Thus, leptin has been shown to be effective for the treatment of diabetes associated with leptin deficiency. However, leptin-deficient states are extremely rare; the great majority of the patients with type 2 diabetes have either normal or elevated leptin levels (23, 24, 25, 26). Previous studies showed that transgenic overexpression of the leptin gene led to increased insulin sensitivity in normal mice (27) and could delay the onset of impaired glucose metabolism in lethal yellow KKAy mice (28). However, thus far, there is no evidence that leptin improves type 2 diabetes where circulating leptin levels are normal.
We developed a transgenic mouse model overexpressing a dominant-negative IGF-I receptor specifically in skeletal muscle [MKR (transgenic overexpression of a skeletal muscle dominant-negative IGF-I receptor with a lysine-to-arginine amino acid) mice] (29). In MKR mice, the formation of hybrid receptors between defective IGF-I receptor and endogenous IGF-I and insulin receptors led to the abrogation of both IGF-I and insulin receptor signaling pathways in skeletal muscle (29). MKR mice develop an early-onset type 2 diabetes, characterized by dyslipidemia; hyperinsulinemia; and secondary insulin resistance in liver and fat, excessive lipid stores in muscle and liver, and -cell dysfunction (29). They have normal serum leptin levels (30, 31). The MKR mouse is therefore an excellent model to examine whether leptin administration would be effective to treat type 2 diabetes with normal circulating leptin. Here we demonstrate that leptin serves as an antidiabetic agent for the MKR mice.
Materials and Methods
Experimental animals
The generation of the MKR mice (FVB/N background) has been described previously (29). All experiments were performed in homozygous MKR male mice, 6–7 wk of age. As controls, sex- and age-matched wild-type (WT) mice were used. Mice were kept on a 12-h light (0600–1800 h), 12-h dark cycle (1800–0600 h) and were fed NIH-07 rodent chow (Zeigler Brothers Inc., Gardners, PA) and water ad libitum. All studies were conducted in accordance with National Institutes of Health guidelines, as approved by the Animal Care and Use Committee of National Institute of Diabetes and Digestive and Kidney Diseases.
Leptin treatment
For leptin infusion, an Alzet miniosmotic pump (model 2002; Alza Corp., Palo Alto, CA) was implanted sc on the back of each mouse. The pumps delivered saline or 10 μg leptin/d (R&D Systems, Minneapolis, MN) for 14 d. After placement of the pump, the mice were housed individually. At the end of the treatment, mice were killed under anesthesia with Avertin in the nonfasting state between 0900 and 1200 h.
Food intake
Mice were caged individually and treated with or without leptin, as described above. The amount of food in the feeding container was measured at d 14 of treatment. Food intake was expressed as grams per day per mouse.
Pair-feeding study
Mice were treated with or without leptin for 2 wk, as described above. The amount of food consumed by the leptin-treated MKR mice was monitored each day at 1800 h, and that amount was given to the one of the saline-treated MKR mice on the next day.
Histology
The left epididymal fat pad was isolated and fixed overnight in 4% paraformaldehyde in PBS. The tissues were then transferred to 70% ethanol and embedded in paraffin. Samples were cut into 5-μm sections and hematoxylin and eosin staining was performed.
Biochemical assays
Blood was obtained from the tail vein and glucose levels were measured using a Glucometer Elite (Bayer Corp., Elkhart, IN). Serum samples were generally obtained from the tail or retroorbital vein in the nonfasting state between 0900 and 1200 h. Triglycerides (Thermo DMA, Louisville, CO) and free fatty acid (FFA) (Roche Applied Science, Indianapolis, IN) were measured according to the manufacturer’s procedures. Insulin (SRI-13K; Linco Research Inc., St. Charles, MO), leptin (ML-82K; Linco Research), and adiponectin (MADP-60HK; Linco Research) were measured by RIAs.
Determination of body composition
Body composition was measured in nonanesthetized mice using the Bruker minispec NMR analyzer mq10 (Bruker Optics Inc., The Woodlands, TX).
Insulin tolerance and glucose tolerance test
Insulin and glucose tolerance tests were performed with 5- or 7-h fasted mice during daylight time. Human insulin (0.75 U/kg body weight) or glucose (2 g/kg body weight) were injected ip. Blood glucose levels were measured at the indicated time points.
Hyperinsulinemic-euglycemic clamps
The clamp studies were performed as developed by Kim et al. (32, 33). The study involved a primed insulin infusion. Mice were treated with or without leptin for 2 wk, as described above. On d 7 of the treatment (4 d before the clamp experiment), mice were anesthetized with 100 mg/kg ketamine and 10 mg/kg xylazine. A catheter was inserted into a lateral incision on the right side of the neck and advanced into the superior vena cava via the right internal jugular vein. The catheter was then sutured into place, according to the protocol of MacLeod and Shapiro (34). The evening before the clamp analysis, mice were fasted for 10 h. The basal rates of glucose turnover were measured by continuous infusion of [3-3H] glucose (0.02 μCi/min) for 120 min that followed a bolus of 2.5 μCi, starting at 0900 h. Blood samples (20 μl) were taken at 90 min and 115 min of the basal period for the determination of plasma [3-3H] glucose concentration. A 120-min hyperinsulinemic-euglycemic clamp was started at 1100 h. Insulin was infused as a bolus of 300 mU/kg over a period of 3 min, followed by continuous insulin infusion at the rate of 2.5 mU/kg·min (Humulin R, Eli Lilly, Indianapolis, IN) to raise plasma insulin concentration to 4 ng/ml. During the clamp study, mice were restrained and blood samples (20 μl) were collected via a small nick in the tail vein at 15-min intervals for the immediate measurement of plasma glucose concentration, and 20% glucose was infused at variable rates to maintain plasma glucose at approximately 140–160 mg/dl. Insulin-stimulated whole-body glucose flux was estimated using a prime continuous infusion of high-pressure liquid chromatography-purified [3-3H] glucose (10-μCi bolus, 0.1 μCi/min; NEN Life Science Products, Boston, MA) throughout the clamps.
To estimate insulin-stimulated glucose transport activity and metabolism in skeletal muscle, 2-deoxy-D-[1-14C] glucose (NEN Life Science Products) was administered as a bolus (10 μCi) at 45 min before the end of clamps. Blood samples (20 μl) were taken at 80, 85, 90, 100, 110, and 120 min of clamp period for the determination of plasma [3H] glucose, 2-deoxy-D-[1-14C] glucose, and 3H2O concentrations. Additional blood samples (10 μl) were collected before and at the end of clamp studies for measurements of plasma insulin concentration. All infusions were performed using microdialysis pumps (CMA/Microdialysis, Acton, MA). At the end of the clamp period, animals were anesthetized with ketamine and xylazine injection. Within 5 min, gastrocnemius muscle from hindlimbs, epididymal and brown adipose tissue, and liver were removed. Each tissue, once exposed, was dissected out within 2 sec, frozen immediately using liquid N2-cooled aluminum blocks, and stored at –70 C for later analysis.
In vivo triglyceride secretion
Triglyceride secretion was measured as described before (35). Briefly, mice were fed fat-free diet (Frosted Flakes; Kellogg Co., Battle Creek, MI) for 4 h and then anesthetized with Avertin. WR1339 (100 μl of a 1:10 dilution in PBS; Sigma, St. Louis, MO) was injected via the tail vein, and plasma was collected at 0, 60, and 120 min after injection. Plasma triglyceride levels were measured as described above. Data are expressed as milligrams triglyceride per kilogram body weight per hour, assuming a plasma volume of 3.5% of body weight.
Triglyceride clearance
Mice were fasted for 4 h and then given 400 μl of peanut oil by a gavage. Plasma was collected hourly for 4 h from the tail vein. Plasma triglyceride levels were measured as described above. To examine triglyceride clearance during inhibition of lipase, mice were fasted for 4 h, injected with WR1339 via the tail vein, and then given 400 μl of peanut oil by a gavage.
Measurement of fatty acid oxidation in skeletal muscle
Soleus muscle was isolated from the anesthetized mice. Isolated soleus muscle strips were preincubated in Krebs-Henseleit buffer consisting of 118.5 mM NaCl, 4.7 mM KCl, 1.2 mM MgSO4, 1.2 mM KH2PO4, 2.5 mM CaCl2, and 25 mM NaHCO3, supplemented with 2% dialyzed BSA and 5 mM glucose for 20 min. Fatty acid oxidation was assayed by using 0.2 mM palmitate and 0.45 μCi/ml [1-C14] palmitate bound to 2% BSA for 30 min. Incubations were carried out at 30 C under the atmosphere of 95% O2-5% CO2. At the end of incubation, the soleus muscle was removed and weighed. A suspended center well containing 200 μl of 2 M NaOH was placed atop the flask. Then 1.25 ml perchloric acid was injected into the assay buffer to stop the reaction. The flask was lightly shaken for 2 h, and then NaOH was counted by the scintillation counter.
Pancreatic perfusions
All pancreatic perfusions were performed on nonfasted mice that were appropriately anesthetized. The procedure was based on that of Grodsky et al. (36) with some modifications. In brief, pancreata were isolated in vivo from the stomach, spleen, and duodenum via several ligations. The auxiliary arteries and the aorta above the celiac axis were ligated and the aorta below the celiac axis and the hepatic portal vein cannulated using PE50 tubing (Intramedic, Parsippany, NJ). Pancreata were then perfused via the arterial cannula with a Krebs-Ringer Buffer solution containing 2% BSA, 1.4 mmol/liter glucose, and 3% dextran. This perfusate solution was maintained at 37 C (pH 7.4) by passing it through a heated chamber while constantly gassing it with a mixture of 95% O2-5% CO2. After 20 min of preperfusion with the Krebs-Ringer buffer solution containing 1.4 mmol/liter glucose, the pancreata were perfused with a linear glucose gradient ranging from 1.4 to 20 mmol/liter over a period of 40 min. Fractions were collected every minute via the portal vein and assayed for insulin by RIA as previously described by Joseph et al. (37).
Total insulin content
At the completion of each perfusion experiment, whole pancreas was isolated and total insulin quantified. Whole pancreas was homogenized with extraction media containing 9.2% concentrated HCl, 5.6% formic acid, 1% trifluoroacetic acid (TFA), and 1% NaCl. A protein solution in isopropanol was isolated using a C-18 sep-pak cartridge (Waters Scientific, Mississauga, Ontario). From the extracted solution, protein was quantified using the Bio-Rad method (Hercules, CA) and insulin was measured by RIA.
Indirect calorimetry
Oxygen consumption and carbon dioxide production were measured using a 4-chamber Oxymax system (Columbus Instruments, Columbus, OH), with one mouse per chamber and testing transgenic mice simultaneously with controls (38). Motor activity (total and ambulating) was determined by infrared beam interruption (Opto-Varimex mini, Columbus Instruments). Mice had free access to food and water. Resting oxygen consumption was calculated as the average of the points with less than one ambulating beam break per minute, omitting the first hour of the experiment. The respiratory exchange ratio (RER), the ratio of carbon dioxide produced to oxygen consumed, was calculated using the same data points. The data for O2 consumption were normalized to (lean mass)0.75.
Tissue triglyceride assay
Tissue triglycerides were extracted with chloroform/methanol as described by Burant et al. (39). After hydrolysis with KOH base, triglycerides were measured radiometrically using a glycerol kinase assay (40).
Measurement of AMP-activated protein kinase (AMPK) kinase activity
Tissue lysates (200 mg protein) were immunoprecipitated with antibody against the 2 catalytic subunit of AMPK (Santa Cruz Biotechnology, Santa Cruz, CA) bound to protein-G/Sepharose beads. AMPK activity was measured using SAMS peptide as substrate (Upstate Biotechnology, Lake Placid, NY) as described before (41). Reaction products were resolved by electrophoresis on a 12% NuPAGE gel (Invitrogen, Carlsbad, CA) followed by Coomassie brilliant blue staining. The radiograms were obtained and quantified using the Image Reader software and Image Gauge software together with a Fuji BAS1800II instrument (Fuji Film, Stamford, CT).
Immunoblotting
To check the phosphorylation and total amount of AMPK, Akt, acetyl coenzyme A carboxylase (ACC), and glucose transporter protein (GLUT)4, the extracted protein samples were subjected to SDS-PAGE and Western blotting. Blots were probed with anti-phospho-AMPK (Thr172), anti-AMPK, anti-phospho-ACC (Ser79), anti-phospho-Akt, anti-Akt (Cell Signaling Technology, Beverly, MA), or anti-GLUT4 antibodies (Biogenesis Inc., Brentwood, NH). Detection of immunoreactive bands was performed using the enhanced chemiluminescence kit (PerkinElmer Life Science Products, Boston, MA). Densitometry was performed by scanning the radiographs using the Mac Bas V2.52 (Fuji Film).
Determination of gene expression
Total RNA was isolated using TRIzol reagent (Invitrogen) following the manufacturer’s instructions. Total RNA (20 μg) was resolved by a 1.25% denaturing agarose gels, transferred to Nytran nylon membrane (Schleicher & Schüell, Keene, NH), and hybridized with 32P-labeled cDNA probes as described before (42). The hybridized radioactivity was measured using Fuji BAS1800II instrument (Fuji Film). The mRNA levels were normalized to the 18S rRNA.
Statistical analysis
All values are expressed as the mean ± SEM. Statistical significance was determined by unpaired Student’s t test or one-way ANOVA with Fisher’s protected least significant difference test using Statview 5.0 software (SAS Institute Inc., Cary, NC). Differences were considered to be statistically significant at P < 0.05.
Results
Leptin decreases adipose tissue mass but does not affect energy expenditure in MKR mice
Leptin (10 μg/d) was administered to WT and MKR mice by sc implanted osmotic pumps for 14 d, resulting in a 10-fold increase in circulating leptin levels (Table 1). In response to treatment, both WT and MKR mice reduced their daily food intake by 22% and body weight by 10% (Table 1). Epididymal fat pad and whole-body fat mass were markedly reduced in the leptin-treated mice, but there were no significant differences in adipose tissue mass between the saline-treated WT and MKR mice (Table 1). The size of adipocytes was smaller after leptin treatment in both WT and MKR mice, and there were an increased number of cells with multilocular fat droplets resembling brown adipocytes (Fig. 1 and data not shown).
It has been shown that leptin replacement increases the metabolic rate and activity levels in leptin-deficient ob/ob mice (6). To determine whether leptin can increase energy expenditure in the MKR mice, we measured their metabolic rates by indirect calorimetry. No major differences were observed in resting and total oxygen consumptions, respiratory exchange ratio, and total activity at both 23 and 30 C in all groups of mice, suggesting that leptin treatment did not affect the metabolic rate in WT and MKR mice (Table 1 and data not shown). Importantly, the leptin-treated WT and MKR mice did not show a decrease in energy expenditure despite the decreased food intake.
Leptin improves the hyperglycemia and hyperinsulinemia in MKR mice
We examined the effect of leptin on blood glucose and serum insulin levels in MKR mice. The hyperglycemia in MKR mice was normalized after leptin treatment (Fig. 2A). The elevated serum insulin level in MKR mice was also significantly decreased (Fig. 2B). In addition, we examined whether the changes in an antidiabetic adipocytokine, adiponectin, was involved in these improvements. Serum adiponectin levels were not changed after leptin treatment (Table 1). These data demonstrated that leptin treatment improves diabetes in non-leptin-deficient and nonobese diabetic mice.
Because blood glucose and serum insulin levels were decreased by leptin in MKR mice, we anticipated that insulin sensitivity might be increased by leptin treatment in MKR mice. We performed insulin tolerance and glucose tolerance tests. In MKR mice, both insulin and glucose tolerance were improved by leptin treatment, suggesting that leptin increases insulin sensitivity in MKR mice (Fig. 2, C and D). In addition, fasting blood glucose was significantly lower in the leptin-treated MKR mice, compared with the saline-treated MKR and both groups of WT mice (Fig. 2D). It is not clear why leptin-treated MKR mice had very low fasting glucose levels. One possible explanation is that they are more sensitive to leptin particularly under fasting condition. Fasting insulin levels were higher in leptin-treated MKR mice, compared with leptin-treated WT mice (0.48 ± 0.13 vs. 0.25 ± 0.03 ng/ml, P = 0.055). Leptin also caused a significant reduction in elevated corticosterone levels in MKR mice (from 420 + 130 to 121 ± 22 ng/ml, P = 0.032). Both factors may contribute to a lower hepatic glucose production. It is also possible that leptin-treated MKR mice poorly adapt to fasting because of their smaller size and inability to down-regulate metabolic rate in response to caloric restriction.
The improvement of diabetes by leptin was not due to the reduced food intake in MKR mice
To exclude the possibility that the improvement of diabetes in MKR mice was due to the reduction of food intake, we pair fed one group of MKR mice treated with saline to the amount of food consumed by the MKR mice treated with leptin on the previous day. Pair feeding did not improve hyperinsulinemia, hypertriglyceridemia, and insulin and glucose tolerance in MKR mice (see Figs. 2, E and F, and 5, C and D, and data not shown). These results suggest that the antidiabetic effect of leptin was independent of caloric restriction in MKR mice.
Leptin enhances insulin sensitivity in liver and adipose tissues of MKR mice
The results of the insulin tolerance test suggested that leptin increased insulin sensitivity in MKR mice. We performed hyperinsulinemic-euglycemic clamps in MKR mice to assess insulin sensitivity in individual tissues. In the basal state (after 10 h of fasting), the leptin-treated MKR mice showed lower plasma glucose than the MKR mice treated with saline (MKR-saline: 241 ± 36 mg/dl; MKR-leptin: 126 ± 35 mg/dl). In addition, basal endogenous glucose production (EGP) was significantly reduced by 30% in the leptin-treated MKR mice (Fig. 3B). These findings suggest that the decreased basal endogenous glucose production partly contributes to the lower basal glucose levels.
The glucose infusion rate during the clamp was increased 4-fold (Fig. 3A), suggesting an increase in insulin sensitivity by leptin treatment. The clamp EGP was significantly lowered by 90% in the leptin-treated MKR mice (Fig. 3B). When the clamp EGP levels were compared with the basal level of each group, it was reduced by 40% in MKR mice treated with saline but further reduced by 90% in MKR mice treated with leptin. Thus, these results suggest that leptin increased hepatic insulin sensitivity in MKR mice.
During the clamp period, whole-body glucose uptake showed a tendency to increase in the leptin-treated MKR mice but did not reach statistical significance (P < 0.064; Fig. 3C). Whereas muscle glucose uptake was unaffected (Fig. 3D), the clamp glucose uptake in white adipose tissue (WAT) (Fig. 3E) and brown adipose tissue (Fig. 3F) was significantly increased by 4- and 10-fold, respectively, after leptin treatment. Thus, in MKR mice, leptin enhanced insulin-stimulated glucose uptake in adipose tissues, and this was associated with only a limited increase in whole-body glucose uptake.
Leptin inhibits insulin secretion in MKR mice
Previous studies have shown that MKR mice have -cell dysfunction (29, 30, 43). To examine whether leptin treatment improves -cell function, we performed pancreatic perfusion experiments in situ. MKR mice treated with saline showed an impaired glucose responsiveness as previously described (Fig. 4A) (30, 43). Insulin secretion under low glucose (1.4–5 mM) conditions showed no significant difference between saline- and leptin-treated MKR mice (Fig. 4B). However, under stimulatory (6–20 mM) glucose conditions, insulin secretion was inhibited in leptin-treated MKR mice (Fig. 4, A and B). Thus, leptin inhibited glucose-stimulated insulin secretion despite no reduction in total insulin content in the pancreas of MKR mice (data not shown). Thus, the improvement of hyperinsulinemia might be, in part, due to the leptin-inhibited insulin secretion and, in part, due to the reduction in insulin resistance.
Leptin improves lipid metabolism in MKR mice
Serum triglyceride (TG), which is abnormally elevated in MKR mice, was normalized after leptin treatment (Fig. 5A). Similarly, leptin reduced serum TG levels in WT mice (Fig. 5A). Serum FFA levels were decreased by leptin treatment in MKR mice (Fig. 5B). These effects of leptin on the circulating lipid levels were not due to the reduced caloric intake because pair feeding had no effect on serum TG and FFA levels in MKR mice (Fig. 5, C and D). Liver TG content was significantly decreased by leptin treatment in both WT and MKR mice (Fig. 5E). However, pair feeding also lowered liver TG content in MKR mice (Fig. 5E). In muscle, TG content in the saline-treated MKR mice was significantly increased as compared with the saline-treated WT mice (Fig. 5D). After leptin treatment, muscle TG content was normalized in MKR mice (Fig. 5F). Muscle TG content was not affected by pair feeding (Fig. 5F). Thus, leptin improved hyperlipidemia and reduced liver and muscle TG contents in MKR mice. Whereas caloric restriction may contribute to the lowering of liver TG content, the rest of the effects appear to be independent of leptin’s effect on food intake.
To demonstrate the mechanisms whereby leptin decreased serum and tissue TG levels, we measured TG secretion rates in WT and MKR mice. To quantify the TG secretion, the rate of increase in circulating TG was measured after inhibiting TG clearance with WR1339. TG secretion reflects whole-body TG production, with liver being the major source. Leptin treatment reduced whole-body TG production by 30% in WT and MKR mice (Fig. 6A), which correlates with the decrease in serum and liver TG contents after leptin treatment in both MKR and WT mice.
We also measured plasma TG levels after an oral lipid load to examine the TG clearance in WT and MKR mice treated with and without leptin. In the saline-treated WT and MKR mice, plasma TG peaked at 2–3 h after the lipid load (384 and 481 mg/dl at 2 h, respectively) and then gradually decreased (Fig. 6B). In the leptin-treated WT and MKR mice, plasma TG rose much less than the saline-treated mice, peaking at 161 and 193 mg/dl at 2 h after the lipid load, respectively. This was not due to poor TG uptake from the intestine because plasma TG levels increased similarly in the saline-treated WT and MKR mice in the absence of TG clearance after lipid load (Fig. 6C). Thus, leptin can enhance TG clearance in MKR mice, suggesting that leptin might be able to increase lipase activity.
Finally, we measured fatty acid oxidation in isolated skeletal muscle of WT and MKR mice with and without leptin infusion. Fatty acid oxidation was significantly inhibited in the saline-treated MKR mice as compared with WT mice. Leptin enhanced fatty acid oxidation in skeletal muscle in MKR mice but not WT mice (Fig. 6D). These data suggested that the enhanced muscle fatty acid oxidation led to the reduction of TG contents in MKR mice after leptin treatment.
All these effects might contribute to the improvement of lipotoxic conditions in the MKR mice.
Molecular mechanisms whereby leptin reduced glucose output and TG content in liver
To explore the molecular mechanisms whereby leptin inhibited glucose production and decreased TG accumulation in liver, we measured the expression of several genes related to glucose and lipid metabolism. The gene expression of glucose-6-phosphatase (G6Pase), which is one of the enzymes that regulates gluconeogenesis, was significantly decreased after leptin treatment in MKR mice (Fig. 7A). In addition, we found that the gene expression of GLUT-2, a low-affinity glucose transport protein that facilitates either glucose uptake or efflux from liver, depending on the nutritional state (44), was significantly reduced after leptin treatment (Fig. 7B). Because the glucose transporters are facilitative transporters, flux of glucose is dependent on a gradient, and therefore they are involved in both influx and efflux of glucose from the tissues. Thus, a reduction in GLUT2 expression could also explain the reduced glucose output from the liver after leptin treatment. Thus, the decreased hepatic glucose output by leptin treatment could be attributed to the reduction in both gluconeogenesis and glucose efflux from the liver.
We demonstrated that the expression of the gene for sterol-regulatory-element-binding protein-1c (SREBP-1c), a transcription factor that globally activates pathways of lipid biosynthesis (45), was significantly reduced in MKR mice after leptin treatment (Fig. 7C). In parallel, the expression of fatty acid synthase (FAS) and stearoyl-CoA desaturase-1 (SCD-1) were inhibited by leptin treatment (Fig. 7, D and F). Furthermore, CD36 gene expression, a fatty acid transporter, was also decreased in the leptin-treated MKR mice. Thus, the reduction of TG levels in liver could be due to the reduction of lipid synthesis and fatty acid uptake.
Leptin enhanced AMPK activity in skeletal muscle of MKR mice
Leptin has been shown to selectively stimulate phosphorylation and activation of the 2 catalytic subunit of AMPK, an enzyme that has been found to be a principal mediator of the leptin effects on fatty acid oxidation in skeletal muscle (8). Because the TG content in skeletal muscle was decreased and fatty acid oxidation was increased in muscle by leptin, we studied AMP kinase activity in muscle. In skeletal muscle, AMP kinase activity was 3-fold increased in the leptin-treated MKR mice (Fig. 7G). In addition, the phosphorylation of AMPK also was increased 1.5-fold by leptin in MKR mice (Fig. 7H). We also checked the phosphorylation of ACC, which is phosphorylated and inhibited by AMPK. The phosphorylation of ACC was increased by leptin in skeletal muscle of MKR mice (data not shown). These results suggest that the leptin-enhanced fatty acid oxidation in skeletal muscle was due to the increased AMPK activity.
Because dominant-negative IGF-I receptor also inhibits insulin receptor signaling (29) and hyperinsulinemic-euglycemic clamp data showed no changes in insulin-stimulated muscle glucose uptake in MKR mice (Fig. 3D), we did not expect to see any changes in insulin signaling in skeletal muscle of MKR mice after leptin treatment. To confirm this hypothesis, we measured phosphorylation of Akt, a downstream mediator of insulin-stimulated glucose uptake (46), in skeletal muscle of the saline- and leptin-treated WT and MKR mice. In WT mice, leptin increased the level of Akt phosphorylation, suggesting an increase in insulin signaling (Fig 8, A and B). In MKR mice, Akt phosphorylation was dramatically lower than in WT mice and was not affected by leptin treatment. Muscle GLUT4 protein levels were comparable in WT and MKR mice and did not change in response to leptin treatment (Fig. 8A). Thus, in skeletal muscle of MKR mice, leptin increased AMPK activity but had no effects on Akt phosphorylation or GLUT4 protein levels.
Discussion
MKR mice are a model for type 2 diabetes. They develop insulin resistance initially in muscle because of genetically impaired IGF-I and insulin receptor signaling (29), which leads to insulin resistance in liver and fat. Subsequently pancreatic -cell dysfunction develops, as demonstrated by the loss of first-phase insulin secretion. This event is associated with the appearance of diabetes (29). Unlike either genetically leptin-deficient or lipodystrophic diabetes models, MKR mice have normal amounts of adipose tissue and serum leptin levels (30, 31).
This study clearly demonstrates that leptin treatment improves insulin resistance and hyperglycemia in a specific mouse model of type 2 diabetes. Leptin treatment enhanced hepatic insulin sensitivity of the MKR mice by suppression of hepatic glucose production during the hyperinsulinemic-euglycemic clamp. Additionally, leptin reduced gene expression of key regulator of glucose metabolism such as G6Pase, a gluconeogenic enzyme in liver. All together, these findings suggest that leptin decreased glucose production through the inhibition of gluconeogenic enzymes. In previous studies, it has been reported that insulin action on hepatic glucose output in normal rats was enhanced by leptin (11, 47). Whereas it was also shown that the suppression of hepatic glycogenolysis primarily contributed to the decreased glucose production, gluconeogenesis was increased and accounted for the majority of glucose production. These events were accompanied by an increase in G6Pase and phosphoenolpyruvate carboxykinase gene expression. The discrepancy between the two studies might be due to differences in the regulation of gluconeogenesis affected by leptin between normal and diabetic animals or different responses of different animal species. In this study, it is also demonstrated that GLUT2 gene expression was decreased after leptin treatment. The liver senses hepatoportal glucose levels in a GLUT2-dependent manner and adjusts glucose influx and efflux to maintain euglycemia (48). Because GLUT2 expression was decreased by leptin treatment, glucose flux in both directions in the liver might be reduced. These events correlate with the reduction of gluconeogenesis. In addition, the decreased glucose influx could induce the reduction of fatty acid synthesis. It is possible that the leptin-suppressed GLUT2 gene expression could be secondary to other effects induced by prolonged leptin treatment (49). Thus, the reduction of both gluconeogenesis and efflux of glucose from liver could lead to the decreased hepatic glucose output after leptin treatment. Because the effect of leptin on hepatic glucose flux is largely mediated by hypothalamus (47, 50, 51), our findings might also be primarily related to the hypothalamic regulation. A recent study reported that the antidiabetic effect of leptin in mice with congenital lipodystrophy is mainly ascribed to its action on the liver (51). Our data also suggest that the improvement of liver function is important in the treatment of diabetes by leptin.
Unlike the liver, there was no improvement of insulin-stimulated glucose uptake in skeletal muscle. Previous studies have shown that chronic or acute leptin treatment in either normal or obese rodents increased glucose uptake and insulin sensitivity in skeletal muscle (7, 9). However, in the MKR mouse model, both the insulin and IGF-I receptor signaling pathways in skeletal muscle are genetically blocked, and therefore, not surprisingly, no leptin effect on glucose uptake could be detected.
A major cause of type 2 diabetes is a progressive insulin resistance of peripheral tissues, followed by a defect in insulin secretion in the islets of Langerhans (52). MKR mice showed islet cell hyperplasia, which resulted from severe insulin resistance in skeletal muscle, liver, and adipose tissues (29, 30, 43). Our previous studies demonstrated that the improvement of insulin sensitivity caused by peroxisomal proliferator-activated receptor- agonist and muscle CD36 overexpression led to the normalization of pancreatic -cells function in MKR mice (30, 43). However, in the present study, leptin did not improve the insulin responses of pancreatic -cells despite its improvement of insulin sensitivity. Instead, leptin inhibited glucose-stimulated insulin secretion. Leptin was shown to inhibit acutely glucose-stimulated insulin secretion in isolated islets from ob/ob mice (53). Similarly, acute physiological increases in plasma leptin levels inhibited glucose-simulated insulin secretion in conscious rats (54). Furthermore, hyperleptinemia achieved by adenovirus-leptin gene therapy had similar effects on insulin secretion, which could be explained by the depletion of lipids from the islets (55). It is likely that in the MKR mouse model, the effect of chronic leptin treatment on inhibition of insulin secretion is stronger than the acute glucose-stimulated insulin secretion.
Leptin treatment improved the hyperlipidemia of MKR mice by enhancing TG clearance and inhibiting TG secretion. Because leptin has been shown to prevent the accumulation of lipid content in nonadipose tissues in normal and lipodystrophic mice (21, 56, 57), the elevated TG levels in liver and skeletal muscle of MKR mice were significantly decreased after leptin treatment. In skeletal muscle, the reduction of TG contents was most likely secondary to the increased fatty acid oxidation, which was derived by the stimulation of AMPK kinase activity. In contrast, in liver, leptin apparently inhibited both lipid biosynthesis and fatty acid uptake by inhibiting gene expression of SREBP-1c, FAS, SCD-1, and CD36. These events might contribute to the reduction of liver TG content after leptin treatment. Our data suggest that a decreased caloric intake may also contribute to the reduction of liver TG content in the leptin-treated MKR mice. In addition to the lipid effects in nonadipose tissues, we observed a reduction in the amount of WAT. Hyperleptinemia in normal rats, induced by means of adenovirus-mediated gene transfer of the leptin gene, has reported a marked depletion of body fat content without increasing circulating FFAs (58). The effect of leptin on the transformation fat-storing adipocytes into fat-burning cells could be related to the depletion of body fat mass (59). This effect on WAT included the reduction in the gene expression of lipogenic enzymes and the increased phosphorylation of AMPK. We did not assess the mechanisms whereby leptin decreases the fat depot, but we can expect that similar molecular events might occur in WAT of MKR mice because we observed an increase in the number of cells with multilocular fat droplets resembling brown adipocytes.
Our previous studies using the MKR mouse model showed that the improvement in lipotoxicity (but not glucotoxicity) led to an improvement in insulin sensitivity and hyperglycemia (30, 31, 43). The improvement of diabetes after leptin treatment could also be due to the improvement of lipotoxicity in MKR mice. Namely, leptin enhanced lipid oxidation in skeletal muscle and apparently inhibited both lipid biosynthesis and fatty acid uptake in liver, leading to the reduction of lipid deposition in both tissues, thereby improving lipotoxicity and improving insulin sensitivity and diabetes.
It has been reported that leptin could be used as an adjunct of insulin therapy in streptozotocin-induced, insulin-dependent diabetic mice (60) and restore euglycemia and glucose turnover in streptozotocin-induced, insulin-dependent diabetic rats (61). Furthermore, in several clinical trials of leptin treatment, it has been shown that leptin improved the glycemic control, increased insulin sensitivity, and decreased TG levels in patients with lipodystrophy and leptin deficiency (19, 20, 62). More recently it has been reported that leptin administration improved fasting hyperglycemia, hyperinsulinemia, and glucose and insulin tolerances in patients with Rabson-Mendenhall syndrome, which show severe insulin resistance with presumed insulin receptor mutations, low levels of serum TG, and normal serum leptin levels (63).
Obesity is emerging as a worldwide epidemic and the number of obese type 2 diabetic patients is consequently increasing. Circulating leptin levels are strongly correlated with fat mass. In general, obese patients and rodents have elevated leptin levels (24, 64). It has been shown that leptin resistance exists in hyperleptinemic states, although the mechanisms involved in leptin resistance remain unclear. However, there was considerable variability in the effect of leptin treatment in obese patients (26). Furthermore, a large proportion of lean patients with type 2 diabetes have leptin levels in normal range (23, 25). Because leptin is currently available for clinical studies, our findings might provide supportive evidence for the use of leptin as a therapy for some type 2 diabetic patients. However, because chronic hyperleptinemia could induce hypertension (65) and reversible unresponsiveness to glucose in -cells (55) in animal models, further studies are necessary to see whether long-term leptin therapy has any side effects
In summary, the present study demonstrates that leptin can reverse diabetes by a combination of improvement in glucose and lipid metabolism in MKR mice having normal circulating leptin levels. These findings are independent of decreased food intake, suggesting that leptin could be a useful form of treatment for not only patients with lipodystrophic diabetes but also type 2 diabetes patients without leptin resistance.
Footnotes
This work was supported in part by a Mentor grant (7-02-MN-15) and a Research grant (7-02-RA-49) from the American Diabetes Association awarded to D.L. and by the Canadian Diabetes Association grants (to M.B.W.). M.B.W. is a Canadian Institutes for Health Research Investigator.
Abbreviations: ACC, Acetyl coenzyme A carboxylase; AMPK, AMP-activated protein kinase; EGP, endogenous glucose production; FAS, fatty acid synthase; FFA, free fatty acid; GLUT, glucose transporter protein; G6Pase, glucose-6-phosphatase; MKR, transgenic overexpression of a skeletal muscle dominant-negative IGF-I receptor with a lysine-to-arginine amino acid; RER, respiratory exchange ratio; SCD-1, stearoyl-CoA desaturase-1; SREBP-1c, sterol-regulatory-element-binding protein-1c; TG, triglyceride; WAT, white adipose tissue; WT, wild type.
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Department of Physiology (Z.A., M.W.), University of Toronto, Toronto, Canada M5S 1A8
Abstract
Leptin has metabolic effects on peripheral tissues including muscle, liver, and pancreas, and it has been successfully used to treat lipodystrophic diabetes, a leptin-deficient state. To study whether leptin therapy can be used for treatment of more common cases of type 2 diabetes, we used a mouse model of type 2 diabetes (MKR mice) that show normal leptin levels and are diabetic due to a primary defect in both IGF-I and insulin receptors signaling in skeletal muscle. Here we show that leptin administration to the MKR mice resulted in improvement of diabetes, an effect that was independent of the reduced food intake. The main effect of leptin therapy was enhanced hepatic insulin responsiveness possibly through decreasing gluconeogenesis. In addition, the reduction of lipid stores in liver and muscle induced by enhancing fatty acid oxidation and inhibiting lipogenesis led to an improvement of the lipotoxic condition. Our data suggest that leptin could be a potent antidiabetic drug in cases of type 2 diabetes that are not leptin resistant.
Introduction
LEPTIN IS AN adipocyte-derived hormone that plays a key role in the regulation of food intake and energy expenditure (1, 2). Generally, leptin regulates energy homeostasis via the long isoform of the leptin receptor, which is highly expressed in hypothalamus (3, 4). It also regulates lipid and glucose metabolism and insulin action independently of its effects on food intake (5, 6). Leptin stimulates fatty acid oxidation and glucose uptake in skeletal muscle (7, 8, 9, 10), inhibits glucose output and lipogenesis in liver (11, 12), and inhibits insulin secretion (13). These metabolic effects of leptin are thought to be mainly exerted via the hypothalamus. However, the long isoform of the leptin receptor is also expressed in peripheral tissues (3, 4, 14). Thus, it is possible that leptin could regulate metabolism acting directly on peripheral tissues. However, the underlying mechanisms by which leptin regulates on metabolism and insulin action remain unclear.
Leptin deficiency contributes to the insulin resistance of the ob/ob mice bearing a mutation in the leptin gene (15, 16) and leptin administration to ob/ob mice ameliorates the hyperglycemia, hypertriglyceridemia, and hyperinsulinemia (6, 17, 18). Leptin deficiency is also a primary cause of insulin resistance in mice and humans with lipodystrophy who have reduced amount of adipose tissue. Leptin replacement therapy improves most of the metabolic abnormalities associated with lipodystrophy (19, 20, 21) and transgenic overexpression of leptin gene also improved insulin resistance and diabetes in a mouse model of lipoatrophic diabetes (22). Thus, leptin has been shown to be effective for the treatment of diabetes associated with leptin deficiency. However, leptin-deficient states are extremely rare; the great majority of the patients with type 2 diabetes have either normal or elevated leptin levels (23, 24, 25, 26). Previous studies showed that transgenic overexpression of the leptin gene led to increased insulin sensitivity in normal mice (27) and could delay the onset of impaired glucose metabolism in lethal yellow KKAy mice (28). However, thus far, there is no evidence that leptin improves type 2 diabetes where circulating leptin levels are normal.
We developed a transgenic mouse model overexpressing a dominant-negative IGF-I receptor specifically in skeletal muscle [MKR (transgenic overexpression of a skeletal muscle dominant-negative IGF-I receptor with a lysine-to-arginine amino acid) mice] (29). In MKR mice, the formation of hybrid receptors between defective IGF-I receptor and endogenous IGF-I and insulin receptors led to the abrogation of both IGF-I and insulin receptor signaling pathways in skeletal muscle (29). MKR mice develop an early-onset type 2 diabetes, characterized by dyslipidemia; hyperinsulinemia; and secondary insulin resistance in liver and fat, excessive lipid stores in muscle and liver, and -cell dysfunction (29). They have normal serum leptin levels (30, 31). The MKR mouse is therefore an excellent model to examine whether leptin administration would be effective to treat type 2 diabetes with normal circulating leptin. Here we demonstrate that leptin serves as an antidiabetic agent for the MKR mice.
Materials and Methods
Experimental animals
The generation of the MKR mice (FVB/N background) has been described previously (29). All experiments were performed in homozygous MKR male mice, 6–7 wk of age. As controls, sex- and age-matched wild-type (WT) mice were used. Mice were kept on a 12-h light (0600–1800 h), 12-h dark cycle (1800–0600 h) and were fed NIH-07 rodent chow (Zeigler Brothers Inc., Gardners, PA) and water ad libitum. All studies were conducted in accordance with National Institutes of Health guidelines, as approved by the Animal Care and Use Committee of National Institute of Diabetes and Digestive and Kidney Diseases.
Leptin treatment
For leptin infusion, an Alzet miniosmotic pump (model 2002; Alza Corp., Palo Alto, CA) was implanted sc on the back of each mouse. The pumps delivered saline or 10 μg leptin/d (R&D Systems, Minneapolis, MN) for 14 d. After placement of the pump, the mice were housed individually. At the end of the treatment, mice were killed under anesthesia with Avertin in the nonfasting state between 0900 and 1200 h.
Food intake
Mice were caged individually and treated with or without leptin, as described above. The amount of food in the feeding container was measured at d 14 of treatment. Food intake was expressed as grams per day per mouse.
Pair-feeding study
Mice were treated with or without leptin for 2 wk, as described above. The amount of food consumed by the leptin-treated MKR mice was monitored each day at 1800 h, and that amount was given to the one of the saline-treated MKR mice on the next day.
Histology
The left epididymal fat pad was isolated and fixed overnight in 4% paraformaldehyde in PBS. The tissues were then transferred to 70% ethanol and embedded in paraffin. Samples were cut into 5-μm sections and hematoxylin and eosin staining was performed.
Biochemical assays
Blood was obtained from the tail vein and glucose levels were measured using a Glucometer Elite (Bayer Corp., Elkhart, IN). Serum samples were generally obtained from the tail or retroorbital vein in the nonfasting state between 0900 and 1200 h. Triglycerides (Thermo DMA, Louisville, CO) and free fatty acid (FFA) (Roche Applied Science, Indianapolis, IN) were measured according to the manufacturer’s procedures. Insulin (SRI-13K; Linco Research Inc., St. Charles, MO), leptin (ML-82K; Linco Research), and adiponectin (MADP-60HK; Linco Research) were measured by RIAs.
Determination of body composition
Body composition was measured in nonanesthetized mice using the Bruker minispec NMR analyzer mq10 (Bruker Optics Inc., The Woodlands, TX).
Insulin tolerance and glucose tolerance test
Insulin and glucose tolerance tests were performed with 5- or 7-h fasted mice during daylight time. Human insulin (0.75 U/kg body weight) or glucose (2 g/kg body weight) were injected ip. Blood glucose levels were measured at the indicated time points.
Hyperinsulinemic-euglycemic clamps
The clamp studies were performed as developed by Kim et al. (32, 33). The study involved a primed insulin infusion. Mice were treated with or without leptin for 2 wk, as described above. On d 7 of the treatment (4 d before the clamp experiment), mice were anesthetized with 100 mg/kg ketamine and 10 mg/kg xylazine. A catheter was inserted into a lateral incision on the right side of the neck and advanced into the superior vena cava via the right internal jugular vein. The catheter was then sutured into place, according to the protocol of MacLeod and Shapiro (34). The evening before the clamp analysis, mice were fasted for 10 h. The basal rates of glucose turnover were measured by continuous infusion of [3-3H] glucose (0.02 μCi/min) for 120 min that followed a bolus of 2.5 μCi, starting at 0900 h. Blood samples (20 μl) were taken at 90 min and 115 min of the basal period for the determination of plasma [3-3H] glucose concentration. A 120-min hyperinsulinemic-euglycemic clamp was started at 1100 h. Insulin was infused as a bolus of 300 mU/kg over a period of 3 min, followed by continuous insulin infusion at the rate of 2.5 mU/kg·min (Humulin R, Eli Lilly, Indianapolis, IN) to raise plasma insulin concentration to 4 ng/ml. During the clamp study, mice were restrained and blood samples (20 μl) were collected via a small nick in the tail vein at 15-min intervals for the immediate measurement of plasma glucose concentration, and 20% glucose was infused at variable rates to maintain plasma glucose at approximately 140–160 mg/dl. Insulin-stimulated whole-body glucose flux was estimated using a prime continuous infusion of high-pressure liquid chromatography-purified [3-3H] glucose (10-μCi bolus, 0.1 μCi/min; NEN Life Science Products, Boston, MA) throughout the clamps.
To estimate insulin-stimulated glucose transport activity and metabolism in skeletal muscle, 2-deoxy-D-[1-14C] glucose (NEN Life Science Products) was administered as a bolus (10 μCi) at 45 min before the end of clamps. Blood samples (20 μl) were taken at 80, 85, 90, 100, 110, and 120 min of clamp period for the determination of plasma [3H] glucose, 2-deoxy-D-[1-14C] glucose, and 3H2O concentrations. Additional blood samples (10 μl) were collected before and at the end of clamp studies for measurements of plasma insulin concentration. All infusions were performed using microdialysis pumps (CMA/Microdialysis, Acton, MA). At the end of the clamp period, animals were anesthetized with ketamine and xylazine injection. Within 5 min, gastrocnemius muscle from hindlimbs, epididymal and brown adipose tissue, and liver were removed. Each tissue, once exposed, was dissected out within 2 sec, frozen immediately using liquid N2-cooled aluminum blocks, and stored at –70 C for later analysis.
In vivo triglyceride secretion
Triglyceride secretion was measured as described before (35). Briefly, mice were fed fat-free diet (Frosted Flakes; Kellogg Co., Battle Creek, MI) for 4 h and then anesthetized with Avertin. WR1339 (100 μl of a 1:10 dilution in PBS; Sigma, St. Louis, MO) was injected via the tail vein, and plasma was collected at 0, 60, and 120 min after injection. Plasma triglyceride levels were measured as described above. Data are expressed as milligrams triglyceride per kilogram body weight per hour, assuming a plasma volume of 3.5% of body weight.
Triglyceride clearance
Mice were fasted for 4 h and then given 400 μl of peanut oil by a gavage. Plasma was collected hourly for 4 h from the tail vein. Plasma triglyceride levels were measured as described above. To examine triglyceride clearance during inhibition of lipase, mice were fasted for 4 h, injected with WR1339 via the tail vein, and then given 400 μl of peanut oil by a gavage.
Measurement of fatty acid oxidation in skeletal muscle
Soleus muscle was isolated from the anesthetized mice. Isolated soleus muscle strips were preincubated in Krebs-Henseleit buffer consisting of 118.5 mM NaCl, 4.7 mM KCl, 1.2 mM MgSO4, 1.2 mM KH2PO4, 2.5 mM CaCl2, and 25 mM NaHCO3, supplemented with 2% dialyzed BSA and 5 mM glucose for 20 min. Fatty acid oxidation was assayed by using 0.2 mM palmitate and 0.45 μCi/ml [1-C14] palmitate bound to 2% BSA for 30 min. Incubations were carried out at 30 C under the atmosphere of 95% O2-5% CO2. At the end of incubation, the soleus muscle was removed and weighed. A suspended center well containing 200 μl of 2 M NaOH was placed atop the flask. Then 1.25 ml perchloric acid was injected into the assay buffer to stop the reaction. The flask was lightly shaken for 2 h, and then NaOH was counted by the scintillation counter.
Pancreatic perfusions
All pancreatic perfusions were performed on nonfasted mice that were appropriately anesthetized. The procedure was based on that of Grodsky et al. (36) with some modifications. In brief, pancreata were isolated in vivo from the stomach, spleen, and duodenum via several ligations. The auxiliary arteries and the aorta above the celiac axis were ligated and the aorta below the celiac axis and the hepatic portal vein cannulated using PE50 tubing (Intramedic, Parsippany, NJ). Pancreata were then perfused via the arterial cannula with a Krebs-Ringer Buffer solution containing 2% BSA, 1.4 mmol/liter glucose, and 3% dextran. This perfusate solution was maintained at 37 C (pH 7.4) by passing it through a heated chamber while constantly gassing it with a mixture of 95% O2-5% CO2. After 20 min of preperfusion with the Krebs-Ringer buffer solution containing 1.4 mmol/liter glucose, the pancreata were perfused with a linear glucose gradient ranging from 1.4 to 20 mmol/liter over a period of 40 min. Fractions were collected every minute via the portal vein and assayed for insulin by RIA as previously described by Joseph et al. (37).
Total insulin content
At the completion of each perfusion experiment, whole pancreas was isolated and total insulin quantified. Whole pancreas was homogenized with extraction media containing 9.2% concentrated HCl, 5.6% formic acid, 1% trifluoroacetic acid (TFA), and 1% NaCl. A protein solution in isopropanol was isolated using a C-18 sep-pak cartridge (Waters Scientific, Mississauga, Ontario). From the extracted solution, protein was quantified using the Bio-Rad method (Hercules, CA) and insulin was measured by RIA.
Indirect calorimetry
Oxygen consumption and carbon dioxide production were measured using a 4-chamber Oxymax system (Columbus Instruments, Columbus, OH), with one mouse per chamber and testing transgenic mice simultaneously with controls (38). Motor activity (total and ambulating) was determined by infrared beam interruption (Opto-Varimex mini, Columbus Instruments). Mice had free access to food and water. Resting oxygen consumption was calculated as the average of the points with less than one ambulating beam break per minute, omitting the first hour of the experiment. The respiratory exchange ratio (RER), the ratio of carbon dioxide produced to oxygen consumed, was calculated using the same data points. The data for O2 consumption were normalized to (lean mass)0.75.
Tissue triglyceride assay
Tissue triglycerides were extracted with chloroform/methanol as described by Burant et al. (39). After hydrolysis with KOH base, triglycerides were measured radiometrically using a glycerol kinase assay (40).
Measurement of AMP-activated protein kinase (AMPK) kinase activity
Tissue lysates (200 mg protein) were immunoprecipitated with antibody against the 2 catalytic subunit of AMPK (Santa Cruz Biotechnology, Santa Cruz, CA) bound to protein-G/Sepharose beads. AMPK activity was measured using SAMS peptide as substrate (Upstate Biotechnology, Lake Placid, NY) as described before (41). Reaction products were resolved by electrophoresis on a 12% NuPAGE gel (Invitrogen, Carlsbad, CA) followed by Coomassie brilliant blue staining. The radiograms were obtained and quantified using the Image Reader software and Image Gauge software together with a Fuji BAS1800II instrument (Fuji Film, Stamford, CT).
Immunoblotting
To check the phosphorylation and total amount of AMPK, Akt, acetyl coenzyme A carboxylase (ACC), and glucose transporter protein (GLUT)4, the extracted protein samples were subjected to SDS-PAGE and Western blotting. Blots were probed with anti-phospho-AMPK (Thr172), anti-AMPK, anti-phospho-ACC (Ser79), anti-phospho-Akt, anti-Akt (Cell Signaling Technology, Beverly, MA), or anti-GLUT4 antibodies (Biogenesis Inc., Brentwood, NH). Detection of immunoreactive bands was performed using the enhanced chemiluminescence kit (PerkinElmer Life Science Products, Boston, MA). Densitometry was performed by scanning the radiographs using the Mac Bas V2.52 (Fuji Film).
Determination of gene expression
Total RNA was isolated using TRIzol reagent (Invitrogen) following the manufacturer’s instructions. Total RNA (20 μg) was resolved by a 1.25% denaturing agarose gels, transferred to Nytran nylon membrane (Schleicher & Schüell, Keene, NH), and hybridized with 32P-labeled cDNA probes as described before (42). The hybridized radioactivity was measured using Fuji BAS1800II instrument (Fuji Film). The mRNA levels were normalized to the 18S rRNA.
Statistical analysis
All values are expressed as the mean ± SEM. Statistical significance was determined by unpaired Student’s t test or one-way ANOVA with Fisher’s protected least significant difference test using Statview 5.0 software (SAS Institute Inc., Cary, NC). Differences were considered to be statistically significant at P < 0.05.
Results
Leptin decreases adipose tissue mass but does not affect energy expenditure in MKR mice
Leptin (10 μg/d) was administered to WT and MKR mice by sc implanted osmotic pumps for 14 d, resulting in a 10-fold increase in circulating leptin levels (Table 1). In response to treatment, both WT and MKR mice reduced their daily food intake by 22% and body weight by 10% (Table 1). Epididymal fat pad and whole-body fat mass were markedly reduced in the leptin-treated mice, but there were no significant differences in adipose tissue mass between the saline-treated WT and MKR mice (Table 1). The size of adipocytes was smaller after leptin treatment in both WT and MKR mice, and there were an increased number of cells with multilocular fat droplets resembling brown adipocytes (Fig. 1 and data not shown).
It has been shown that leptin replacement increases the metabolic rate and activity levels in leptin-deficient ob/ob mice (6). To determine whether leptin can increase energy expenditure in the MKR mice, we measured their metabolic rates by indirect calorimetry. No major differences were observed in resting and total oxygen consumptions, respiratory exchange ratio, and total activity at both 23 and 30 C in all groups of mice, suggesting that leptin treatment did not affect the metabolic rate in WT and MKR mice (Table 1 and data not shown). Importantly, the leptin-treated WT and MKR mice did not show a decrease in energy expenditure despite the decreased food intake.
Leptin improves the hyperglycemia and hyperinsulinemia in MKR mice
We examined the effect of leptin on blood glucose and serum insulin levels in MKR mice. The hyperglycemia in MKR mice was normalized after leptin treatment (Fig. 2A). The elevated serum insulin level in MKR mice was also significantly decreased (Fig. 2B). In addition, we examined whether the changes in an antidiabetic adipocytokine, adiponectin, was involved in these improvements. Serum adiponectin levels were not changed after leptin treatment (Table 1). These data demonstrated that leptin treatment improves diabetes in non-leptin-deficient and nonobese diabetic mice.
Because blood glucose and serum insulin levels were decreased by leptin in MKR mice, we anticipated that insulin sensitivity might be increased by leptin treatment in MKR mice. We performed insulin tolerance and glucose tolerance tests. In MKR mice, both insulin and glucose tolerance were improved by leptin treatment, suggesting that leptin increases insulin sensitivity in MKR mice (Fig. 2, C and D). In addition, fasting blood glucose was significantly lower in the leptin-treated MKR mice, compared with the saline-treated MKR and both groups of WT mice (Fig. 2D). It is not clear why leptin-treated MKR mice had very low fasting glucose levels. One possible explanation is that they are more sensitive to leptin particularly under fasting condition. Fasting insulin levels were higher in leptin-treated MKR mice, compared with leptin-treated WT mice (0.48 ± 0.13 vs. 0.25 ± 0.03 ng/ml, P = 0.055). Leptin also caused a significant reduction in elevated corticosterone levels in MKR mice (from 420 + 130 to 121 ± 22 ng/ml, P = 0.032). Both factors may contribute to a lower hepatic glucose production. It is also possible that leptin-treated MKR mice poorly adapt to fasting because of their smaller size and inability to down-regulate metabolic rate in response to caloric restriction.
The improvement of diabetes by leptin was not due to the reduced food intake in MKR mice
To exclude the possibility that the improvement of diabetes in MKR mice was due to the reduction of food intake, we pair fed one group of MKR mice treated with saline to the amount of food consumed by the MKR mice treated with leptin on the previous day. Pair feeding did not improve hyperinsulinemia, hypertriglyceridemia, and insulin and glucose tolerance in MKR mice (see Figs. 2, E and F, and 5, C and D, and data not shown). These results suggest that the antidiabetic effect of leptin was independent of caloric restriction in MKR mice.
Leptin enhances insulin sensitivity in liver and adipose tissues of MKR mice
The results of the insulin tolerance test suggested that leptin increased insulin sensitivity in MKR mice. We performed hyperinsulinemic-euglycemic clamps in MKR mice to assess insulin sensitivity in individual tissues. In the basal state (after 10 h of fasting), the leptin-treated MKR mice showed lower plasma glucose than the MKR mice treated with saline (MKR-saline: 241 ± 36 mg/dl; MKR-leptin: 126 ± 35 mg/dl). In addition, basal endogenous glucose production (EGP) was significantly reduced by 30% in the leptin-treated MKR mice (Fig. 3B). These findings suggest that the decreased basal endogenous glucose production partly contributes to the lower basal glucose levels.
The glucose infusion rate during the clamp was increased 4-fold (Fig. 3A), suggesting an increase in insulin sensitivity by leptin treatment. The clamp EGP was significantly lowered by 90% in the leptin-treated MKR mice (Fig. 3B). When the clamp EGP levels were compared with the basal level of each group, it was reduced by 40% in MKR mice treated with saline but further reduced by 90% in MKR mice treated with leptin. Thus, these results suggest that leptin increased hepatic insulin sensitivity in MKR mice.
During the clamp period, whole-body glucose uptake showed a tendency to increase in the leptin-treated MKR mice but did not reach statistical significance (P < 0.064; Fig. 3C). Whereas muscle glucose uptake was unaffected (Fig. 3D), the clamp glucose uptake in white adipose tissue (WAT) (Fig. 3E) and brown adipose tissue (Fig. 3F) was significantly increased by 4- and 10-fold, respectively, after leptin treatment. Thus, in MKR mice, leptin enhanced insulin-stimulated glucose uptake in adipose tissues, and this was associated with only a limited increase in whole-body glucose uptake.
Leptin inhibits insulin secretion in MKR mice
Previous studies have shown that MKR mice have -cell dysfunction (29, 30, 43). To examine whether leptin treatment improves -cell function, we performed pancreatic perfusion experiments in situ. MKR mice treated with saline showed an impaired glucose responsiveness as previously described (Fig. 4A) (30, 43). Insulin secretion under low glucose (1.4–5 mM) conditions showed no significant difference between saline- and leptin-treated MKR mice (Fig. 4B). However, under stimulatory (6–20 mM) glucose conditions, insulin secretion was inhibited in leptin-treated MKR mice (Fig. 4, A and B). Thus, leptin inhibited glucose-stimulated insulin secretion despite no reduction in total insulin content in the pancreas of MKR mice (data not shown). Thus, the improvement of hyperinsulinemia might be, in part, due to the leptin-inhibited insulin secretion and, in part, due to the reduction in insulin resistance.
Leptin improves lipid metabolism in MKR mice
Serum triglyceride (TG), which is abnormally elevated in MKR mice, was normalized after leptin treatment (Fig. 5A). Similarly, leptin reduced serum TG levels in WT mice (Fig. 5A). Serum FFA levels were decreased by leptin treatment in MKR mice (Fig. 5B). These effects of leptin on the circulating lipid levels were not due to the reduced caloric intake because pair feeding had no effect on serum TG and FFA levels in MKR mice (Fig. 5, C and D). Liver TG content was significantly decreased by leptin treatment in both WT and MKR mice (Fig. 5E). However, pair feeding also lowered liver TG content in MKR mice (Fig. 5E). In muscle, TG content in the saline-treated MKR mice was significantly increased as compared with the saline-treated WT mice (Fig. 5D). After leptin treatment, muscle TG content was normalized in MKR mice (Fig. 5F). Muscle TG content was not affected by pair feeding (Fig. 5F). Thus, leptin improved hyperlipidemia and reduced liver and muscle TG contents in MKR mice. Whereas caloric restriction may contribute to the lowering of liver TG content, the rest of the effects appear to be independent of leptin’s effect on food intake.
To demonstrate the mechanisms whereby leptin decreased serum and tissue TG levels, we measured TG secretion rates in WT and MKR mice. To quantify the TG secretion, the rate of increase in circulating TG was measured after inhibiting TG clearance with WR1339. TG secretion reflects whole-body TG production, with liver being the major source. Leptin treatment reduced whole-body TG production by 30% in WT and MKR mice (Fig. 6A), which correlates with the decrease in serum and liver TG contents after leptin treatment in both MKR and WT mice.
We also measured plasma TG levels after an oral lipid load to examine the TG clearance in WT and MKR mice treated with and without leptin. In the saline-treated WT and MKR mice, plasma TG peaked at 2–3 h after the lipid load (384 and 481 mg/dl at 2 h, respectively) and then gradually decreased (Fig. 6B). In the leptin-treated WT and MKR mice, plasma TG rose much less than the saline-treated mice, peaking at 161 and 193 mg/dl at 2 h after the lipid load, respectively. This was not due to poor TG uptake from the intestine because plasma TG levels increased similarly in the saline-treated WT and MKR mice in the absence of TG clearance after lipid load (Fig. 6C). Thus, leptin can enhance TG clearance in MKR mice, suggesting that leptin might be able to increase lipase activity.
Finally, we measured fatty acid oxidation in isolated skeletal muscle of WT and MKR mice with and without leptin infusion. Fatty acid oxidation was significantly inhibited in the saline-treated MKR mice as compared with WT mice. Leptin enhanced fatty acid oxidation in skeletal muscle in MKR mice but not WT mice (Fig. 6D). These data suggested that the enhanced muscle fatty acid oxidation led to the reduction of TG contents in MKR mice after leptin treatment.
All these effects might contribute to the improvement of lipotoxic conditions in the MKR mice.
Molecular mechanisms whereby leptin reduced glucose output and TG content in liver
To explore the molecular mechanisms whereby leptin inhibited glucose production and decreased TG accumulation in liver, we measured the expression of several genes related to glucose and lipid metabolism. The gene expression of glucose-6-phosphatase (G6Pase), which is one of the enzymes that regulates gluconeogenesis, was significantly decreased after leptin treatment in MKR mice (Fig. 7A). In addition, we found that the gene expression of GLUT-2, a low-affinity glucose transport protein that facilitates either glucose uptake or efflux from liver, depending on the nutritional state (44), was significantly reduced after leptin treatment (Fig. 7B). Because the glucose transporters are facilitative transporters, flux of glucose is dependent on a gradient, and therefore they are involved in both influx and efflux of glucose from the tissues. Thus, a reduction in GLUT2 expression could also explain the reduced glucose output from the liver after leptin treatment. Thus, the decreased hepatic glucose output by leptin treatment could be attributed to the reduction in both gluconeogenesis and glucose efflux from the liver.
We demonstrated that the expression of the gene for sterol-regulatory-element-binding protein-1c (SREBP-1c), a transcription factor that globally activates pathways of lipid biosynthesis (45), was significantly reduced in MKR mice after leptin treatment (Fig. 7C). In parallel, the expression of fatty acid synthase (FAS) and stearoyl-CoA desaturase-1 (SCD-1) were inhibited by leptin treatment (Fig. 7, D and F). Furthermore, CD36 gene expression, a fatty acid transporter, was also decreased in the leptin-treated MKR mice. Thus, the reduction of TG levels in liver could be due to the reduction of lipid synthesis and fatty acid uptake.
Leptin enhanced AMPK activity in skeletal muscle of MKR mice
Leptin has been shown to selectively stimulate phosphorylation and activation of the 2 catalytic subunit of AMPK, an enzyme that has been found to be a principal mediator of the leptin effects on fatty acid oxidation in skeletal muscle (8). Because the TG content in skeletal muscle was decreased and fatty acid oxidation was increased in muscle by leptin, we studied AMP kinase activity in muscle. In skeletal muscle, AMP kinase activity was 3-fold increased in the leptin-treated MKR mice (Fig. 7G). In addition, the phosphorylation of AMPK also was increased 1.5-fold by leptin in MKR mice (Fig. 7H). We also checked the phosphorylation of ACC, which is phosphorylated and inhibited by AMPK. The phosphorylation of ACC was increased by leptin in skeletal muscle of MKR mice (data not shown). These results suggest that the leptin-enhanced fatty acid oxidation in skeletal muscle was due to the increased AMPK activity.
Because dominant-negative IGF-I receptor also inhibits insulin receptor signaling (29) and hyperinsulinemic-euglycemic clamp data showed no changes in insulin-stimulated muscle glucose uptake in MKR mice (Fig. 3D), we did not expect to see any changes in insulin signaling in skeletal muscle of MKR mice after leptin treatment. To confirm this hypothesis, we measured phosphorylation of Akt, a downstream mediator of insulin-stimulated glucose uptake (46), in skeletal muscle of the saline- and leptin-treated WT and MKR mice. In WT mice, leptin increased the level of Akt phosphorylation, suggesting an increase in insulin signaling (Fig 8, A and B). In MKR mice, Akt phosphorylation was dramatically lower than in WT mice and was not affected by leptin treatment. Muscle GLUT4 protein levels were comparable in WT and MKR mice and did not change in response to leptin treatment (Fig. 8A). Thus, in skeletal muscle of MKR mice, leptin increased AMPK activity but had no effects on Akt phosphorylation or GLUT4 protein levels.
Discussion
MKR mice are a model for type 2 diabetes. They develop insulin resistance initially in muscle because of genetically impaired IGF-I and insulin receptor signaling (29), which leads to insulin resistance in liver and fat. Subsequently pancreatic -cell dysfunction develops, as demonstrated by the loss of first-phase insulin secretion. This event is associated with the appearance of diabetes (29). Unlike either genetically leptin-deficient or lipodystrophic diabetes models, MKR mice have normal amounts of adipose tissue and serum leptin levels (30, 31).
This study clearly demonstrates that leptin treatment improves insulin resistance and hyperglycemia in a specific mouse model of type 2 diabetes. Leptin treatment enhanced hepatic insulin sensitivity of the MKR mice by suppression of hepatic glucose production during the hyperinsulinemic-euglycemic clamp. Additionally, leptin reduced gene expression of key regulator of glucose metabolism such as G6Pase, a gluconeogenic enzyme in liver. All together, these findings suggest that leptin decreased glucose production through the inhibition of gluconeogenic enzymes. In previous studies, it has been reported that insulin action on hepatic glucose output in normal rats was enhanced by leptin (11, 47). Whereas it was also shown that the suppression of hepatic glycogenolysis primarily contributed to the decreased glucose production, gluconeogenesis was increased and accounted for the majority of glucose production. These events were accompanied by an increase in G6Pase and phosphoenolpyruvate carboxykinase gene expression. The discrepancy between the two studies might be due to differences in the regulation of gluconeogenesis affected by leptin between normal and diabetic animals or different responses of different animal species. In this study, it is also demonstrated that GLUT2 gene expression was decreased after leptin treatment. The liver senses hepatoportal glucose levels in a GLUT2-dependent manner and adjusts glucose influx and efflux to maintain euglycemia (48). Because GLUT2 expression was decreased by leptin treatment, glucose flux in both directions in the liver might be reduced. These events correlate with the reduction of gluconeogenesis. In addition, the decreased glucose influx could induce the reduction of fatty acid synthesis. It is possible that the leptin-suppressed GLUT2 gene expression could be secondary to other effects induced by prolonged leptin treatment (49). Thus, the reduction of both gluconeogenesis and efflux of glucose from liver could lead to the decreased hepatic glucose output after leptin treatment. Because the effect of leptin on hepatic glucose flux is largely mediated by hypothalamus (47, 50, 51), our findings might also be primarily related to the hypothalamic regulation. A recent study reported that the antidiabetic effect of leptin in mice with congenital lipodystrophy is mainly ascribed to its action on the liver (51). Our data also suggest that the improvement of liver function is important in the treatment of diabetes by leptin.
Unlike the liver, there was no improvement of insulin-stimulated glucose uptake in skeletal muscle. Previous studies have shown that chronic or acute leptin treatment in either normal or obese rodents increased glucose uptake and insulin sensitivity in skeletal muscle (7, 9). However, in the MKR mouse model, both the insulin and IGF-I receptor signaling pathways in skeletal muscle are genetically blocked, and therefore, not surprisingly, no leptin effect on glucose uptake could be detected.
A major cause of type 2 diabetes is a progressive insulin resistance of peripheral tissues, followed by a defect in insulin secretion in the islets of Langerhans (52). MKR mice showed islet cell hyperplasia, which resulted from severe insulin resistance in skeletal muscle, liver, and adipose tissues (29, 30, 43). Our previous studies demonstrated that the improvement of insulin sensitivity caused by peroxisomal proliferator-activated receptor- agonist and muscle CD36 overexpression led to the normalization of pancreatic -cells function in MKR mice (30, 43). However, in the present study, leptin did not improve the insulin responses of pancreatic -cells despite its improvement of insulin sensitivity. Instead, leptin inhibited glucose-stimulated insulin secretion. Leptin was shown to inhibit acutely glucose-stimulated insulin secretion in isolated islets from ob/ob mice (53). Similarly, acute physiological increases in plasma leptin levels inhibited glucose-simulated insulin secretion in conscious rats (54). Furthermore, hyperleptinemia achieved by adenovirus-leptin gene therapy had similar effects on insulin secretion, which could be explained by the depletion of lipids from the islets (55). It is likely that in the MKR mouse model, the effect of chronic leptin treatment on inhibition of insulin secretion is stronger than the acute glucose-stimulated insulin secretion.
Leptin treatment improved the hyperlipidemia of MKR mice by enhancing TG clearance and inhibiting TG secretion. Because leptin has been shown to prevent the accumulation of lipid content in nonadipose tissues in normal and lipodystrophic mice (21, 56, 57), the elevated TG levels in liver and skeletal muscle of MKR mice were significantly decreased after leptin treatment. In skeletal muscle, the reduction of TG contents was most likely secondary to the increased fatty acid oxidation, which was derived by the stimulation of AMPK kinase activity. In contrast, in liver, leptin apparently inhibited both lipid biosynthesis and fatty acid uptake by inhibiting gene expression of SREBP-1c, FAS, SCD-1, and CD36. These events might contribute to the reduction of liver TG content after leptin treatment. Our data suggest that a decreased caloric intake may also contribute to the reduction of liver TG content in the leptin-treated MKR mice. In addition to the lipid effects in nonadipose tissues, we observed a reduction in the amount of WAT. Hyperleptinemia in normal rats, induced by means of adenovirus-mediated gene transfer of the leptin gene, has reported a marked depletion of body fat content without increasing circulating FFAs (58). The effect of leptin on the transformation fat-storing adipocytes into fat-burning cells could be related to the depletion of body fat mass (59). This effect on WAT included the reduction in the gene expression of lipogenic enzymes and the increased phosphorylation of AMPK. We did not assess the mechanisms whereby leptin decreases the fat depot, but we can expect that similar molecular events might occur in WAT of MKR mice because we observed an increase in the number of cells with multilocular fat droplets resembling brown adipocytes.
Our previous studies using the MKR mouse model showed that the improvement in lipotoxicity (but not glucotoxicity) led to an improvement in insulin sensitivity and hyperglycemia (30, 31, 43). The improvement of diabetes after leptin treatment could also be due to the improvement of lipotoxicity in MKR mice. Namely, leptin enhanced lipid oxidation in skeletal muscle and apparently inhibited both lipid biosynthesis and fatty acid uptake in liver, leading to the reduction of lipid deposition in both tissues, thereby improving lipotoxicity and improving insulin sensitivity and diabetes.
It has been reported that leptin could be used as an adjunct of insulin therapy in streptozotocin-induced, insulin-dependent diabetic mice (60) and restore euglycemia and glucose turnover in streptozotocin-induced, insulin-dependent diabetic rats (61). Furthermore, in several clinical trials of leptin treatment, it has been shown that leptin improved the glycemic control, increased insulin sensitivity, and decreased TG levels in patients with lipodystrophy and leptin deficiency (19, 20, 62). More recently it has been reported that leptin administration improved fasting hyperglycemia, hyperinsulinemia, and glucose and insulin tolerances in patients with Rabson-Mendenhall syndrome, which show severe insulin resistance with presumed insulin receptor mutations, low levels of serum TG, and normal serum leptin levels (63).
Obesity is emerging as a worldwide epidemic and the number of obese type 2 diabetic patients is consequently increasing. Circulating leptin levels are strongly correlated with fat mass. In general, obese patients and rodents have elevated leptin levels (24, 64). It has been shown that leptin resistance exists in hyperleptinemic states, although the mechanisms involved in leptin resistance remain unclear. However, there was considerable variability in the effect of leptin treatment in obese patients (26). Furthermore, a large proportion of lean patients with type 2 diabetes have leptin levels in normal range (23, 25). Because leptin is currently available for clinical studies, our findings might provide supportive evidence for the use of leptin as a therapy for some type 2 diabetic patients. However, because chronic hyperleptinemia could induce hypertension (65) and reversible unresponsiveness to glucose in -cells (55) in animal models, further studies are necessary to see whether long-term leptin therapy has any side effects
In summary, the present study demonstrates that leptin can reverse diabetes by a combination of improvement in glucose and lipid metabolism in MKR mice having normal circulating leptin levels. These findings are independent of decreased food intake, suggesting that leptin could be a useful form of treatment for not only patients with lipodystrophic diabetes but also type 2 diabetes patients without leptin resistance.
Footnotes
This work was supported in part by a Mentor grant (7-02-MN-15) and a Research grant (7-02-RA-49) from the American Diabetes Association awarded to D.L. and by the Canadian Diabetes Association grants (to M.B.W.). M.B.W. is a Canadian Institutes for Health Research Investigator.
Abbreviations: ACC, Acetyl coenzyme A carboxylase; AMPK, AMP-activated protein kinase; EGP, endogenous glucose production; FAS, fatty acid synthase; FFA, free fatty acid; GLUT, glucose transporter protein; G6Pase, glucose-6-phosphatase; MKR, transgenic overexpression of a skeletal muscle dominant-negative IGF-I receptor with a lysine-to-arginine amino acid; RER, respiratory exchange ratio; SCD-1, stearoyl-CoA desaturase-1; SREBP-1c, sterol-regulatory-element-binding protein-1c; TG, triglyceride; WAT, white adipose tissue; WT, wild type.
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