Differential Effects of Interleukin-6 and -10 on Skeletal Muscle and Liver Insulin Action In Vivo

RESEARCH DESIGN AND METHODS

Animals and surgery.

Male C57BL/6 mice at ∼12 weeks of age were obtained from The Jackson Laboratory and studied after 2–3 weeks of acclimatization. Mice were housed under controlled temperature (23°C) and lighting (12 h of light, 0700–1900; 12 h of dark, 1900–0700) with free access to water and standard mouse diet. All procedures were approved by the Yale University Animal Care and Use Committee. At least 4 days before hyperinsulinemic-euglycemic clamp experiments, mice were anesthetized with an intraperitoneal injection of ketamine (100 mg/kg body wt) and xylazine (10 mg/kg body wt), and an indwelling catheter was inserted in the right internal jugular vein as previously described (27). On the day of clamp experiment, a three-way connector was attached to the jugular vein catheter to intravenously deliver solutions (e.g., glucose, insulin), and the blood samples were obtained from the tail vessel (27).

Hyperinsulinemic-euglycemic clamps to assess insulin action and signaling in vivo.

After an overnight fast, saline (control, n = 8) or mouse recombinant IL-6 (0.5 μg/h, n = 5; Sigma, St. Louis, MO) was infused for 2 h of the pretreatment period and a blood sample (60 μl) was collected at the end for the measurements of glucose, insulin, and fatty acids. The IL-6 pretreatment period was followed by a 2-h hyperinsulinemic-euglycemic clamp with a primed-continuous infusion of human insulin (Humulin; Eli Lilly, Indianapolis, IN) at a rate of 15 pmol · kg⁻¹ · min⁻¹ to raise plasma insulin within a physiological range and continuous IL-6 infusion. Blood samples (20 μl) were collected at 20-min intervals for the immediate measurement of plasma glucose concentration, and 20% glucose was infused at variable rates to maintain plasma glucose at basal concentrations. Basal and insulin-stimulated whole-body glucose turnover was estimated with a continuous infusion of [3-³H]glucose (Perkin Elmer Life and Analytical Sciences, Boston, MA) during the pretreatment period (0.05 μCi/min) and throughout the clamps (0.1 μCi/min). All infusions were performed using microdialysis pumps (CMA/Microdialysis, North Chelmsford, MA). To estimate insulin-stimulated glucose uptake in individual tissues, 2-deoxy-d-[1-¹⁴C]glucose (2-[¹⁴C]DG) (Perkin Elmer) was administered as a bolus (10 μCi) at 75 min after the start of clamps as previously described (27). Blood samples were taken during and at the end of clamps for measurement of plasma [³H]glucose, ³H₂O, 2-[¹⁴C]DG concentrations, insulin, and/or fatty acids concentrations (27). At the end of the clamps, mice were anesthetized with sodium pentobarbital injection. Within 5 min, three muscles (gastrocnemius, tibialis anterior, and quadriceps) from both hindlimbs, epididymal white adipose tissue, interscapular brown adipose tissue, liver, and heart were taken as previously described (27).

Biochemical assays and calculation.

The glucose concentration during the clamps was analyzed using 10 μl plasma by a glucose oxidase method on a Beckman Glucose Analyzer 2 (Beckman, Fullerton, CA). Plasma insulin concentration was measured by radioimmunoassay using kits from Linco Research (St. Charles, MO). Plasma fatty acid concentration was determined using an acyl-CoA oxidase-based colorimetric kit (Wako Chemicals, Richmond, VA). Plasma concentrations of [3-³H]glucose, 2-[¹⁴C]DG, and ³H₂O were determined after deproteinization of plasma samples as previously described (27). The radioactivity of ³H in tissue glycogen was determined by digesting tissue samples in KOH and precipitating glycogen with ethanol. For the determination of tissue 2-[¹⁴C]DG-6-phosphate content, tissue samples were homogenized, and the supernatants were subjected to an ion-exchange column to separate 2-[¹⁴C]DG-6-phosphate from 2-[¹⁴C]DG (27).

Rates of basal and insulin-stimulated whole-body glucose turnover were determined as the ratio of the [³H]glucose infusion rate (disintegrations per minute [dpm]) to the specific activity of plasma glucose (dpm/μmol) at the end of the basal period and during the final 30 min of the clamps, respectively. Hepatic glucose production (HGP) during the clamps was determined by subtracting the steady-state glucose infusion rate from the whole-body glucose turnover rate. Whole-body glycolysis and glycogen/lipid synthesis were calculated as previously described (27). Glucose uptake in individual tissues was calculated from the plasma 2-[¹⁴C]DG profile, which was fitted with a double exponential or linear curve using MLAB (Civilized Software, Bethesda, MD) and tissue 2-[¹⁴C]DG-6-phosphate content. Skeletal muscle glycolysis and glycogen synthesis were calculated as previously described (27).

Insulin signaling analysis.

Skeletal muscle (gastrocnemius) and liver samples were obtained at the end of the clamps to measure in vivo activities of insulin receptor substrate (IRS)-1–and IRS-2–associated phosphatidylinositol (PI) 3-kinase, respectively. These activities were assessed by immunoprecipitating tissue lysates with polyclonal IRS-1/IRS-2 antibodies (Upstate Biotechnology, Lake Placid, NY) coupled with protein A-sepharose beads (Sigma) and assessing the incorporation of ³²P into PI to yield PI-3-monophosphate. The immune complex was washed, and the activity was determined as previously described (28). Insulin-stimulated activity of protein kinase C (PKC)-λ/ζ was determined by immunoprecipitating tissue lysates with a polyclonal PKC-ζ antibody (Santa Cruz Biotechnology, Santa Cruz, CA) that recognized both PKC-λ and -ζ, coupled with protein A-sepharose beads as previously described (28). Insulin-stimulated Akt activity was determined by immunoprecipitating tissue lysates with a polyclonal Akt antibody (Upstate Biotechnology) that recognized both Akt1 and Akt2, coupled with protein G-sepharose beads (Amersham Pharmacia Biotechnology, Piscataway, NJ) as previously described (28).

Measurements of phospho-STAT3 expression and fatty acyl-CoA in skeletal muscle and liver.

To determine STAT3 tyrosine phosphorylation in IL-6–treated skeletal muscle, tissue samples were obtained after 10-min postintraperitoneal IL-6 injection (1 μg) or 2-h intravenous IL-6 infusion (0.5 μg/h). Muscle lysates (gastrocnemius) were resolved by SDS-PAGE (8% gel) and transferred to nitrocellulose membranes. The nitrocellulose membranes were blocked and incubated with phospho-specific STAT3 (Tyr705) polyclonal antibody (Cell Signaling, Beverly, MA). The membranes were washed and incubated with horseradish peroxidase secondary antibody (Amersham Pharmaceuticals, Arlington Heights, IL), and the bands were visualized using the enhanced chemiluminescence system (Amersham Pharmaceuticals).

To determine the intramuscular and intrahepatic concentrations of fatty acyl-CoA using liquid chromatography tandem mass spectrometry, skeletal muscle (quadriceps) and liver samples were homogenized and prepared using a modified method of Bligh and Dyer (29). A PE Sciex API 3000 tandem mass spectrometer (Applied Biosystems, Foster City, CA) interfaced with the TurboIonSpray ionization source was used for the analysis. The intracellular concentrations of long-chain fatty acyl-CoAs (C16:0, C16:1, C18:0, C18:1, C18:2, and C18:3) were detected in negative electrospray mode, and C17 CoA ester was used as an internal standard. The doubly charged ions of these compounds were transmitted, and singly charged product ions were quantified in multiple reaction mode.

Effects of IL-10 co-treatment on IL-6 and lipid-induced insulin resistance in vivo.

To examine the effects of IL-10 on IL-6–induced insulin resistance, mouse recombinant IL-10 (0.5 μg/h; Sigma) was co-infused with IL-6 (0.5 μg/h) for 2 h of the pretreatment period (n = 7), and a 2-h hyperinsulinemic-euglycemic clamp was conducted in awake mice. Moreover, the effects of IL-10 on lipid-induced insulin resistance were examined by infusing lipid (5 ml · kg⁻¹ · h⁻¹, Liposyn II, triglyceride emulsion, 20% wt/vol; Abbott, Chicago) and heparin (6 units/h) with or without IL-10 (0.2 μg/h) for 5 h (n = 5 for each group), and the clamp experiment was conducted in awake mice. The effects of IL-10 treatment alone on insulin action were determined in an additional group of mice treated with a 2-h infusion of saline (n = 6) or IL-10 (0.5 μg/h, n = 7) followed by clamp experiments.

For insulin-signaling assays (i.e., IRS-1/IRS-2–associated PI 3-kinase, PKC-λ/ζ, and Akt/protein kinase B [PKB] activities), skeletal muscle (gastrocnemius) and liver samples were obtained at the end of clamp experiments. For phospho-STAT3 expression and fatty acyl-CoA levels, skeletal muscle (gastrocnemius and quadriceps, respectively) and/or liver samples were obtained after a 2-h intravenous co-infusion of IL-10 and IL-6 (0.5 μg/h for each) or 5-h co-infusion of IL-10 (0.2 μg/h) and lipid (5 ml · kg⁻¹ · h⁻¹).

Statistical analysis.

Data are expressed as means ± SE. The significance of the difference in mean values among control, IL-6 treated, IL-6/IL-10 co-treated, lipid-infused, and lipid/IL-10 co-treated groups was evaluated using the Duncan’s multiple range test. The significance of the difference in mean values between control and IL-10–treated mice was evaluated using the Student’s t test.

RESULTS

IL-6 decreased skeletal muscle insulin action and signaling in vivo.

IL-6 treatment did not affect basal concentrations of glucose, insulin, and fatty acids compared with the controls (Table 1). Although previous studies have shown the inhibitory effects of IL-6 on pancreatic β-cell function (30,31), our 2-h infusion of IL-6 did not affect circulating insulin levels. Furthermore, previous studies have shown that IL-6 affected the hypothalamus-pituitary-adrenal cortex axis and increased circulating cortocosterone levels in humans (12,32). However, our findings of normal glucose and fatty acids concentrations after acute IL-6 treatment suggested that the dose of IL-6 used in this study did not significantly affect the hypothalamus-pituitary-adrenal cortex axis. During the clamp experiments, the plasma insulin concentration was raised to ∼450 pmol/l, whereas the plasma glucose concentration was maintained at ∼7 mmol/l by a variable infusion of glucose in both groups (Table 2).

The rates of glucose infusion required to maintain euglycemia increased rapidly in the controls and reached a steady state within 90 min. There was a markedly blunted insulin response during the hyperinsulinemic-euglycemic clamp studies in the IL-6–treated mice, as reflected by a significantly lower steady-state glucose infusion rate (Table 2). Insulin-stimulated whole-body glucose turnover was decreased by ∼25% after IL-6 treatment (144 ± 5 vs. 186 ± 12 μmol · kg⁻¹ · min⁻¹ in the controls) (Fig. 1A). The rate of insulin-stimulated glucose uptake in skeletal muscle in vivo was assessed using 2-[¹⁴C]DG injection during clamps in awake mice. The IL-6–induced whole-body insulin resistance was mostly accounted for by 40% decreases in insulin-stimulated skeletal muscle (gastrocnemius) glucose uptake (127 ± 14 vs. 218 ± 23 nmol · g⁻¹ · min⁻¹ in the IL-6–treated mice vs. controls) (Fig. 1B). Insulin-stimulated whole-body glycolysis was significantly reduced, whereas whole-body glycogen/lipid synthesis showed a tendency to decrease after IL-6 treatment (Fig. 2A and B). The pattern of changes in skeletal muscle glucose metabolic flux paralleled that in whole-body glucose metabolism after IL-6 treatment (Fig. 2C and D). These results demonstrated that IL-6 treatment caused significant impairment in skeletal muscle insulin action in vivo.

Recent studies in IRS-1 and IRS-2 gene–disrupted mice have suggested that IRS-1 is important in insulin activation of muscle glucose metabolism (i.e., insulin-stimulated glucose transport and glycogen synthase activity), whereas IRS-2 is more important in mediating insulin activation of hepatic glucose metabolism (i.e., insulin-mediated suppression of basal HGP) (1,33). IL-6 treatment significantly reduced insulin-stimulated IRS-1–associated PI 3-kinase activity (39 ± 11 vs. 148 ± 9 arbitrary units in the IL-6–treated mice vs. controls), a key intracellular mediator of insulin signaling in skeletal muscle (Fig. 1C). These findings demonstrate that IL-6–induced skeletal muscle insulin resistance was most likely secondary to IL-6–induced defects in skeletal muscle insulin signaling. In contrast, insulin-stimulated activities of PKC-λ/ζ and Akt/PKB were not significantly altered among groups (7.0 ± 0.6 vs. 8.5 ± 1.1 arbitrary units for PKC-λ/ζ activity and 537 ± 120 vs. 483 ± 105 arbitrary units for Akt/PKB activity in the IL-6–treated mice vs. controls). Interestingly, IL-6 treatment significantly raised intramuscular total fatty acyl-CoA levels (sum of C16:0, C16:1, C18:0, C18:1, C18:2, and C18:3) in the skeletal muscle (quadriceps) compared with the controls (20 ± 4 vs. 8 ± 4 nmol/g in the IL-6–treated mice vs. controls) (Fig. 1D). Furthermore, IL-6 treatment increased tyrosine phosphorylation of STAT3 in skeletal muscle, indicating that the effect of IL-6 on skeletal muscle was mediated by IL-6–induced activation of the gp130 complex (Fig. 3).

IL-6–induced hepatic insulin resistance in vivo.

The rates of basal HGP were not different among the groups, but insulin’s ability to suppress basal HGP was significantly reduced after IL-6 treatment, reflecting IL-6–induced hepatic insulin resistance (57 ± 13 vs. 3 ± 8 μmol · kg⁻¹ · min⁻¹ for clamp HGP in the IL-6–treated mice vs. controls) (Fig. 4A and B). This is the first in vivo demonstration of insulin resistance induced by IL-6 treatment in mice and supports previous in vitro findings in hepatocytes (17). Insulin-stimulated IRS-2–associated PI 3-kinase activity in liver was decreased by 50% after IL-6 treatment (508 ± 107 vs. 935 ± 79 arbitrary units in the IL-6–treated mice vs. controls) (Fig. 4C). In contrast to the findings in skeletal muscle, intrahepatic fatty acyl-CoA levels were reduced after IL-6 treatment (Fig. 4D).

IL-10 prevented IL-6–induced insulin resistance in skeletal muscle and liver in vivo.

We examined the effects of 2-h treatment of IL-10 and 2-h co-treatment of IL-10 and IL-6 on in vivo insulin action during hyperinsulinemic-euglycemic clamps in awake mice. Plasma glucose concentrations after saline or IL-10 treatment did not differ among groups (7.8 ± 0.5 or 8.4 ± 0.5 mmol/l, respectively). Steady-state rates of glucose infusion as well as hepatic and peripheral insulin actions were not affected by IL-10 treatment (Fig. 5). Furthermore, plasma glucose, insulin, and fatty acids concentrations during basal or clamp periods did not differ after co-treatment of IL-10 and IL-6 compared with the controls or IL-6–treated mice (Tables 1 and 2). Remarkably, IL-10 co-treatment completely restored IL-6–induced blunted rates of glucose infusion during clamps (Table 2). Insulin-stimulated whole-body glucose turnover was normalized after IL-10 co-treatment (191 ± 10 μmol · kg⁻¹ · min⁻¹) compared with the IL-6–treated mice, and this was accounted for by increases in insulin-stimulated skeletal muscle glucose uptake after IL-10 co-treatment (237 ± 20 nmol · g⁻¹ · min⁻¹) (Fig. 1A and B). Similarly, rates of insulin-stimulated whole-body glycogen/lipid synthesis and skeletal muscle glycolysis were increased after IL-10 co-treatment compared with the IL-6–treated mice (Fig. 2B and C).

The protective effects of IL-10 on IL-6–induced skeletal muscle insulin resistance were secondary to their effects on skeletal muscle insulin signaling. After IL-10 co-treatment, insulin-stimulated IRS-1–associated PI 3-kinase activity was significantly increased compared with the IL-6–treated mice (Fig. 1C). These changes in skeletal muscle insulin action and signaling were associated with significantly lower intramuscular fatty acyl-CoA levels in the IL-10 co-treated mice (13 ± 4 nmol/g) compared with the IL-6–treated mice (Fig. 1D). Furthermore, similar to the protective effects of IL-10 on IL-6–induced skeletal muscle insulin resistance, insulin’s ability to suppress basal HGP (1 ± 5 μmol · kg⁻¹ · min⁻¹ for clamp HGP) and insulin-mediated IRS-2–associated PI 3-kinase activity (1,023 ± 239 arbitrary units) were normalized after IL-10 co-treatment (Fig. 4B and C). Intrahepatic fatty acyl-CoA levels remained reduced after co-treatment of IL-10 compared with the controls (Fig. 4D).

IL-10 prevented lipid-induced insulin resistance in skeletal muscle in vivo.

Intralipid was infused for 5 h to raise circulating fatty acid levels (Table 1) and induce insulin resistance in skeletal muscle (23). As expected, lipid infusion significantly decreased insulin-stimulated whole-body and skeletal muscle glucose uptake (142 ± 12 and 83 ± 18 μmol · kg⁻¹ · min⁻¹ for whole-body glucose turnover and skeletal muscle glucose uptake, respectively) (Fig. 1 and 2). These changes were due to lipid-induced decreases in insulin-stimulated IRS-1–associated PI 3-kinase activity in skeletal muscle and were further associated with increased intramuscular fatty acyl-CoA levels (24 ± 11 nmol/g) (Fig. 1C and D). Co-treatment of IL-10 with lipid infusion did not affect circulating fatty acid levels, which remained elevated similar to the lipid-infused mice (Table 1). In contrast, IL-10 co-treatment completely prevented lipid-induced insulin resistance in skeletal muscle, as reflected by normal insulin-stimulated glucose uptake (181 ± 21 nmol · g⁻¹ · min⁻¹) and insulin signaling in skeletal muscle (Fig. 1B and C). Additionally, this protective effect of IL-10 was associated with reduced intramuscular fatty acyl-CoA levels in the IL-10 co-treated with lipid mice (15 ± 2 nmol/g) compared with the lipid-infused mice (Fig. 1D). Neither lipid infusion nor lipid and IL-10 co-treatment affected hepatic insulin action, consistent with our previous results (23).

DISCUSSION

To examine the effects of IL-6 and IL-10 on tissue-specific insulin action, we performed 2-h hyperinsulinemic-euglycemic clamps after a 2-h IL-6, IL-10, or IL-6 and IL-10 co-treatment. Our findings showed that acute IL-6 treatment caused insulin resistance in skeletal muscle and liver that was associated with significant defects in insulin signaling and increases in intramuscular fatty acyl-CoA levels. Additionally, IL-6 treatment increased STAT3 tyrosine phosphorylation in skeletal muscle, suggesting the direct effects of IL-6 on skeletal muscle insulin resistance. In contrast, IL-10 co-treatment completely prevented IL-6–induced insulin resistance in skeletal muscle and liver. The protective effects of IL-10 were associated with normalization of insulin signaling and intramuscular fatty acyl-CoA levels. Moreover, IL-10 co-treatment prevented skeletal muscle insulin resistance mediated by a 5-h lipid infusion, and this effect was also associated with normal insulin signaling and intramuscular fatty acyl-CoA levels. Taken together, our findings demonstrated for the first time that acute IL-6 treatment caused skeletal muscle and hepatic insulin resistance in vivo and that IL-10 co-treatment prevented IL-6–and lipid-induced insulin resistance.

Previous studies have shown that IL-6 altered hepatic glucose metabolism (34,35). Ritchie (34) showed that overnight treatment of cultured rat hepatocytes with IL-6 increased labeled glucose release from prelabeled glycogen pools, whereas Kanemaki et al. (35) showed that IL-6 stimulated hepatic glycogen phosphorylase, a rate-limiting enzyme in glycogenolysis. Further supporting the effects of IL-6 to increase HGP, Senn et al. (17) recently found that IL-6 pretreatment reduced insulin-stimulated IRS-1 tyrosine phosphorylation as well as IRS-1–associated PI 3-kinase activity in isolated hepatocytes. In contrast to the potential effects of IL-6 on hepatic insulin resistance, Stouthard et al. (18) showed that IL-6 treatment increased glucose transport in 3T3-L1 adipocytes. Furthermore, Wallenius et al. (36) generated IL-6–deficient mice that were found to develop obesity and become glucose intolerant. Overall, it is clear that in vitro studies with IL-6 have yielded inconsistent results with regard to its effects on whole-body glucose homeostasis. Additionally, the effects of IL-6 on insulin action in skeletal muscle, quantitatively the most important insulin-responsive organ in the body (37), have not been previously examined.

The present findings demonstrated that IL-6 treatment significantly decreased insulin-stimulated glucose uptake in skeletal muscle in vivo, and the underlying mechanism involved defects in insulin-mediated IRS-1–associated PI 3-kinase activity, which is an important intracellular mediator of muscle insulin signaling (1). In addition to the IL-6–induced peripheral insulin resistance, IL-6 pretreatment also blunted insulin’s ability to suppress basal HGP in vivo and insulin-mediated IRS-2–associated PI 3-kinase activity in the liver. The biological activities of IL-6 are initiated by binding to a high-affinity receptor complex, consisting of two membrane glycoproteins: the 80-kDa ligand binding component and 130-kDa signal-transducing component (gp130). Upon binding of IL-6, dimerization of gp130 is followed by activation of Janus-activated kinase, which promotes the tyrosine phosphorylation of gp130 (15). This event triggers the recruitment of signal-transducing molecules, such as SHP-2 and STAT3, leading to the activation of suppressor of cytokine signaling (SOCS)-3 (38–40). Recent studies have demonstrated the inhibitory effects of SOCS-3 on insulin-signaling molecules in isolated hepatocytes/HepG2 cells (41) and 3T3-L1 adipocytes (42,43), and the underlying mechanism may involve ubiquitin-mediated degradation of IRS-1 and IRS-2 proteins (44). To determine whether IL-6–mediated activation of STAT3 and SOCS-3 is involved in skeletal muscle insulin resistance, we determined the tyrosine phosphorylation of STAT3 in skeletal muscle of mice treated with IL-6 for 10 min and found that IL-6 treatment profoundly increased tyrosine phosphorylation of STAT3 in skeletal muscle compared with the controls. The effects of IL-6–mediated STAT3 activation in skeletal muscle may be further supported by the findings of Zhang et al. (45) in which IL-6 was shown to increase IL-6 receptor expression in skeletal muscle. Overall, one hypothesis by which IL-6 caused skeletal muscle insulin resistance involves IL-6–induced activation of STAT3 and SOCS-3 in skeletal muscle, subsequently promoting SOCS-3–mediated degradation of IRS-1 and IRS-2 and leading to defects in insulin signaling and action.

An alternative hypothesis involves IL-6–induced increases in intramuscular fatty acyl-CoA levels. Previous studies have shown a strong inverse relationship between intramuscular fat content and insulin sensitivity in both animal models and humans (46–48). We have also demonstrated that muscle-specific overexpression of lipoprotein lipase, the rate-controlling enzyme in fatty acid uptake into cells (49), caused muscle-specific insulin resistance due to intramuscular accumulation of fatty acid–derived metabolites (47). The underlying mechanism may involve activation of serine kinase cascade, of which PKC-θ and/or IKK-β may play a role, by fatty acid–derived metabolites (e.g., fatty acyl-CoA) leading to the serine phosphorylation of IRS-1 (50–54). In this regard, Yuan et al. (52) have demonstrated that activation of IKK-β resulted in defects in skeletal muscle insulin signaling. Additionally, serine phosphorylation of IRS-1 has been shown to reduce tyrosine phosphorylation of IRS-1 and interfere with its ability to recruit and activate PI 3-kinase, as occurs upon treatment with TNF-α and okadaic acid (55,56). Thus, IL-6–induced insulin resistance in skeletal muscle may involve increases in intramuscular fatty acyl-CoA levels and their subsequent deleterious effects on insulin signaling and action. The mechanism by which IL-6 increases intramuscular fat content is unknown but may involve increases in fatty acid uptake and/or decreases in fatty acid oxidation. Moreover, since IL-6 pretreatment did not raise intrahepatic fatty acyl-CoA levels, the mechanism by which IL-6 caused hepatic insulin resistance might involve activation of STAT3 signaling pathway and/or other fatty acid metabolites in liver.

Interestingly, we found that co-treatment of the anti-inflammatory cytokine IL-10 completely prevented IL-6–induced defects in both hepatic and skeletal muscle insulin action. The protective effects of IL-10 on IL-6–induced insulin resistance involved normalization of insulin signaling and reduction in intramuscular fatty acyl-CoA levels compared with the IL-6–treated mice. Moreover, we demonstrated that co-treatment of IL-10 also prevented lipid-induced defects in skeletal muscle insulin signaling and action, and this was associated with IL-10–mediated reduction in intramuscular fatty acyl-CoA levels. Since IL-10 prevented both IL-6–induced and lipid-induced insulin resistance in skeletal muscle, the underlying mechanism may involve a common pathway mediated by both IL-6 and lipid infusion. In this regard, we recently demonstrated that skeletal muscle insulin resistance induced by 5-h lipid infusion was due to increases in intramuscular fatty acyl-CoA levels, activation of PKC-θ, and subsequent defects in insulin signaling (57). Thus, we hypothesize that IL-6–induced insulin resistance in skeletal muscle is partly mediated via increases in intramuscular fatty acid–derived metabolites and by potential activation of PKC-θ and/or IKK leading to defects in insulin signaling and action. Furthermore, the protective effects of IL-10 on IL-6–induced and lipid-induced insulin resistance may involve IL-10–mediated decreases in intramuscular fatty acid–derived metabolites and/or direct inhibition of IKK. In contrast, because intrahepatic fatty acyl-CoA levels were reduced after IL-6 or IL-6/IL-10 co-treatment, different mechanisms may be responsible for cytokine-induced alteration in hepatic insulin action.

In conclusion, our findings, for the first time, demonstrate that IL-6 treatment causes insulin resistance in skeletal muscle and liver in vivo and that co-treatment of the anti-inflammatory cytokine IL-10 protects IL-6–induced and lipid-induced insulin resistance. These findings demonstrate the important role of IL-6 in the pathogenesis of insulin resistance and further implicate IL-10 as a potential therapeutic target in the treatment of insulin resistance.