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Am J Physiol Endocrinol Metab 293: E1772-E1781, 2007. First published October 16, 2007; doi:10.1152/ajpendo.00158.2007
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Increased lipid accumulation and insulin resistance in transgenic mice expressing DGAT2 in glycolytic (type II) muscle

Malin C. Levin,1,* Mara Monetti,1,* Matthew J. Watt,2 Mini P. Sajan,3 Robert D. Stevens,4 James R. Bain,4 Christopher B. Newgard,4 Robert V. Farese, Sr.,3 and Robert V. Farese, Jr.1,5,6

1Gladstone Institute of Cardiovascular Disease, San Francisco, California; 2St. Vincent's Institute of Medical Research, Fitzroy, Australia; 3James A. Haley Veterans Hospital, Department of Medicine, University of South Florida, Tampa, Florida; 4Stedman Nutrition and Metabolism Center, Duke University, Durham, North Carolina; 5Departments of Medicine and of Biochemistry and Biophysics; and 6Diabetes Center, University of California, San Francisco, California

Submitted 9 March 2007 ; accepted in final form 27 September 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Insulin resistance and type 2 diabetes are frequently accompanied by lipid accumulation in skeletal muscle. However, it is unknown whether primary lipid deposition in skeletal muscle is sufficient to cause insulin resistance or whether the type of muscle fiber, oxidative or glycolytic fiber, is an important determinant of lipid-mediated insulin resistance. Here we utilized transgenic mice to test the hypothesis that lipid accumulation specifically in glycolytic muscle promotes insulin resistance. Overexpression of DGAT2, which encodes an acyl-CoA:diacylglycerol acyltransferase that catalyzes triacylglycerol (TG) synthesis, in glycolytic muscle of mice increased the content of TG, ceramides, and unsaturated long-chain fatty acyl-CoAs in young adult mice. This lipid accumulation was accompanied by impaired insulin signaling and insulin-mediated glucose uptake in glycolytic muscle and impaired whole body glucose and insulin tolerance. We conclude that DGAT2-mediated lipid deposition specifically in glycolytic muscle promotes insulin resistance in this tissue and may contribute to the development of diabetes.

acyl-CoA:diacylglycerol acyltransferase 2; skeletal muscle; glycolytic fibers; triacylglycerols

INSULIN RESISTANCE AND TYPE 2 DIABETES are frequently associated with lipid accumulation in skeletal muscle (30, 48, 56). In humans, increased intramyocellular lipid content correlates with insulin resistance (45). In both rodents and humans, intravenous infusion of lipid or chronic high-fat feeding results in the accumulation of lipids, including triacylglycerol (TG), and insulin resistance in skeletal muscle (6, 12, 23, 33). Mice with genetic modifications that increase muscle lipids, such as muscle-specific overexpression of lipoprotein lipase and CD36 (26, 31), also exhibit insulin resistance. On the other hand, endurance exercise training increases TG deposition in skeletal muscle and is associated with increased insulin sensitivity (22). Thus, the relationship between lipid accumulation in muscle and insulin resistance remains incompletely understood.

One factor that has not been extensively studied with respect to intramuscular lipid deposition is fiber type. Skeletal muscle is composed mainly of two types of fibers, oxidative (type I, slow-twitch, red) and glycolytic (type II, fast-twitch, white), whose proportions can vary greatly. Marathon runners, for example, have ~80% type I fibers in leg muscles (21), whereas sprinters have ~75% type II fibers (14). It is unknown whether the type of muscle fiber is an important consideration in lipid-associated insulin resistance. Oxidative muscle normally stores more TG and is more insulin sensitive than glycolytic muscle (43, 44). Moreover, endurance training increases both TG content and insulin sensitivity in oxidative muscle (22). Thus, increased lipid content in oxidative muscle may not be detrimental. On the other hand, glycolytic muscle stores less TG and is less insulin sensitive than oxidative muscle (43, 44). The relationship of lipid accumulation in glycolytic muscle and insulin resistance has not been directly tested.

In this study, we tested the hypothesis that lipid accumulation in glycolytic muscle would promote insulin resistance. We generated transgenic mice that overexpress human acyl-CoA:diacylglycerol acyltransferase 2 (DGAT2) specifically in glycolytic muscle [muscle creatine kinase (MCK)-DGAT2 mice]. DGAT enzymes catalyze the final step of TG synthesis, a reaction that covalently links diacylglycerol (DG) and fatty acyl-CoA substrates (10). DGAT2 is one of two DGAT enzymes (11) and is more potent (51) and specific (57) for TG biosynthesis. We reasoned that increasing DGAT2 expression in glycolytic muscle would promote the accumulation of TG and possibly other lipids in this tissue. Here we report the phenotype of MCK-DGAT2 mice, focusing on the effects of excessive lipid accumulation on insulin signaling and action in glycolytic muscle.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Mice. The MCK-DGAT2 transgene was made with a 4-kb MCK promoter from the pCCLMCK-II vector (a gift from R. Eckel, University of Colorado, Denver, CO) (34), a 1.2-kb human DGAT2 cDNA, and a 2.2-kb human growth hormone minigene (a gift from J. Marth, University of California, San Diego, CA). The transgene was introduced into C57BL/6NHsd fertilized eggs (Harlan Teklad, Madison, WI) by microinjection. Founder animals were bred with C57BL/6NHsd mice, and an inbred transgenic mouse line was established. Mice were housed in a pathogen-free barrier facility (12:12-h light-dark cycle) and fed rodent chow (Ralston Purina, St. Louis, MO). The mice were studied at 3 mo of age unless otherwise stated. For high-fat diet experiments, mice were fed a Western-type diet (21% fat and 0.2% cholesterol by weight; Harlan Teklad) for 24 wk starting at 12 wk of age. Skeletal muscle samples of mixed (total gastrocnemius), oxidative (red head of the gastrocnemius and soleus), or glycolytic (white heads of the gastrocnemius) muscle were used for analyses.

Genotyping. Genotyping was performed by PCR of tail genomic DNA with the primers forward 5'-AGCTGGTGAAGACACACAACC-3' and reverse 5'-TGATGATAGCATTGCCACTCC-3', which amplify an ~800-bp DGAT2 fragment in wild-type (WT) mice and a ~300-bp fragment in transgenic mice.

Real-time quantitative PCR. Mouse tissues were pulverized in liquid nitrogen, and total RNA was isolated with RNA Stat-60 (Tel-test, Friendswood, TX) according to the manufacturer's protocol. RNA (2.5 µg) was reverse transcribed with Superscript III reverse transcriptase and oligo(dT) primers (Invitrogen, Carlsbad, CA) to generate cDNA. Real-time quantitative PCR was performed with the ABI PerkinElmer Prism 7700 (Applied Biosystems, Foster City, CA) and SYBR Green detection of amplified products. Relative mRNA abundance was normalized to the internal standard cyclophilin (forward primer 5'-TGGAAGAGCACCAAGA CAGACA-3' and reverse primer 5'-TGCCGGAGTCGACAATGAT-3'). The primers used for DGAT2 (forward 5'-AGTGGCAAT GCTATCATCATCGT-3' and reverse 5'-TCTTCTGGACCCATCGGCCCCAGGA-3') are located in regions where the human and mouse DGAT2 cDNA sequences are 100% identical and therefore amplify both gene products. Real-time PCR was also used to quantify troponin I (slow; forward primer 5'-TGCCGGAAGTTGAGAGGAAATCCAAGAT-3' and reverse primer 5'-CCAGCACCTTCAGCTTCAGGTCCTTGAT-3'), myoglobin (forward primer 5'-GCACAAG ATCCCGGTCAAGTACCTGGAG-3' and reverse primer 5'-CTGACGAAGGCCACTTTG CACCTCTG-3'), MCK (forward primer 5'-GATGTCATCCAGACTGGGGTGGACAACC-3' and reverse primer 5'-TGAACTCGCCCGTCAGGCTGTTGAG-3'), glucose transporter 1 (GLUT1; forward primer 5'-CATTTTAGGATTCGCCCATTC-3' and reverse primer 5'-CAGT GCTTCCAACTGGTCTCA-3'), and GLUT4 (forward primer 5'-CTGTCCTGAGAGCCCCA GAT-3' and reverse primer 5'-CAGGCGCTTTAGACTCTTTCG-3').

DGAT activity assays. Mouse tissue was pulverized in liquid nitrogen and homogenized in buffer [buffer A, 50 mM Tris·HCl (pH 7.4), and 250 mM sucrose] containing proteinase inhibitors. To prepare microsomes, the homogenates were centrifuged three times at 4°C (600 g for 5 min, 10,000 g for 10 min, and 100,000 g for 1 h); after each centrifugation, the pellets were resuspended in Tris-sucrose buffer. DGAT assays were performed with microsome proteins (200 µg for red and white gastrocnemius and 50 µg for liver and heart) in an assay mix containing buffer A, 5 mM MgCl2, 1.25 mg/ml BSA, 200 µM 1,2-dioleoyl-sn-glycerol (Sigma-Aldrich, St. Louis, MO) in acetone, and 25 µM [14C]oleoyl-CoA (53.0 mCi/mmol). Lipids were extracted with chloroform-methanol (2:1, vol/vol) and separated by silica gel G-60 TLC plates with hexane-ethyl ether-acetic acid (80:20:1) solvent. The TG bands were scraped, and radioactivity was measured by scintillation counting.

Lipid analyses of skeletal muscle. Mouse tissue was pulverized in liquid nitrogen and homogenized in buffer A containing proteinase inhibitors. Lipids were extracted with chloroform-methanol (2:1) and separated on silica gel G-60 TLC plates in hexane-ethyl ether-acetic acid (80:20:1). The TG bands were scraped, and TG was analyzed as described (49) with triolein as a standard.

DG and ceramides were analyzed as described (46). Skeletal muscle was freeze-dried and dissected free of visible connective tissue, and lipids were extracted with chloroform-methanol-PBS + 0.2% SDS (1:2:0.8). DG kinase (Sigma-Aldrich) and [{gamma}-32P]ATP (15 mCi/mmol cold ATP) were added to lysates preincubated with cardiolipin/octylglucoside. After 2 h, the reaction was stopped by adding chloroform-methanol (2:1), and extracted lipids were spotted onto TLC plates and developed in chloroform-acetate-methanol-acetic acid-water (100:40:30:20:10). 32P-labeled phosphatidic acid (corresponding to DG) and ceramide-1-phosphate (corresponding to ceramide) were identified and scraped from the TLC plate for scintillation counting (Tri-Carb 2500TR; Packard, Canberra, Australia).

Skeletal muscle acyl-CoA esters were analyzed with a method based on that of Magnes et al. (41) that utilizes an extraction procedure described by Deutsch et al. (16). CoAs were purified by solid-phase extraction using Oasis HLB cartridges (1 cm3, 30 mg; Waters). The esters were eluted with 0.55 ml of acetonitrile-water (35:65 vol/vol) containing 15 mM ammonium hydroxide. The acyl-CoAs were analyzed by flow injection analysis using positive electrospray ionization on a Quattro micro, triple quadrupole mass spectrometer (Waters), with acetonitrile-water (60:40, vol/vol) containing 30 mM ammonium hydroxide as the mobile phase. Spectra were acquired in the multichannel acquisition mode by monitoring the neutral loss of 507 amu (phosphoadenosine diphosphate) and scanning from mass-to-charge ratio 890–1,060. Heptadecanoyl CoA was used as an internal standard, and endogenous CoAs were quantified with calibrators prepared by spiking liver homegenates with authentic CoAs (Sigma) that had saturated acyl chain lengths C8–C18 and unsaturated species of C16:1, C18:2, C18:1, and C20:4. Corrections for the heavy isotope effects, mainly 13C, to the adjacent m + 2 spectral peaks in a particular chain length cluster were made empirically by referring to the observed spectra for the analytical standards.

Insulin-signaling analyses. After an overnight fast, mice received PBS or insulin (1 U/kg body wt) intraperitoneally. After 15 min, muscles were excised and snap-frozen in liquid nitrogen. For analysis of insulin-stimulated PKB/Akt phosphorylation, the tissue was pulverized in liquid nitrogen, homogenized in buffer A containing proteinase inhibitors, and centrifuged at 800 g for 5 min. The supernatant protein (50 µg) was analyzed by SDS-PAGE and immunoblotting with antibodies that detect total Akt or phospho-Akt (Ser473; Cell Signaling Technology, Danvers, MA).

IRS-1-associated phosphoinositide 3-kinase (PI3K), PKC{lambda}/{zeta}, PKB, and PKC{theta} activity assays were performed as described (13, 29, 50). Briefly, PBS or insulin (1 U/kg body wt) was injected intraperitoneally, and muscle tissues were removed after 15 min. For IRS-1/PI3K, tissue lysates were immunoprecipitated with polyclonal antibodies that recognize IRS-1 (Upstate Biotechnology, Lake Placid, NY), and precipitates were incubated with [{gamma}-32P]ATP. Aliquots of the mixture were separated by thin-layer chromatography, and 32P radioactivity of phosphatidylinositol 3-phosphate was measured. For PKC{lambda}/{zeta} studies, tissue lysates were immunoprecipitated with polyclonal antibodies that recognize specific isoforms of PKC (Santa Cruz Biotechnology, Santa Cruz, CA). Precipitates were incubated for 8 min with [{gamma}-32P]ATP and the serine analog of PKC{varepsilon} pseudosubstrate (Biosource International, Camarillo, CA). Aliquots of the mixtures were spotted on p81 filter paper, and 32P radioactivity was measured. PKB activity was measured with a kit (Upstate Biotechnology).

In vivo glucose uptake. In vivo glucose uptake was measured as described (58), with some modifications. Briefly, after an overnight fast, mice received an intraperitoneal injection of saline containing 50 µCi/kg 2-deoxy-D-[3H]glucose (specific activity, 9.0 Ci/mmol; Amersham Biosciences, GE Health Care Life Sciences, Piscataway, NJ) with or without insulin (1 U/kg body wt). Injections also contained 2.5 µCi/kg L-[14C]glucose (specific activity, 55 mCi/mmol; Amersham Biosciences), which served as a control for glucose that diffused into the intercellular (but not intracellular) space. Injection of 2-deoxyglucose did not affect serum glucose or insulin levels. After 15 min, blood was collected, serum glucose was measured, and the gastrocnemius, soleus, heart, and white adipose tissue (WAT) were excised and snap-frozen in liquid nitrogen. The tissues were homogenized in ice-cold buffer A, and radioactivity was counted in serum and homogenates. The glucose-specific activity was calculated by dividing serum radioactivity by serum glucose (cpm/nmol). Cellular uptake of deoxyglucose was calculated by subtracting the nonspecific uptake and dividing the corrected value by tissue weight and glucose-specific activity.

Plasma analyses. Mice were fasted overnight (16 h) for plasma lipid and 7 h for plasma insulin determinations. Blood was obtained from tail veins of unanesthetized mice. Plasma free fatty acid levels were measured with a kit (Wako Pure Chemical Industries, Osaka, Japan), as were TG levels (Roche Diagnostics, Indianapolis, IN). ELISA kits were used to measure plasma levels of insulin (Linco Research, St. Charles, MO) and leptin (R&D Systems, Minneapolis, MN).

Glucose and insulin tolerance tests. For glucose tolerance tests, mice were fasted overnight, and glucose (1 g/kg body wt) was administered intraperitoneally. For insulin tolerance tests, mice were fasted for 4 h, and bovine insulin (1 U/kg body wt; Sigma-Aldrich) was administered intraperitoneally. Blood glucose concentrations were measured with a One-Touch Ultra Glucometer (Lifescan, Milpitas, CA).

Statistical analyses. Data are shown as means ± SE. Measurements were compared with the two-tailed t-test or Mann-Whitney rank sum test. Glucose and insulin tolerance tests were compared with repeated-measures ANOVA and Bonferroni post hoc analysis. Areas under the curves were calculated by using the trapezoid rule.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Generation of mice overexpressing DGAT2 in glycolytic skeletal muscle. Using a transgenic construct containing the MCK promoter (Fig. 1A), we generated transgenic mice that overexpress human DGAT2 cDNA in skeletal muscle. Total (murine and human) DGAT2 mRNA levels were threefold higher in mixed fiber (gastrocnemius and soleus) skeletal muscle of the transgenic mice but were normal in heart and liver (Fig. 1B). Because the MCK promoter induces gene expression at higher levels in glycolytic than in oxidative muscle (28, 37), we examined DGAT2 expression in muscles containing different fiber types. DGAT2 mRNA levels were increased sixfold in glycolytic muscle (white heads of gastrocnemius) of MCK-DGAT2 mice but were not increased significantly in oxidative muscle (red head of gastrocnemius and soleus) (Fig. 1C). In WT mice, endogenous DGAT2 mRNA levels were sevenfold higher in oxidative than in glycolytic muscle. DGAT1 mRNA levels were similar in skeletal muscle of MCK-DGAT2 and WT mice (not shown). The expression of markers for oxidative [troponin I (slow) and myoglobin] and glycolytic (MCK) fibers verified the authenticity of the tissue samples and showed that DGAT2 overexpression did not result in a fiber type switch (Fig. 1D). DGAT activity was increased 60% specifically in glycolytic muscle of MCK-DGAT2 mice (Fig. 1E). The smaller proportional increase in DGAT activity than in DGAT2 mRNA levels in glycolytic muscle is consistent with observations that the in vitro assay may not optimally detect DGAT2 activity (51).


Figure 1
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  		Fig. 1. Generation of muscle creatine kinase (MCK)-acyl-CoA:diacylglycerol acyltransferase 2 (DGAT2) transgenic mice. A: MCK-DGAT2 transgene. The construct contains the MCK promoter, human DGAT2 cDNA, and a bovine growth hormone polyA signal. B: DGAT2 mRNA levels in tissues. mRNA levels were measured by real-time PCR in tissues of 3-mo-old male mice (n = 4/genotype). Sk, skeletal. *P < 0.05 vs. wild type (WT). C: increased DGAT2 mRNA levels in glycolytic muscle in MCK-DGAT2 mice. mRNA levels were measured in muscle of 3-mo-old male mice (n = 7/genotype). **P < 0.001 vs. WT; {dagger}P < 0.001 vs. WT oxidative muscle. D: analysis of muscle type-specific gene expression in WT and MCK-DGAT2 mice. Expression of troponin I (slow), myoglobin, and MCK in oxidative and glycolytic muscle in 3-mo-old male mice (n = 6 WT and n = 5 MCK-DGAT2). *P < 0.001 vs. WT glycolytic muscle; {dagger}P < 0.001 vs. WT oxidative muscle. E: increased DGAT activity in glycolytic muscle of MCK-DGAT2 mice (n = 10/genotype for glycolytic muscle, n = 6/genotype for oxidative muscle, and n = 5/genotype for heart). *P < 0.05 vs. WT glycolytic muscle. DGAT activity assays were performed using microsomes and 5 mM MgCl2.

 
In 3-mo-old mice, DGAT2 overexpression increased lipid deposition in glycolytic muscle. TG levels increased 1.8-fold specifically in this fiber type (Fig. 2A). In contrast, TG levels were similar in oxidative muscle of MCK-DGAT2 and WT mice, as were TG levels in WAT and liver. We also measured the skeletal muscle content of several lipids, DG, ceramides, and long-chain fatty acyl-CoAs, that have been implicated in impaired insulin signaling (27, 55, 59). In glycolytic muscle, DG levels were 42% lower in MCK-DGAT2 mice (Fig. 2B). In contrast, the contents of ceramides (63% higher) (Fig. 2C) and of several unsaturated fatty acyl CoA species (Fig. 2D) were increased in glycolytic muscle in MCK-DGAT2 mice. The contents of these lipids were similar in oxidative muscle of WT and MCK-DGAT2 mice. The differences in DGAT2 mRNA expression and TG accumulation in glycolytic muscle were maximal in young adult mice (~3 mo old). By 6 mo of age, TG content in glycolytic muscle was further increased to similar levels in mice of both genotypes, and phenotypic differences between genotypes were no longer apparent; a similar lack of effect of genotype was seen in mice fed a high-fat diet for 24 wk (Supplemental Fig. S1; Supplemental Material for this article is available at the AJP-Endocrinology and Metabolism web site). Therefore, all subsequent phenotypic studies were performed in 3-mo-old mice.


Figure 2
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  		Fig. 2. Lipid content in muscles of 3-mo-old male WT and MCK-DGAT2 mice on a chow diet. A: triacylglycerol levels in tissues of MCK-DGAT2 mice [n = 7–9/genotype for oxidative and glycolytic muscle, n = 5/genotype for white adipose tissue (WAT) and liver]. B: diacylglycerol levels in muscle in MCK-DGAT2 mice (n = 5 WT and n = 9 MCK-DGAT2). C: ceramide levels in WT and MCK-DGAT2 mice (n = 9/genotype). D: long-chain fatty acyl-CoA (LCFA-CoA) levels in WT and MCK-DGAT2 mice (n = 5 WT and n = 7 MCK-DGAT2). *P < 0.05 and **P < 0.01 vs. WT glycolytic muscle. Lipid content of tissues was measured as described in METHODS.

 
Impaired insulin signaling in glycolytic muscle of MCK-DGAT2 mice. Next, we examined insulin signaling in homogenates of muscle of WT and MCK-DGAT2 mice. In mixed muscle, baseline and insulin-stimulated IRS-associated PI3K activity levels were similar in the two groups (Fig. 3A). However, in glycolytic muscle, insulin-stimulated PI3K activity was clearly decreased in the MCK-DGAT2 mice. Insulin-stimulated PKB activity levels in oxidative muscle were similar in WT and MCK-DGAT2 mice but were significantly decreased in glycolytic muscle in MCK-DGAT2 mice (Fig. 3B). We also examined insulin-stimulated PKC{lambda}/{zeta} activity, a sensitive marker of insulin signaling (29), in oxidative and glycolytic muscle (Fig. 3C). PKC{lambda}/{zeta} activity increased about twofold in both types of muscle in WT mice and 1.4-fold in oxidative muscle of MCK-DGAT2 mice. However, insulin failed to stimulate PKC{lambda}/{zeta} activity in glycolytic muscle of MCK-DGAT2 mice. The activity of PKC{theta}, which has been implicated in lipid-mediated insulin resistance (23), was not different in skeletal muscle of WT and MCK-DGAT2 mice (Fig. 4).


Figure 3
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  		Fig. 3. Impaired insulin signaling in glycolytic muscle of MCK-DGAT2 mice. A: insulin-stimulated insulin substrate-1 (IRS-1)-dependent phosphoinositide 3-kinase (PI3K) activity in mixed and glycolytic skeletal muscle. (n = 3–5 mice/group). **P < 0.01 vs. basal; *P < 0.05 vs. basal. B: insulin-stimulated PKB activity in oxidative and glycolytic muscle. (n = 3–5 mice/group). **P < 0.01 vs. basal; *P < 0.05 vs. basal; {dagger}P < 0.05 vs. WT insulin-stimulated muscle. C: insulin-stimulated PKC{lambda}/{zeta} activity in oxidative and glycolytic muscle. (n = 8–11). *P < 0.05 vs. basal; **P < 0.01 vs. basal. Saline or insulin was administered intraperitoneally to 3-mo-old male mice after an overnight fast. After 15 min, muscles were excised and snap-frozen in liquid nitrogen. PI3K, PKB, and PKC{lambda}/{zeta} activities were measured as described in METHODS.

 

Figure 4
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  		Fig. 4. No difference in basal PKC{theta} activity in oxidative and glycolytic muscle of WT and MCK-DGAT2 mice. Basal PKC{theta} activity in oxidative and glycolytic muscle of WT and MCK-DGAT2 mice. (n = 4/group). After an overnight fast, muscles were excised from 3-mo-old male mice and snap-frozen in liquid. PKC{theta} activity was measured as described in METHODS.

 
Impaired glucose uptake in glycolytic muscle of MCK-DGAT2 mice. Because glycolytic muscle of MCK-DGAT2 mice exhibited impaired insulin signaling, we examined glucose uptake in vivo. In WT muscle, insulin stimulated the uptake of 2-deoxyglucose in both oxidative and glycolytic skeletal muscle, heart muscle, and WAT (Fig. 5); uptake was >10-fold higher in oxidative than in glycolytic muscle at baseline and increased more than twofold in both tissues in response to insulin. In MCK-DGAT2 mice, insulin stimulated 2-deoxyglucose uptake in oxidative muscle, heart, and WAT but not in glycolytic muscle. In quadriceps muscle (a mixed muscle that is rich in glycolytic fibers), insulin signaling and insulin-stimulated glucose uptake were also impaired in MCK-DGAT2 mice (Supplemental Fig. S2). Interestingly, basal 2-deoxyglucose uptake trended higher in both glycolytic and oxidative muscle of MCK-DGAT2 mice (Fig. 5), suggesting an increase in glucose utilization in muscle tissue when DGAT2 is overexpressed.


Figure 5
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  		Fig. 5. Impaired insulin-stimulated glucose uptake in vivo in glycolytic muscle of MCK-DGAT2 mice. Insulin-stimulated uptake of 2-deoxyglucose (2-DOG) in tissues of MCK-DGAT2 (n = 8–9/genotype for oxidative muscle, n = 5–6/genotype for glycolytic muscle, n = 4–6/genotype for heart, and n = 6–7/genotype for WAT). *P < 0 vs. basal. After an overnight fast, 3-mo-old male mice received an intraperitoneal injection of saline containing 2-deoxy-D-[3H]glucose with or without insulin (1 U/kg body wt). After 15 min, blood was collected, and the gastrocnemius, soleus, heart, and WAT were excised and snap-frozen in liquid nitrogen. The tissues were analyzed, and glucose uptake was calculated as described in METHODS. ND, not detectable.

 
Impaired glucose and insulin tolerance in MCK-DGAT2 mice. The fasting plasma levels of glucose, insulin, and leptin were normal in MCK-DGAT2 mice (data not shown). To determine whether the impairment in insulin signaling in glycolytic muscle altered systemic glucose metabolism, we performed glucose and insulin tolerance tests. In MCK-DGAT2 mice, glucose tolerance was significantly impaired at 15, 30, and 60 min after a glucose challenge (Fig. 6A), and insulin tolerance was impaired at 60 min (Fig. 6B). For each test, the area under the curve (arbitrary units) was significantly greater for MCK-DGAT2 mice than for WT mice (17.3 ± 0.78 vs. 20.0 ± 0.78; n = 10–12, P < 0.05) and insulin tolerance tests (9.60 ± 0.41 vs. 12.6 ± 1.2; n = 6–10, P < 0.05).


Figure 6
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  		Fig. 6. Impaired glucose and insulin tolerance in MCK-DGAT2 mice. A: glucose tolerance in 3-mo-old male mice on a chow diet (n = 10 WT and n = 12 MCK-DGAT2). Glucose was administered intraperitoneally after an overnight fast, and blood glucose was measured at the indicated times. *P < 0.05 vs. WT. B: insulin tolerance in 3-mo-old male mice on a chow diet (n = 10 WT and n = 6 MCK-DGAT2). Insulin was administered intraperitoneally after a 4-h fast, and blood glucose was measured at the indicated times. *P < 0.05 vs. WT.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
This study shows that increased lipid deposition mediated by DGAT2 expression specifically in glycolytic muscle of mice promotes insulin resistance in this tissue. Young adult MCK-DGAT2 mice had increased content of TG, ceramides, and unsaturated fatty acyl-CoAs in glycolytic muscle, which was associated with impaired insulin signaling and insulin-stimulated glucose uptake in this tissue. Furthermore, the insulin resistance in this tissue resulted in impairments in whole-body glucose and insulin tolerance. These findings suggest that excessive lipid deposition specifically in glycolytic muscle may be an important contributing factor to insulin resistance.

Skeletal muscle consists primarily of oxidative and glycolytic fibers, whose proportions differ in various muscle groups (2) and between individuals (36, 60). Oxidative fibers have a higher capacity to store and utilize lipids and higher mitochondrial content and activities of respiratory enzymes (3, 19, 24). Indeed, the increase in skeletal muscle TG content with endurance exercise occurs primarily in oxidative muscle (22). In contrast, glycolytic fibers rely mainly on carbohydrates as an energy source and do not store as much lipid (56a), probably reflecting their lower capacity to oxidize lipids. Interestingly, few studies have examined the contribution of lipid deposition in specific fiber types to insulin resistance. However, one study suggested that glycolytic muscle may be a major site for insulin resistance because it has high activity levels of enzymes that interfere with insulin signaling (18) and because insulin resistance increases the proportion of glycolytic fibers in skeletal muscle (36).

DGAT2 overexpression in glycolytic muscle caused lipid accumulation and clearly promoted insulin resistance in this tissue. The activities of IRS-associated PI3K and two downstream targets, PKB and PKC{lambda}/{zeta}, were all suppressed in response to insulin in 3-mo-old MCK-DGAT2 mice. Notably, insulin had almost no ability to stimulate the activity of atypical PKC{lambda}/{zeta} in glycolytic muscle in MCK-DGAT2 mice. PKC{lambda}/{zeta} is a sensitive and specific marker for insulin action in skeletal muscle that correlates highly with insulin's effects on glucose transport (4, 7). Insulin-stimulated PKB activity was also suppressed in glycolytic muscle, albeit to a lesser extent. One possible explanation for this minor discrepancy is that PKB may be relatively more sensitive to insulin than PKC{lambda}/{zeta} (54) and that this signaling is therefore partially maintained. Alternatively, lipids may provoke insulin resistance in part by a mechanism that is specific to PKC{lambda}/{zeta}. For example, defective PKC{lambda}/{zeta} activity is associated with an impaired responsiveness to phosphatidylinositol 3,4,5-phosphate (PIP3) (5), possibly due to desensitization or inhibition of the PIP3 binding domain (20). Other studies of insulin resistance in skeletal muscle in obese humans and in rats fed a high-fat diet have shown decreases in insulin-stimulated PKC{lambda}/{zeta} activity without or with smaller decreases in PKB activity (29, 32).

The insulin resistance in glycolytic muscle of MCK-DGAT2 mice was also demonstrated by an impaired ability of insulin to stimulate in vivo deoxyglucose uptake in this tissue. This result agrees with the signaling results in that insulin-stimulated PKC{lambda}/{zeta} activity is highly correlated with glucose uptake in muscle (4, 7). Interestingly, basal uptake of deoxyglucose was increased in both glycolytic and oxidative muscle in the MCK-DGAT2 mice. The explanation for this is unclear. Perhaps glucose uptake and metabolism were upregulated to generate fatty acids for esterification by DGAT2. Supporting this possibility, the content of long-chain fatty acyl-CoAs in glycolytic muscle was increased in MCK-DGAT2 mice.

The insulin resistance in glycolytic muscle of MCK-DGAT2 mice was accompanied by impairments in whole body glucose and insulin tolerance. Thus, the effects of DGAT2 overexpression in glycolytic muscle were sufficient to cause derangements in whole body physiology. This was surprising because deoxyglucose uptake was more than 90% lower in glycolytic than in oxidative muscle. However, rodent muscle mass is predominantly (>80%) glycolytic (15), so phenotypic changes in this tissue could clearly be expected to have systemic consequences.

The phenotypic effects we observed were transient and found only in 3-mo-old MCK-DGAT2 mice, when the intramuscular lipid deposition differences between mice of different genotypes were maximal. Effects of the transgene were not found in older mice or mice that had been a fed a high-fat diet. These latter conditions were generally associated with more lipid deposition in glycolytic muscle and, as expected, more glucose intolerance in WT mice. Thus, the effects of age and diet with respect to lipid accumulation in glycolytic muscle appear to trump the effects of the transgene in this model.

In our study, the insulin resistance in glycolytic muscle was associated with increases in the content of TG, ceramides, and unsaturated fatty acyl-CoAs, all lipids that have been implicated in contributing to insulin resistance. Of these, TGs are the least likely candidate since they are stored intracellularly in lipid droplets and are generally thought to be inert. However, much evidence (25, 53, 55) has implicated ceramides and fatty acyl-CoAs in tissue dysfunction and insulin resistance. Ceramide accumulates in skeletal muscle of insulin-resistant obese humans (1), and reduced ceramide levels in skeletal muscle of obese humans or in genetic rodent models are associated with improved insulin action (9, 17). Ceramide levels were highest, and glucose intolerance the most impaired, in mice of either genotype fed a high-fat diet. It is unclear how DGAT2 overexpression caused an increase in ceramide content in glycolytic muscle of MCK-DGAT2 mice. The mRNA expression levels of serine palmitoyltransferase, the rate-limiting enzyme in ceramide biosynthesis, were not increased (data not shown), suggesting an alternative mechanism. One possibility is increased flux through the ceramide biosynthesis pathway, possibly driven by increased fatty acid synthesis. Long-chain fatty acyl-CoAs have also been implicated in insulin resistance in skeletal muscle (25), although little data exist comparing the effects of saturated and unsaturated species. DG has also been implicated in insulin resistance, possibly through its activation of PKC{theta} activity (35, 59). However, the MCK-DGAT2 mice had reduced rather than increased levels of DG in glycolytic muscle, and PKC{theta} activity was not altered in glycolytic muscle of MCK-DGAT2 mice, excluding this as a mechanism for the insulin resistance in this study. Finally, other mechanisms that we did not examine, such as inflammation or mitochondrial dysfunction (8, 52), might also have contributed to the insulin resistance in this model.

The levels of DGAT2 overexpression in MCK-DGAT2 mice appear to be physiologically relevant. Although the level of DGAT2 mRNA overexpression in glycolytic muscle was relatively high (~6-fold increase), the total DGAT2 mRNA levels were similar to those in oxidative muscle of young mice and to those in glycolytic muscle in older or high-fat-fed mice. Also, the increase in TG deposition in glycolytic muscle was 50% lower than the increase in mRNA and was similar to the levels in the oxidative muscle of young WT mice. We generated several lines with higher DGAT2 mRNA levels in skeletal muscle but elected not to study these mice because the higher mRNA expression levels did not further increase muscle TG levels, and the overexpression in these lines was not restricted to skeletal muscle.

Our results contrast interestingly with a recent study by Liu et al. (39) in which transgenic overexpression of DGAT1 in skeletal muscle resulted in increased TG deposition with increased insulin sensitivity. In that study, DGAT1 overexpression was in muscles of both fiber types and was associated with decreases in both DG and ceramide content in muscle. These contrasting results suggest that overexpression of DGAT1 and DGAT2 have dramatically different effects on muscle metabolism. This is not surprising since the enzymes have different characteristics. For example, DGAT1 has broader substrate specificity (57) and an apparently higher Km than DGAT2 (51) and may play a greater role in protecting cells from toxicity of lipids (38). Additionally, increased DGAT1 activity has been implicated in the "exercise paradox" in which increased muscle TG is associated with increased insulin sensitivity (39, 47). DGAT2, on the other hand, may be more important in the esterification of fatty acids of de novo synthesis (42, 51) rather than in protecting cells from lipid-mediated cytotoxicity.

In summary, selective DGAT2-mediated lipid accumulation in glycolytic muscle of mice results in insulin resistance in this tissue and impaired glucose tolerance systemically. To our knowledge, this is the first report to selectively examine the effects of lipid accumulation on insulin signaling in different muscle fiber types. As such, our results highlight the importance of considering glycolytic and oxidative muscle separately when evaluating the effects of lipids on insulin resistance. More attention to this aspect of insulin resistance may yield insights with implications for the pathogenesis and treatment of human insulin resistance and type 2 diabetes.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by fellowships from the Hillblom Foundation and Throne-Holst Foundation (to M. C. Levin), a postdoctoral fellowship from the American Diabetes Association (to M. Monetti), the National Institute of Diabetes and Digestive and Kidney Diseases (5R01-DK-065599 to R. V. Farese, Jr.), an extramural research facilities improvement program project grant (C06 RR018928), a Peter Doherty postdoctoral fellowship from the National Health and Medical Research Council of Australia (M. J. Watt), an Australian Research Council Discovery Project (M. J. Watt), the Department of Veterans Affairs Merit Review Program, and the J. David Gladstone Institutes.


   ACKNOWLEDGMENTS
 
We thank S. Espineda for assistance with generation of transgenic mice, R. Bituin for assistance in mouse husbandry, S. Ordway and G. Howard for editorial assistance, M. Chang and D. Jones for manuscript preparation, and M. Schambelan for comments on the manuscript.


   FOOTNOTES
 

Address for reprint requests and other correspondence: R. V. Farese, Jr., Gladstone Institute of Cardiovascular Disease, 1650 Owens St., San Francisco, CA 94158 (e-mail: bfarese{at}gladstone.ucsf.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* These authors contributed equally. Back


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