Loss of Resistin Improves Glucose Homeostasis in Leptin Deficiency

Heterozygous resistin-deficient mice (+/−) on the original 129/C57BL/6J mixed background (7) were backcrossed onto a C57BL/6J background (The Jackson Laboratories, Bar Harbor, ME) for seven generations, followed by heterozygous matings to produce wild-type and resistin knockout mice. Resistin (+/−) mice were mated with C57BL/6J-Lep^ob/+ mice (The Jackson Laboratories), and double heterozygotes were then mated to generate ob/ob, resistin knockout ob/ob (Rko-ob/ob), resistin knockout (Rko), and wild-type mice. The mice were weaned at 3 weeks and housed n = 5 per cage in a 12-h light/dark cycle (lights on 0700) at an ambient temperature of 22°C until testing. Food and water were provided ad libitum. A subgroup of Rko and wild-type mice was fed a high-fat (45%) diet (Research Diets, New Brunswick, NJ) from age 6–10 weeks to induce insulin resistance (13). Experimental procedures were in accordance with regulations of the animal care and use committee of the University of Pennsylvania.

Body weight and food intake were measured weekly. Body composition was assessed at 4–22 weeks using dual-emission X-ray absorptiometry (PIXImus DEXA; General Electric, Madison, WI), and energy expenditure was measured using Oxymax EqualFlow indirect calorimetry, as previously described (13,14).

Glucose tolerance testing was performed in 10-week-old mice. After overnight fasting, glucose (2 g/kg) was injected intraperitoneally and tail blood glucose was measured at time 0, 15, 30, 60, 90, and 120 min later using a glucometer (OneTouch Ultra II; Johnson & Johnson). The response to a bolus injection of insulin was assessed. Mice were deprived of food for 5 h (0800–1300), human insulin (0.75 units/kg i.p. Humulin R; Eli Lilly, Indianapolis, IN) was injected intraperitoneally, and tail blood glucose was measured before insulin treatment (time 0 min) and 15, 30, 60, 90, and 120 min later.

Insulin clamp was performed in 10-week-old male wild-type and Rko mice after 4 weeks on a regular chow or high-fat diet and in ob/ob and Rko-ob/ob mice on regular chow diet. The mice were catheterized via the right internal jugular vein under sodium pentobarbital anesthesia and allowed 5 days to recover (15,16). Food was removed at 0800, and resistin elution buffer (vehicle, 100 μl) or resistin (0.25 μg/g) was infused intravenously for 3 h. Then, 2 h later [3-³H]glucose was infused intravenously to assess basal glucose kinetics, followed by hyperinsulinemic-euglycemic clamp for 120 min. A priming intravenous injection of human insulin (40 mU/kg, Humulin; Eli Lilly) was administered, followed by continuous infusion at 10 mU · kg⁻¹ · min⁻¹. Tail blood samples (10 μl) were collected at 20-min intervals for measurement of glucose, and 30% glucose was infused to maintain blood glucose between 120 and 140 mg/dl. Insulin-stimulated whole-body glucose flux was assessed by injecting 10 mCi high-performance liquid chromatography–purified [3-³H]glucose bolus and infusing continuously at 0.1 mCi/min. Once steady state was attained for 60 min, insulin transport in various tissues was determined by injecting 2-deoxy-d-[1-¹⁴C]glucose (10 mCi) 45 min before the end of clamps. Then, 20-μl blood samples were collected at 10-min intervals for measurement of ³[H]glucose, ³H₂O, and 2-deoxy-d-[1-¹⁴C]glucose. Insulin was measured in 10-μl blood samples drawn at the start and end of clamps. The mice were killed at the end of the experiment, and soleus muscles, perigonadal white adipose tissue, and liver were harvested, frozen in liquid nitrogen, and stored at −80°C until processing. The glucose infusion rate (GIR), endogenous glucose production (mostly HGP), rate of glucose disposal (R_d), and tissue glucose uptake were determined as previously described (15,16).

The tissues were homogenized in buffer containing 30 mmol/l Na-HEPES (pH 7.4); 2.5 mmol/l EGTA; 3 mmol/l EDTA; 32% glycerol; 20 mmol/l KCl; 40 mmol/l-glycerophosphate; 40 mmol/l NaF; 4 mmol/l NaPPi; 1 mmol/l Na₃VO₄; 0.1% Nonidet P-40; 2 mmol/l diisopropyl fluorophosphate; 2 mmol/l phenylmethylsulfonyl fluoride; 5 mmol/l aprotinin, leupeptin, and pepstatin A; and 1 mmol/l dithiothreitol. Then, 30–40 μg protein was resolved by SDS-PAGE (4–20% gel) and transferred to nitrocellulose membranes (Bio-Rad). Immunoblotting was performed using antibodies to suppressor of cytokine signaling-3 (SOCS-3), Akt, p-Akt (Ser 473), p-acetyl CoA carboxylase (p-ACC), and ACC (Cell Signaling, Beverly, MA) (7,8). Antibodies to AMP-activated protein kinase (AMPK) and p-AMPK were provided by Morris Birnbaum (University of Pennsylvania) (17). Immunoreactivity was visualized, using enhanced chemiluminescence, and quantified by laser densitometry.

For tissue chemistry, 10-week-old wild-type, Rko, ob/ob, and Rko-ob/ob mice on regular chow diet were killed, using CO₂, after fasting (0800–1300). Blood was obtained through cardiac puncture, and brown adipose tissue (BAT) was rapidly excised, frozen in liquid nitrogen, and stored at −80°C. Insulin, adiponectin, and corticosterone were measured in serum, using specific enzyme-linked immunosorbent assays and radioimmunoassays, and triglycerides, cholesterol, and nonesterified fatty acids were measured using enzyme assays as previously described (13,14). The mRNA expression of uncoupling protein-1 (UCP-1) and peroxisome proliferator–activated receptor-γ coactivator-1α (PGC-1α) was measured using real-time PCR and normalized to 36B4 level (14).

Data are the means ± SE. Changes in various parameters were analyzed by ANOVA and pairwise differences assessed using Neuman-Keul’s test (GraphPad Prism, San Diego, CA). P < 0.05 was considered significant.

C57BL/6J-ob/ob mice showed a significant weight increase at 7 weeks and beyond, when resistin was ablated (Fig. 1A). Body fat also increased in Rko-ob/ob mice (Fig. 1B). Resistin deficiency resulted in a slight increase in weight in older male C57BL/6J wild-type mice (Fig. 1A), although this was not significant. Food intake was nearly doubled in ob/ob and Rko-ob/ob mice compared with wild-type and Rko mice, but there was no difference between ob/ob and Rko-ob/ob mice (Fig. 1C). In contrast, oxygen consumption (Vo₂) was reduced by 50% in ob/ob (1,243 ± 68 ml · kg⁻¹ · h⁻¹) versus wild-type mice (2,894 ± 103) (P < 0.0001), and it decreased further during the light cycle in Rko-ob/ob mice (P < 0.001) (Fig. 1D). Resistin deficiency did not affect locomotor activity as assessed by photobeams (data not shown). However, UCP-1 expression was drastically reduced in BAT of Rko-ob/ob mice (Fig. 1E), and PGC-1α expression was suppressed in BAT of Rko and Rko-ob/ob mice (Fig. 1F), indicating that resistin deficiency exacerbates obesity by decreasing thermogenesis.

The loss of resistin did not affect glucose tolerance in Rko mice on regular chow diet, although the sensitivity to a bolus intraperitoneal insulin injection was improved (Fig. 2A and B). In contrast, glucose tolerance and insulin sensitivity both improved significantly in Rko-ob/ob mice (Fig. 2C and D). Serum glucose and insulin levels were significantly lower in Rko-ob/ob mice, consistent with enhancement of insulin sensitivity (Table 1). Serum triglycerides were also lower in Rko-ob/ob than ob/ob mice, but cholesterol, nonesterified fatty acids, adiponectin, and corticosterone were not affected (Table 1).

Loss of resistin did not affect the basal glucose level (128 ± 4.2 vs. 135 ± 3.9 mg/dl) or insulin (0.74 ± 0.08 vs. 0.87 ± 0.1 ng/ml) in wild-type mice on chow diet. Moreover, the GIR (134 ± 6.9 vs. 130 ± 9.8 mg · kg⁻¹ · min⁻¹), HGP (10.1 ± 2 vs. 9.7 ± 0.3), and R_d (144 ± 5.2 vs. 139 ± 9.6) were not affected by resistin deficiency in the presence of normal leptin alleles, when insulin was increased 10-fold during hyperinsulinemic-euglycemic clamp. In contrast, resistin deficiency in Rko-ob/ob mice doubled the GIR (Fig. 3A). HGP was not affected in Rko-ob/ob mice, but R_d increased significantly (Fig. 3C). The enhancement of R_d in Rko-ob/ob mice was associated with increases in glucose uptake in muscle, white adipose tissue, and BAT (Fig. 3D–F). These changes in Rko-ob/ob mice were reversed by intravenous resistin infusion before the clamp (Fig. 3A and C–F).

Phosphorylation of Akt increased (Fig. 4A) and the SOCS-3 level decreased (Fig. 4B) in Rko-ob/ob muscle, and they were reversed by resistin treatment (Fig. 4A and B). In contrast, loss of resistin in Rko-ob/ob mice did not alter the phosphorylation of AMPK and its downstream target, ACC (Fig. 4C and D). In support of the failure of resistin to affect HGP in Rko-ob/ob mice, neither the loss of resistin nor resistin treatment affected the phosphorylation of Akt, AMPK (Fig. 4E and F), and ACC or the SOCS-3 level in livers of Rko-ob/ob mice (data not shown).

After 4 weeks on a high-fat diet, resistin deficiency increased GIR (Fig. 5A) and suppressed HGP by 25% (Fig. 5B). Furthermore, R_d was enhanced in Rko mice on high-fat diet (Fig. 5C). The rise in R_d was associated with higher glucose uptake in muscle and BAT (Fig. 5D–F). Resistin treatment prevented these changes in glucose kinetics (Fig. 5A–F).

Akt phosphorylation increased in Rko muscle (Fig. 6A), whereas SOCS-3 and phosphorylated AMPK (pAMPK)/AMPK and phosphorylated ACC (pACC)/ACC were reduced (Fig. 6A–D). Resistin treatment reversed pAkt and SOCS-3 levels in Rko muscle (Fig. 6A and B), but not pAMPK/AMPK or pACC/ACC (Fig. 6C and D). Resistin deficiency enhanced Akt phosphorylation in Rko liver (Fig. 6E), consistent with increased HGP (Fig. 5B). Resistin treatment blunted hepatic insulin response in Rko, as evidenced by the reduction in pAkt/Akt level (Fig. 6E). In contrast to muscle, pAMPK/AMPK (Fig. 6F), pACC/ACC, and SOCS-3 levels in liver were not affected by resistin when Rko mice were fed a high-fat diet (data not shown).

Resistin was named for its putative role as a mediator of insulin resistance in rodents (10). In the original studies, circulating resistin level, measured by immunoblotting, was elevated in obese mice (10). Administration of recombinant resistin resulted in glucose intolerance, whereas immunoneutralization of resistin in diet-induced obese mice lowered glucose levels (10). These results have been corroborated by subsequent studies (4–8,18). For example, transgenic overexpression of resistin induces insulin resistance in liver and muscle (5,6). Ablation of the resistin gene in mice lowers fasting glucose by decreasing gluconeogenesis, whereas treatment of resistin knockout mice with recombinant resistin enhances HGP (7). Antisense-mediated “knockdown” of resistin decreased hepatic insulin resistance and gluconeogenesis in mice on a high-fat diet (8). Resistin appears to increase insulin resistance in various tissues at least in part by decreasing phosphorylation of AMPK (7,8,19). Studies have also suggested resistin stimulates hepatic gluconeogenesis and inhibits insulin signal transduction in adipocytes by inducing SOCS-3 (20).

We have addressed the interplay between resistin and leptin, an adipocyte hormone that regulates weight and glucose metabolism. Leptin-deficient ob/ob mice develop early-onset obesity, severe insulin resistance, steatosis, neuroendocrine deficits, and diabetes (21). There are similarities between the nutritional regulation of leptin and resistin in that both adipocyte hormones are reduced by fasting and increased by feeding (12,22). This effect is mediated partly through insulin and glucose (12,22). We have previously shown that serum resistin is increased in ob/ob mice, despite a marked reduction in gene expression per adipocyte (8,12). A functional link between leptin and resistin is suggested by regulation of resistin mRNA expression and protein levels in ob/ob mice by leptin treatment (12,23).

An important result of the current study is that the loss of resistin in ob/ob mice increases body weight and fat by reducing energy expenditure. Although it has been reported that resistin transiently inhibits food intake in rats (24), we did not observe significant changes in feeding between wild-type and Rko and ob/ob and Rko-ob/ob mice. Both ob/ob and Rko-ob/ob mice were hyperphagic. We found UCP-1 mRNA and PGC-1α levels to be reduced in BAT of Rko-ob/ob compared with ob/ob mice. These results uncover a crucial role of resistin in the regulation of thermogenesis, but it remains to be determined whether resistin acts in the brain or peripheral tissues to control metabolic rate. Our observation that resistin deficiency exacerbates obesity is consistent with previous reports (9,22). Treatment with conditioned medium from COS cells expressing resistin inhibited the differentiation of 3T3-L1 cells to adipocytes (22). In contrast, transgenic expression of a dominant inhibitory form of resistin resulted in excess adiposity (9).

Despite being more obese, Rko-ob/ob mice exhibited drastic reductions in glucose and insulin and an enhancement of insulin sensitivity compared with ob/ob mice. The main effect of resistin in the setting of leptin deficiency was to decrease insulin-mediated glucose disposal, particularly in skeletal muscle. This increase in muscle insulin resistance in Rko-ob/ob mice was consistent with the reduction in Akt phosphorylation in muscle. Moreover, SOCS-3 level increased in Rko-ob/ob muscle after resistin treatment, consistent with the hypothesis that SOCS-3 is a mediator of resistin-related insulin resistance (20). Our result on resistin’s effect on muscle insulin sensitivity in Rko-ob/ob mice contrasts with the previous finding that resistin deficiency primarily decreases glucose output in DIO mice (5). Those studies were performed in mixed 129/BL6 mice, whereas the Rko-ob/ob mice were bred to the C57BL/6J strain.

To further elucidate the role of resistin in glucose homeostasis, we performed hyperinsulinemic-euglycemic clamps in Rko mice bred to the C57BL/6J background and fed a high-fat diet. This mouse strain is particularly susceptible to diet-induced obesity and insulin resistance (13). GIR was increased significantly in Rko mice and reversed by resistin treatment. However, in contrast to Rko-ob/ob mice, resistin had dual effects to increase glucose production and decrease peripheral glucose uptake in DIO Rko mice. Accordingly, resistin infusion decreased Akt phosphorylation in muscle and liver of Rko mice. Phosphorylation of AMPK and ACC and the level of the negative insulin regulator SOCS-3 were all reduced in Rko muscle; however, only SOCS-3 was partially reversed by resistin treatment. In contrast, these putative mediators of resistin action were not altered in Rko liver. The reasons for these differences are unclear, but they may indicate different resistin thresholds in insulin target tissues.

We have established that high resistin levels confer insulin resistance in models of obesity. Insulin sensitivity is improved in Rko mice on high-fat diet and in Rko-ob/ob mice, and it is reversed by acute resistin treatment. Loss of resistin also impairs thermogenesis and exacerbates obesity in Rko-ob/ob mice, suggesting a role in energy homeostasis. These studies reveal that the actions of resistin depend on the presence or absence of leptin, as well as interaction with diet. In humans, where resistin appears to be primarily derived from macrophages rather than adipocytes, some studies have failed to demonstrate a significant association between resistin and adiposity and glucose (25,26), whereas others have shown increased resistin levels in obesity, inflammation, and type 2 diabetes (27–29). Although the connection between resistin and insulin sensitivity in humans has yet to be proven, our findings raise the possibility of cross talk between resistin and leptin in the regulation of energy and glucose homeostasis.

This study was supported by National Institutes of Health Grants PO1DK49210 (to M.A.L. and R.S.A.) and RO1DK62348 (to R.S.A.). Clamp and various metabolic studies were performed by the Penn DERC (Diabetes and Endocrinology Research Center) Mouse Phenotyping Core (P30DK19525).

Our thanks to Ronadip Banerjee and Shamina Rangwala for providing breeder resistin knockout mice.