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Endocrinology Vol. 145, No. 9 4094-4102
Copyright © 2004 by The Endocrine Society

Increased Insulin Sensitivity in Paternal Gnas Knockout Mice Is Associated with Increased Lipid Clearance

Min Chen, Martin Haluzik, Nicole J. Wolf, Javier Lorenzo, Kelly R. Dietz, Marc L. Reitman and Lee S. Weinstein

Metabolic Diseases Branch (M.C., N.J.W., J.L., L.S.W.) and Diabetes Branch (M.H., K.R.D., M.L.R.), National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892

Address all correspondence and requests for reprints to: Dr. Lee S. Weinstein, Metabolic Diseases Branch, National Institute of Diabetes and Digestive and Kidney Diseases/National Institutes of Health, Building 10, Room 8C101, Bethesda, Maryland 20892-1752. E-mail: leew{at}amb niddk.nih.gov.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The G protein {alpha}-subunit Gs{alpha} is required for hormone-stimulated cAMP generation. The Gs{alpha} gene Gnas is a complex gene with multiple imprinted gene products. Mice with heterozygous disruption of the Gnas paternal allele (+/p–) are partially Gs{alpha} deficient and totally deficient in XL{alpha}s, a neuroendocrine-specific Gs{alpha} isoform that is expressed only from the paternal Gnas allele. We previously showed that these mice are hypermetabolic and lean and have increased insulin sensitivity. We now performed hyperinsulinemic-euglycemic clamp studies, which confirmed the markedly increased whole body insulin sensitivity in +/p– mice. +/p– mice had 1.4-, 7- and 3.8-fold increases in insulin-stimulated glucose uptake in muscle and white and brown adipose tissue, respectively, and markedly suppressed endogenous glucose production from the liver. This was associated with increased phosphorylation of insulin receptor and a downstream effector (Akt kinase) in both liver and muscle in response to insulin. Triglycerides cleared more rapidly in +/p– mice after a bolus administered by gavage. This was associated with decreased liver and muscle triglyceride content and increased muscle acyl-CoA oxidase mRNA expression. Resistin and adiponectin were overexpressed in white adipose tissue of +/p– mice, although there was no difference in serum adiponectin levels. The lean phenotype and increased insulin sensitivity observed in +/p– mice is likely a consequence of increased lipid oxidation in muscle and possibly other tissues. Further studies will clarify whether XL{alpha}s deficiency is responsible for these effects and if so, the mechanism by which XL{alpha}s deficiency leads to this metabolic phenotype.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE HETEROTRIMERIC G PROTEIN {alpha}-subunit Gs{alpha} is a ubiquitously expressed protein that couples seven transmembrane receptors to adenylyl cyclase and is required for stimulation of intracellular cAMP generation in response to many hormones and other extracellular signals (1). Gs{alpha} mediates the cAMP response to glucagon and ß-adrenergic agonists, hormones that counteract the actions of insulin, and cAMP is known to mitigate the metabolic actions of insulin in several cellular contexts (2, 3, 4, 5). In adipocytes, Gs{alpha} is also critical for the lipolytic response to ß-adrenergic agonists, which is mediated through cAMP (6, 7). Heterozygous null Gs{alpha} mutations lead to Albright hereditary osteodystrophy, a syndrome that is characterized by obesity as well as other skeletal and neurological abnormalities (1, 8, 9).

The gene encoding Gs{alpha} (GNAS at 20q13.3 in humans; Gnas on chromosome 2 in mice) is a very complex gene with multiple promoters and first exons that splice onto a common set of downstream exons (exons 2–13) to generate at least three distinct protein products (1). Another level of complexity results from the fact that these promoters are imprinted, leading to some gene products being expressed from the maternal allele and others being expressed exclusively from the paternal allele. The most downstream promoter and first exon generates transcripts encoding Gs{alpha}. Gs{alpha} is biallelically expressed in most tissues but is expressed primarily from the maternal allele in some tissues, such as renal proximal tubules, thyroid, and pituitary (10, 11, 12). This explains why maternally, but not paternally, inherited Gs{alpha} mutations results in PTH, TSH, and gonadotropin resistance (13). Other GNAS/Gnas gene products include NESP55, a chromogranin-like protein expressed exclusively from the maternal allele, and XL{alpha}s, a Gs{alpha} isoform expressed exclusively from the paternal allele (14, 15, 16). Both proteins are expressed primarily in neuroendocrine tissues, and little is known about their biological function (17, 18, 19, 20, 21, 22), although it has been shown recently that XL{alpha}s is capable of mediating receptor-stimulated cAMP generation (23). A fourth promoter and first exon (exon 1A) generates RNA transcripts only from the paternal allele; these transcripts are probably not translated (24, 25).

We previously generated mice with an insertion in Gnas exon 2 that disrupts the Gs{alpha} coding sequence (26). The homozygotes were embryonically lethal and heterozygotes with disruption of the maternal (m–/+) or paternal (+/p–) allele had distinct phenotypes as would be predicted by Gnas imprinting. Interestingly, the m–/+ mice had decreased energy expenditure and activity levels leading to obesity, whereas the +/p– mice were hypermetabolic, hyperactive, and very lean (27). M–/+ and +/p– mice had decreased and increased urinary norepinephrine excretion, respectively, suggesting that perhaps the opposite metabolic phenotypes are due to opposite changes in sympathetic nervous system activity. Interestingly, both groups of animals had improved glucose tolerance and increased insulin sensitivity, although this effect was much greater in the lean +/p– mice (28).

In the present study, we examined the metabolic phenotype in +/p– mice in more detail. Using hyperinsulinemic-euglycemic clamp studies, we show that +/p– mice have increased insulin sensitivity in the liver, skeletal muscle, brown adipose tissue (BAT), and white adipose tissue (WAT) associated with increased phosphorylation of the insulin receptor (IR) and the downstream effector kinase Akt in both liver and muscle. We also demonstrate that +/p– mice have significantly accelerated triglyceride clearance after oral lipid load and increased expression of the lipid oxidation enzyme acyl-CoA oxidase (AOX) in skeletal muscle. Although both resistin and adiponectin are overexpressed in WAT of +/p– mice, there was no difference in serum adiponectin levels. We suggest that increased lipid clearance and oxidation, which may be the result of increased sympathetic nervous system activity, leads to leanness and increased insulin sensitivity in +/p– mice. Loss of XL{alpha}s expression is a likely candidate to be the underlying molecular defect that leads to the +/p– metabolic phenotype.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Mice with insertion of a neomycin resistance cassette into exon 2 of Gnas were previously created by targeted mutagenesis (26). Male mutants, which were in a CD-1 background, were mated to wild-type CD-1 females to generate +/p– mice. Animals were maintained on a 12-h light, 12-h dark cycle (0600/1800 h) and a standard pellet diet (NIH-07, 5% fat by weight). All experiments were performed on 12-wk-old male mice and wild-type littermates were used as controls. For ß3-adrenergic agonist studies, daily ip injections of CL316243 (1 mg/kg body weight per day) (Sigma, St. Louis, MO) or saline vehicle were administered for 4 wk in normal CD-1 mice (Charles River, Wilmington, MA). Animal experiments were approved by the National Institute of Diabetes and Digestive and Kidney Diseases Animal Care and Use Committee.

In vivo insulin response to glucose
Mice were fasted overnight and given an ip glucose injection (3 mg/g body weight) after avertin administration (0.25 mg/g body weight). Blood was collected in heparinized capillary tubes from the retroorbital vein, and plasma was collected after centrifugation. Insulin was measured by RIA and glucose was measured by glucometer.

Biochemical and hormonal assays
Plasma glucose was measured using a Glucometer Elite (Bayer, Elkhart, IN). Plasma insulin (SRI-13K; Linco Research, St. Charles, MO), serum corticosterone (kit 07120102; ICN Pharmaceuticals, Orangeburg, NY), serum adiponectin (kit MADP-60HK; Linco), and serum glucagon (kit GL-32K; Linco) were measured by RIA.

Surgery and animal handling
Catheter insertion was adapted from MacLeod and Shapiro (29). Operations were carried out under ketamine (100 mg/kg; Fort Dodge Animal Health, Fort Dodge, IA) and xylazine (10 mg/kg; Phoenix Scientific, St. Joseph, MO) anesthesia. The silastic catheter (inner diameter, 0.30 mm; outer diameter, 0.64 mm, 508–001, Dow Corning, Midland, MI), filled with heparin solution (100 United States Pharmacopeia U/ml in 0.9% NaCl) was inserted via a right lateral neck incision, advanced into the superior vena cava via the right internal jugular vein, and sutured in place. The distal end of the catheter was knotted, tunneled sc, exteriorized first at the dorsal cervical midline, and then further tunneled sc and exteriorized in the dorsal midline 2 cm above the tail. A silk suture was fastened around the catheter at the neck site. On the day of the clamp, the catheter was externalized by pulling the suture through the dorsal cervical incision site.

Hyperinsulinemic-euglycemic clamp
The clamps were performed 4–6 d after catheter placement as described previously (30, 31). Clamps began at 0700 h, after 12 h of fasting. Mice were placed in a restrainer (552-BSRR; PlasLabs, Lansing, MI), and the catheter was externalized. The tip of the tail was cut before the start of the first infusion, and all subsequent blood samples were drawn from this site. Blood was collected into heparinized microcapillary tubes (Fisher Scientific, Pittsburgh, PA) and centrifuged for 10 sec to obtain plasma. Basal endogenous glucose production rate was determined by continuously infusing [3-3H]glucose (3 µCi bolus, then 0.02 µCi/min, 740 GBq/mmol; NET 331C; NEN Life Science Products, Boston, MA). Samples for determination of plasma [3-3H]glucose concentration were taken after 90 and 115 min of basal infusion. Basal insulin concentration was measured using 10 µl of the 90-min sample. After 120 min of basal [3-3H]glucose infusion, the hyperinsulinemic-euglycemic clamp was begun with a primed continuous infusion of human insulin (300 mIU/kg bolus over 3 min, then 2.5 mIU/kg/min; Humulin R; Eli Lilly, Indianapolis, IN). Plasma glucose was measured at 15- and 10-min intervals during the first and second hour of the clamp, respectively, and 20% glucose was infused at a rate adjusted to keep plasma glucose at approximately 110 mg/dl.

Insulin-stimulated whole-body glucose uptake was measured using a primed continuous infusion of [3-3H]glucose (10 µCi bolus, 0.1 µCi/min) throughout the clamps. Insulin-stimulated glucose uptake in tissues was measured using a bolus injection of 2-deoxy-D-[1-14C]glucose (10 µCi in 5 µl of 0.9% saline, 2.1 GBq/mmol; NEC 495; NEN Life Science Products) at 70 min after the start of the insulin infusion. Blood samples (20 µl) were withdrawn at 80, 85, 90, 100, 110, and 120 min after start of the insulin infusion to measure plasma 3H and 14C. Clamp insulin levels were measured in 5 µl plasma from the 110-min time point. All infusions were performed using a microdialysis pump (model CMA 102; CMA/Microdialysis, Acton, MA). Gastight syringes (10 µl; Hamilton Co., Reno, NV) were used for bolus injections. After 120 min of insulin infusion, animals were anesthetized with ketamine/xylazine solution. Tissues were immediately removed, frozen in liquid nitrogen, and stored at –70 C. The total volume of blood withdrawn, which was not replaced, was approximately 300 µl per animal.

In vivo glucose flux analysis
For the determination of plasma [3-3H]glucose and 2-deoxy-D-[1-14C]glucose concentrations, plasma was deproteinized with ZnSO4 and Ba(OH)2, dried under vacuum at room temperature to remove 3H2O, resuspended in water, and counted in BioSafe II scintillation fluid (Research Products International, Mount Prospect, IL) using a Beckman LS60001C (Beckman Coulter, Inc., Fullerton, CA) with correction for background, counting efficiency, and channel cross-over. For determination of tissue 2-deoxy-D-[1-14C]glucose-6-phosphate, tissue samples were homogenized in distilled water (50 mg tissue/500 µl water) and an aliquot was counted to determine total 14C. The remainder of the homogenate was subjected to anion-exchange chromatography (model 731-6211, Bio-Rad Laboratories, Hercules, CA) to separate nonmetabolized 2-deoxyglucose (neutral 14C counts eluted with 6 ml distilled water) from 2-deoxyglucose-6-phosphate (anionic 14C counts eluted with 6 ml of 0.2 M formic acid/0.5 M ammonium acetate).

Calculations
Basal endogenous glucose production was calculated as the ratio of the preclamp [3-3H]glucose infusion rate (disintegrations per minute per minute) to the specific activity of the plasma glucose (mean of the 90 and 115 min preclamp values in disintegrations per minute per micromole). Clamp whole-body glucose uptake was calculated as the ratio of the [3-3H] glucose infusion rate (disintegrations per minute per minute) to the specific activity of plasma glucose (disintegrations per minute per micromole) during the last 30 min of the clamp (mean of the 90- to 120-min clamp samples). Whole-body glycolysis was determined from the rate of increase in plasma 3H2O determined by linear regression using the 90- to 120-min points. Plasma 3H2O concentrations were measured from the difference between nondried vs. dried plasma 3H counts. Clamp endogenous glucose production was determined by subtracting the average glucose infusion rate in the last 30 min of clamp from the whole-body glucose uptake. Whole-body glycogen synthesis was estimated by subtracting the whole-body glycolysis from the whole-body glucose uptake, which assumes that glycolysis and glycogen synthesis account for the majority of insulin-stimulated glucose uptake (32). Tissue glucose uptake was calculated from the plasma 2-deoxy-D-[1-14C] glucose concentration profile (using plasma 14C counts at 80–120 min, the area under the curve was calculated by trapezoidal approximation) and tissue 2-deoxy-D-[1-14C] glucose-6-phosphate content as described previously (33).

Tissue triglyceride and glycogen content
Liver and muscle triglyceride contents were measured by solvent extraction followed by a radiometric assay for glycerol (34). To measure tissue glycogen content, tissue (~100 mg) was homogenized in 600 µl of 30% KOH and incubated at 97 C for 15 min. Cold 95% ethanol (3 ml) was added into each tube and incubated at –30 C for 1 h. After centrifugation at 3300 rpm for 30 min at 4 C, pellets were washed with cold 95% ethanol three times and dissolved in 200 µl distilled water. Samples were then incubated in 100 µl of solution containing 1 U/ml glucokinase, 50 mM triethanolamine hydrochloride (pH 9.0) 2 mM MgCl2, 1 mg/ml BSA, and 40 µM [{gamma}-32P]ATP at 30 C for 30 min, and then 100 µl of 2N HClO4 with 0.2 mM H3PO4 were added and samples incubated at 90 C for 40 min. After adding 50 µl of 100 mM ammonium molybdate and 50 µl of 200 mM triethylamine, samples were centrifuged at 3000 rpm for 30 min. Tissue glucose was measured as incorporation of {gamma}-32P ATP and calculated using a standard curve with various glucose concentrations. All reagents were purchased from Sigma.

Portal vein insulin injection and immunoblot analysis
Mice were fasted overnight and anesthetized by ip avertin (0.25 mg/g body weight). Insulin (100 µl of 15 µg/ml, Sigma) was injected via the portal vein. At 2 or 4 min after injection, liver and hind leg muscles were dissected and immediately frozen. Tissues were then homogenized using a Polytron in 50 mM Tris-HCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 150 mM NaCl, 1 mM EGTA, 1 mM phenylmethylsulfonyl fluoride, 1 mM Na3VO4, 1 mM NaF (pH 7.4) with protease inhibitor cocktail (Roche Biomedical, Indianapolis, IN). Tissue lysates were centrifuged in an Eppendorf microcentrifuge at 1100 rpm for 10 min at 4 C, and supernatants were used for immunoprecipitation. Protein concentrations were determined using the dye method (Bio-Rad). Immunoprecipitations were performed on tissue extracts (1 mg protein) using either an antiinsulin receptor (Transduction Laboratories, Lexington, KY) or an anti-Akt antibody (Upstate Biotechnology, Lake Placid, NY) as per manufacturer’s instructions, followed by immunoblot analysis using an antiphosphotyrosine (4G10, Upstate Biotechnology) or anti-Akt1 phospho-Ser473 antibody (Upstate Biotechnology), respectively. Antibody binding was determined by chemiluminescence (ECL kit; Amersham, Arlington Heights, IL), and bands were quantified using NIH Image version 1.55 software. To normalize for the amount of IR and Akt protein, the same blots were stripped and hybridized with antiinsulin receptor antibody or anti-Akt antibody, respectively.

Triglyceride clearance test
Clearance of triglycerides (400 µl peanut oil delivered by gavage) from the circulation was measured in mice after a 4-h fast. Blood was taken before gavage and hourly for 6 h after gavage and plasma triglycerides were measured (kit 337-B; Sigma).

RNA analysis
Total RNA was isolated from epididymal WAT using the TRIzol method (Life Technologies, Inc.-BRL, Gaithersburg, MD). Mouse adiponectin and resistin cDNA probes were generated by RT-PCR using WAT as a template and verified by restriction mapping. Adiponectin primers were: sense, 5'-AGAGAAGGGAGAGAAAGGAGATGC-3' and antisense, 5'-TGGTCGTAGGTGAAGAGAACGG-3'. Resistin primers were: sense, 5'-CCCTCCTTTTCCTTTTCTTCCTTG-3' and antisense, 5'-TTTTCTTCACGAATGTCCCACG-3'. Northern analysis was performed using 15 µg total WAT RNA per sample. Mouse adiponectin and resistin signals were quantified using a BAS1500 phosphor imager (Fuji, Tokyo, Japan) and normalized to 18S RNA, which was quantified by ethidium bromide using NIH Image version 1.55 software. The peroxisomal proliferator-activated receptor-{gamma} coactivator-1 (PGC-1) probe was a PCR product spanning nucleotides 1475–3009 of mouse PGC-1 (GenBank accession no. AF0499330). Probes for metabolic enzymes were previously described (35).

Statistical analysis
Data are expressed means ± SEM. Statistical significance between the groups was determined with SigmaStat (SPSS Inc., Chicago, IL) using paired Student’s t test (except where noted, in which case an unpaired t test was performed).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Anatomical and biochemical characteristics of paternal Gnas knockout mice
As reported previously, +/p– mice had significantly decreased body weight and body fat content as compared with control group (27) (data not shown). Whereas basal glucose levels of +/p– mice after overnight fasting were not different from control group, insulin levels were 2.5-fold lower (Table 1Go). Because low body weight can be a sign of adrenal insufficiency, we measured serum fasting corticosterone levels. Fasting corticosterone levels were similar in +/p– and wild-type mice (445 ± 84 ng/ml in +/p– mice vs. 461 ± 92 ng/ml in wild-type mice, n = 5 per group), ruling out adrenal insufficiency as an explanation for the +/p– metabolic phenotype. There were also no differences in fasting serum glucagon levels (24.6 ± 5.0 pg/ml in +/p– mice vs. 18.3 ± 0.9 pg/ml in wild-type mice, n = 5 per group).


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  		TABLE 1. Metabolic parameters in mice undergoing hyperinsulinemic-euglycemic clamp

 
To rule out the possibility that low insulin levels in +/p– mice result from a pancreatic islet cell defect in insulin secretion, we determined the first- and second-phase insulin response to an ip glucose bolus (3 mg/g) (Fig. 1Go). In this experiment glucose levels were lower than wild type before injection but subsequently followed a similar curve after glucose injection. Consistent with prior results, insulin levels were lower than normal at baseline. After injection the +/p– mice had a rapid rise in insulin levels that were similar or greater than wild type, and by 5 min after injection, both groups of mice had similar levels of insulin, which persisted until the end of the experiment (30 min). Based on these results, it is unlikely that +/p– mice have a defect in insulin secretion.



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  		FIG. 1. Glucose-stimulated insulin release in wild-type (+/+, triangles) and +/p– mice (squares). Plasma insulin (A) and plasma glucose (B) in overnight fasted, 12-wk-old male mice were determined before and 2, 5, 15, and 30 min after ip glucose administration (3 mg/g body weight). Values are expressed as mean ± SEM (n = 4 per group).

 
Paternal Gnas knockout mice have increased insulin sensitivity in liver, muscle, and adipose tissue
We performed hyperinsulinemic-euglycemic clamp studies to better characterize the insulin sensitivity of +/p– mice at an organ-specific level. Studies were performed after an overnight fast, and therefore the animals likely had low liver glycogen levels at the start of the clamp. The basal endogenous glucose production rate after overnight fasting was comparable in +/p– and wild-type mice (Fig. 2Go). The presence of a normal endogenous glucose production rate in the face of lower insulin levels in +/p– mice (Table 1Go) suggests that their livers are overly sensitive to insulin. Consistent with normal fasting endogenous glucose production, which primarily reflects the rate of gluconeogenesis by the liver, the steady-state mRNA levels of two gluconeogenic enzymes, phosphoenolpyruvate carboxykinase and glucose-6-phosphatase as well as PGC-1, which promotes the expression of gluconeogenic enzymes, were unchanged in the livers of +/p– mice (Table 2Go). Suppression of endogenous glucose production during the clamp measures the acute metabolic response of the liver to insulin. Clamp endogenous glucose production rate was completely suppressed in +/p– mice, whereas only 49% suppression was observed in wild-type mice (Fig. 2Go), indicating that the livers of +/p– mice have markedly greater insulin sensitivity than normal. In fact, the average absolute endogenous glucose production rate in +/p– mice was slightly negative, probably as a result of increased glycogen cycling in the liver.



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  		FIG. 2. Whole-body measures of glucose metabolism in wild-type (+/+) and +/p– mice during hyperinsulinemic-euglycemic clamp studies. Basal (preclamp) and clamp endogenous glucose production rate, glucose infusion rate, and rates of whole-body glucose uptake, glycolysis, and glycogen synthesis in 12-wk-old male wild-type (black bars) and +/p– mice (open bars) are shown as mean ± SEM (n = 6–7 per group). *, P < 0.05 vs. wild-type group. Details describing how these rates were calculated are described in Materials and Methods under Calculations.

 

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  		TABLE 2. mRNA expression of metabolic enzymes and PGC-1 in +/p– mice

 
Insulin-stimulated whole-body glucose uptake during the clamp is primarily a reflection of insulin sensitivity of skeletal muscle. The rate of whole-body glucose uptake was 54% greater in +/p– mice, compared with wild-type mice (Fig. 2Go), whereas muscle-specific glucose uptake was 70% greater in +/p– mice (Fig. 3Go). The increase in insulin-stimulated glucose uptake in muscle during the clamp was similar or slightly greater than the increase observed in isolated skeletal muscle from +/p– mice (28). Interestingly, the most pronounced differences in tissue insulin sensitivity were detected in adipose tissues. Glucose uptake in WAT and BAT of +/p– mice was 7- and 3.8-fold higher than in wild-type mice, respectively (Fig. 3Go). Most of the increase in whole-body glucose uptake was accounted for by an increase in whole-body glycolysis, with little or no increase in whole-body glycogen synthesis (Fig. 2Go). Consistent with the mutants having normal glycogen synthesis rates, we did not detect changes in liver or muscle glycogen content (Fig. 4Go, C and D). Also, the glucose infusion rate, which reflects both the rates of endogenous glucose production and uptake during the clamp, was significantly greater in +/p– mice (Fig. 2Go). In summary, hyperinsulinemic-euglycemic clamp studies revealed that +/p– mice have increased insulin sensitivity in liver, skeletal muscle, WAT, and BAT.



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  		FIG. 3. Tissue-specific glucose uptake rates in wild-type (+/+) and +/p– mice during hyperinsulinemic-euglycemic clamp studies. Glucose uptake rates in gastrocnemius muscle, WAT, and BAT during hyperinsulinemic-euglycemic clamp studies in 12-wk-old male wild-type (black bars) and +/p– mice (open bars) are shown as mean ± SEM (n = 5–7 per group). *, P < 0.05 vs. wild-type group. Details describing how these rates were calculated are described in Materials and Methods under Calculations.

 


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  		FIG. 4. Tissue triglyceride and glycogen content. A, Liver triglyceride content (micromoles per gram of tissue) was measured in liver (A) and skeletal muscle (B). Liver (C) and muscle (D) glycogen content (nanomoles per gram of tissue) in 12-wk-old male wild-type (+/+; black bars) and +/p– mice (open bars) is shown as mean ± SEM (n = 4–6 per group). *, P < 0.05 vs. wild-type group by unpaired t test for liver and paired t test for muscle.

 
Paternal Gnas knockout mice have increased insulin receptor and Akt phosphorylation in response to insulin in liver and skeletal muscle
To determine whether the increased in vivo insulin sensitivity in +/p– mice was associated with altered responsiveness of the proximal insulin signaling pathway, we examined tyrosine phosphorylation of IR and Ser473 phosphorylation of the downstream effector Akt in response to insulin administration in vivo (Fig. 5Go). Immunoblot analysis with anti-IR and anti-Akt antibodies demonstrated that expression levels of both proteins were unaffected in +/p– mice. Moreover, there were negligible levels of IR or Akt phosphorylation after injection of saline vehicle (data not shown), indicating that there was no basal signaling in the absence of insulin. However, IR phosphorylation at 2 min after insulin injection was more than 2-fold higher in liver and more than 5-fold higher in muscle of +/p– mice, compared with their wild-type littermates, indicating that in +/p– mice, insulin signaling is enhanced at the level of the IR in these tissues. At 4 min IR phosphorylation in muscles was increased by only 76% in +/p– mice. IR activation leads to serine/threonine phosphorylation and activation of Akt kinase through activation of phosphatidylinositol-3 kinase, and this pathway is required for insulin-stimulated glucose uptake. Consistent with higher levels of IR phosphorylation, phosphorylation of Akt1-Ser473 in response was higher in liver and muscle of +/p– mice. In muscle, the differences in IR phosphorylation were greatest at 2 min, whereas the differences in Akt phosphorylation were greater at 4 min.



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  		FIG. 5. Insulin-stimulated IR and Akt phosphorylation in liver and skeletal muscle. A, Twelve-week-old male +/p– mice and wild-type littermates (+/+) were killed 2 min after insulin administration into the portal vein, and IR tyrosine phosphorylation was assessed in liver (left) and muscle (right). After immunoprecipitation with an anti-IR antibody, immunoblotting was performed with an antiphosphotyrosine (top of each panel) or anti-IR antibody (bottom of each panel). To the right of each panel is the relative IR phosphorylation in +/p– mice expressed as the percentage of IR phosphorylation in +/+ littermates (mean ± SEM, *, P < 0.05 by one-tailed t test). B, The same tissue extracts were immunoprecipitated with anti-Akt antibody followed by immunoblotting with anti-Akt1 Ser473 (top of each panel) or anti-Akt antibodies (bottom of each panel). For muscle, results at both 2 and 4 min are shown.

 
Paternal Gnas knockout mice have accelerated triglyceride clearance
It is now well established that insulin sensitivity is generally inversely related to the levels of triglycerides and other lipids in serum and nonadipose tissues (36, 37, 38). We previously showed that circulating levels of cholesterol and triglycerides were significantly lower than normal in +/p– mice (27). We now measured liver and muscle triglyceride content and found both to also be lower than normal in +/p– mice (Fig. 4Go, A and B). To examine triglyceride metabolism more directly, we measured serum triglyceride levels before and after gavage of 400 µl peanut oil (Fig. 6Go). Consistent with previous results (27), basal triglyceride levels after 4 h of fasting were lower in +/p– mice than in wild-type mice (P = 0.051). Triglyceride clearance during the 6-h period after gavage was markedly accelerated in +/p– mice with an areas under the curve in +/p– mice that were 46% less than those of wild-type mice (P < 0.05). This is consistent with previous results showing that +/p– have an increased resting metabolic rate (27).



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  		FIG. 6. Triglyceride clearance in wild-type and +/p– mice. Serum triglycerides were measured at baseline (time 0) and hourly for 6 h after administration of peanut oil by gavage in 12-wk-old male wild-type (+/+; black circles) and +/p– mice (white circles). Values are shown as mean ± SEM (n = 4–5 per group). The areas under curves (mean ± SEM) are 1160 ± 237 for wild-type and 630 ± 73 for +/p– mice (P < 0.05).

 
We next examined the expression of enzymes required for lipid oxidation to determine whether increased expression of these enzymes might account for the apparent increase in lipid metabolism observed in these mice. mRNA expression of the enzymes AOX and carnitine palmitoyltransferase (CPT1) was unaffected in the livers of +/p– mice (Table 2Go). In skeletal muscle CPT1 mRNA levels were unaffected, whereas AOX mRNA levels were increased in +/p– mice (Table 2Go). We detected no changes in PGC-1 mRNA levels in skeletal muscle, liver, or WAT (Table 2Go).

Adiponectin and resistin are overexpressed in WAT of paternal Gnas knockout mice
Adipocytes have been shown to produce and secrete various circulating factors that have effects on insulin sensitivity and lipid metabolism. Resistin was originally described as a factor secreted from adipose tissue in proportion to the level of adiposity that leads to insulin resistance (39), although more recent studies suggest that resistin expression is not correlated with adiposity and question its importance in the regulation of insulin action (40, 41, 42, 43). Adiponectin is another adipose-derived factor that is expressed in inverse proportion to adiposity and stimulates lipid metabolism and insulin sensitivity in liver and muscle through activation of AMP kinase (44). To determine the potential role of these adipocytokines in the +/p– metabolic phenotype, we measured the levels of their mRNAs in epididymal WAT from +/p– mice and littermate controls. As shown in Fig. 7AGo, resistin mRNA levels were approximately 3-fold higher in +/p– mice, compared with littermate controls, whereas adiponectin mRNA levels were approximately 3-fold higher in +/p– mice. Therefore, in this model reduced adiposity is associated with increased expression of both adiponectin and resistin.



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  		FIG. 7. Adiponectin and resistin expression in WAT tissue. A, Results of Northern analysis of total WAT RNA (15 µg/lane) from male +/p– mice and their +/+ littermates after hybridizing with resistin- (top panel) and adiponectin-specific cDNA probes (middle panel). The 18S rRNA bands after ethidium bromide staining are shown in the bottom panel. To the right is the relative expression of resistin and adiponectin in +/p– WAT expressed as the percentage of expression in +/+ littermates (mean ± SEM, P < 0.05 for both by t test). B, Adiponectin expression in 8-wk-old male wild-type CD1 mice after 4 wk of treatment with CL316243 (CL, +) or saline vehicle (–). Adiponectin expression in CL-treated mice expressed as the percent of expression in untreated mice is shown on the right (mean ± SEM).

 
Despite the apparent overexpression of adiponectin in WAT of +/p– mice, these mice do not appear to have increased circulating levels of adiponectin (serum levels 7.36 ± 0.57 µg/ml in +/p– mice vs. 8.65 ± 0.93 µg/ml in wild-type mice, n = 5 pairs). This may reflect the fact that +/p– mice have significantly reduced WAT mass, which compensates for the overexpression of adiponectin per unit of WAT mass, as was found in mice lacking acyl-CoA/diacylglycerol acyltransferase 1 (45). In any case, it appears unlikely that adiponectin overexpression accounts for the increased lipid metabolism and insulin sensitivity in +/p– mice because there is no increase in circulating levels. TNF{alpha} mRNA levels in WAT were not significantly increased in +/p– mice, and we were unable to detect TNF{alpha} in serum of either mutant or wild-type mice using currently available assays (data not shown).

We previously showed that urinary norepinephrine levels were elevated in +/p– mice, and therefore the metabolically active adipocytes in the mice may be due to increased sympathetic stimulation (27). If this is the case, then the increased adiponectin expression that we observed in WAT in these mice may be a consequence of chronic ß-adrenergic stimulation. To test this hypothesis, we treated wild-type CD1 mice for 4 wk with the ß3-adrenergic agonist CL316243 and measured the effects on adiponectin mRNA expression in WAT (Fig. 7BGo). Chronic ß-adrenergic stimulation resulted in a significant increase in adiponectin expression, which supports the hypothesis that increased adiponectin in +/p– WAT tissue may be secondary to chronically elevated levels of sympathetic activity.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We previously demonstrated that disruption of the Gnas paternal allele within a common downstream exon leads to decreased body weight and body fat content and increased whole-body insulin sensitivity as shown by insulin and glucose tolerance tests (27, 28). Here we further extend this observation by showing that in addition to increased muscle glucose uptake, the overall metabolic phenotype of +/p– mice also includes enhanced insulin sensitivity in liver, WAT, and BAT. In fact, the most impressive changes in glucose uptake rate were detected in WAT and BAT with 7- and 3.8-fold increases over control levels, respectively. We also showed that insulin-stimulated IR and Akt phosphorylation was increased in both liver and muscle of +/p– mice, suggesting increased activation of the IR and its downstream effectors accounts for some or all of the increased insulin sensitivity in these tissues. Basal IR and Akt phosphorylation in the absence of insulin were unaffected in +/p– mice, indicating that the insulin pathway is not constitutively activated in these tissues. This is consistent with prior results showing that insulin-stimulated but not basal glucose uptake was greater than normal in skeletal muscles isolated from +/p– mice (28).

It is now well established that obesity-related disturbances in lipid metabolism, such as increased circulating fatty acids and triglycerides as well as excessive deposition of triglycerides and/or other lipid metabolites in nonadipose tissues, can cause insulin resistance (36, 37, 38) and that therapeutic approaches that lower tissue lipid content improve insulin sensitivity (46, 47). We have shown in this and prior studies (27) that +/p– mice have decreased levels of serum triglycerides and show in this study that these mice also have decreased triglyceride levels in liver and muscle. This likely contributes to the increased insulin sensitivity of +/p– mice, particularly the increased sensitivity present in their livers. We also show that +/p– mice have more rapid triglyceride clearance than normal, suggesting that lower serum and tissue triglyceride levels in these mice result from increased rates of lipid metabolism. This is consistent with our prior results that showed that +/p– mice become lean because of increased metabolic rate, rather than decreased food intake (27). The low levels of liver triglyceride in the face of increased lipid clearance suggests that free fatty acids are directed to oxidative pathways, rather than being reesterified into triglyceride. Although mRNA expression of enzymes involved in lipid oxidation or PGC-1 (which stimulates the expression of enzymes involved in fatty acid oxidation) was unaffected in livers of +/p– mice, it is possible that there is increased expression or activity of these proteins. cAMP has been shown to directly affect the activity of CPT1 in hepatocytes (48, 49). AOX mRNA expression was increased in the muscle of these mice, suggesting that increased lipid oxidation in muscle may be a significant contributor to the +/p– metabolic phenotype.

What are the factors that lead to increased lipid oxidation and insulin sensitivity in +/p– mice? Although adipose-derived circulating factors such as leptin and adiponectin can directly stimulate glucose uptake and fatty acid oxidation (44, 50), they probably do not contribute to the metabolic phenotype in +/p– mice because serum leptin levels are low, rather than high (27), and serum adiponectin levels are unchanged in +/p– mice. The fact that serum adiponectin levels are unchanged in +/p– mice probably reflects the opposing effects of adiponectin overexpression in adipose tissue and decreased adipose tissue mass, similar to what has been previously described in the diacylglycerol acyltransferase 1 (Dgat1–/–) mouse (45). Adiponectin expression in adipose tissue is inversely proportional to the level of lipid stores, and therefore the adiponectin overexpression observed in this study is consistent with other animal models. We show that chronic ß-adrenergic stimulates adiponectin expression in WAT, and therefore adiponectin overexpression in the adipose tissue of +/p– mice might be the result of chronic sympathetic overstimulation. Although ß-adrenergic agents have been shown to lower adiponectin acutely (51, 52), chronic ß-adrenergic stimulation was shown to increase adiponectin in one prior study (53). We show that resistin is also overexpressed in our lean +/p– mice. Although originally reported that resistin expression is related to obesity (39), more recent studies show resistin expression to be low in several models of obesity (40, 41, 42, 43). Our results provide another demonstration that resistin expression is not always correlated with obesity. Previous reports suggest that acute ß-adrenergic stimulation suppresses resistin expression (54), whereas chronic ß-adrenergic stimulation induces resistin expression (55).

One possible explanation for the increased lipid clearance in +/p– mice is that these mice are hypermetabolic due to increased activity of the sympathetic nervous system. This is consistent with prior results showing that +/p– mice have increased metabolic rate at both ambient and thermoneutral temperatures, are hyperactive, and have increased urinary norepinephrine excretion. Moreover, both BAT and WAT from these mice have a histological appearance consistent with increased metabolic stimulation, and uncoupling protein 1 expression is increased in BAT from +/p– mice (27), all consistent with increased adrenergic stimulation of these tissues. Increased sympathetic stimulation of metabolically active tissues such as BAT may also indirectly stimulate glucose uptake independent of insulin by stimulating lipid metabolism (56).

What genetic mechanism might underlie the +/p– phenotype? The fact that mice with the same Gnas disruption in the maternal allele do not develop the same metabolic phenotype (although they do also have somewhat increased insulin sensitivity) suggests that partial Gs{alpha} deficiency per se does not explain the phenotype. A more likely explanation is that +/p– mice develop a specific metabolic phenotype due to loss of expression of a paternal-specific Gnas gene product. This explanation is supported by preliminary observations in mice with paternal deletion of Gs{alpha} exon 1 (which are partially deficient in Gs{alpha} but not other Gnas gene products) demonstrating that these mice lack the metabolic phenotype observed in +/p– mice (57). XL{alpha}s is the only known protein that is expressed exclusively from the paternal Gnas allele and is therefore the most likely candidate. This is supported by preliminary observations in XL{alpha}s knockout mice that show that these mice also develop decreased body mass (Plagge, A. and G. Kelsey, personal communication).

Little is known about the importance of XL{alpha}s in vivo. XL{alpha}s is similar to Gs{alpha} except that it has a long amino terminal domain encoded by its specific alternative first exon, and it has been shown to be capable of mediating receptor-stimulated cAMP generation (23). Unlike Gs{alpha}, which is expressed ubiquitously, XL{alpha}s expression is limited primarily to neuroendocrine tissues. XL{alpha}s expression occurs early in the development of the central nervous system and within the distribution of the sympathetic trunk (18, 19, 22). Given that +/p– mice may have increased sympathetic activity, we speculate that XL{alpha}s might normally mediate pathways within the sympathetic nervous system that negatively regulate sympathetic activity. Loss of XL{alpha}s would then be predicted to increase sympathetic activity, which would lead to increased lipid oxidation and glucose uptake in metabolically active tissues. It has been noted that the distribution of XL{alpha}s in the central nervous system somewhat resembles that of pituitary adenylyl cyclase-activating polypeptide (PACAP) receptors (17). Although it is not known whether XL{alpha}s mediates the stimulation of cAMP by PACAP in the central nervous system, it is interesting to note that PACAP knockout mice also develop a severely lean phenotype and abnormal glucose and lipid metabolism (58).

Whereas XL{alpha}s appears to have an important role in mice, it is important to note that patients with paternal null GNAS mutations within common downstream exons, who have genetic defects similar to that in +/p– mice and who are also presumably XL{alpha}s-deficient, do not develop a metabolic syndrome similar to +/p– mice (1). Therefore, there may be species-specific differences in the role that XL{alpha}s plays in metabolic regulation. Further studies in the ever increasing number of genetically altered mouse models will allow us to further explore the role of XL{alpha}s, Gs{alpha}, and other Gnas gene products in energy and glucose metabolism.


   Acknowledgments
 
The authors acknowledge Oksana Gavrilova and Stephanie Pack for their technical support and insightful comments.


   Footnotes
 
This work was supported by the Intramural Research Program of the National Institute of Diabetes and Digestive and Kidney Diseases, Department of Health and Human Services.

M.C. and M.H. contributed equally to the study.

Present address for M.H.: 3 Department of Medicine, 1 Faculty of Medicine, Charles University, Prague 2, Czech Republic.

Present address for N.J.W.: University of Maryland, College Park, Maryland 20742.

Present address for K.R.D.: University of Minnesota, Minneapolis, Minnesota 55455.

Present address for M.L.R.: Merck Research Laboratories, Rahway, New Jersey 07065.

Abbreviations: AOX, Acyl-CoA oxidase; BAT, brown adipose tissue; CPT1, carnitine palmitoyltransferase; IR, insulin receptor; PACAP, pituitary adenylyl cyclase-activating polypeptide; PGC-1, peroxisomal proliferator-activated receptor-{gamma} coactivator-1; WAT, white adipose tissue.

Received January 14, 2004.

Accepted for publication May 21, 2004.


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